ELISA Introduction

Enzyme-linked immunosorbent assay (ELISA) is widely used in immunology to detect the presence of proteins or other antibodies in a sample. For instance, it is used as an initial detection tool for HIV, based on the interaction of an antibody with antigen presented by the virus. There are several different ELISA methods: indirect ELISA, sandwich ELISA and competitive ELISA are the most commonly applied. All three methods have similar steps:

1) Attach antigens or primary antibodies or their complexes to a solid surface.

2) Wash away unbound substances.

3) Block exposed sites on the solid surface.

4) Add detection antibodies and/or enzyme-conjugated secondary antibodies.

5) Develop color by adding substrates that react with the enzymes.

For a detailed step-by-step guide to Indirect ELISA download the pdf protocol below.

Immunohistochemistry Introduction

Immunohistochemistry (IHC) allows you to visualize proteins in tissue while remaining its microstructure. It helps to demonstrate the exact position and distribution of the protein of interest in the analyzed tissue section. The advantage of this visualization is that it allows for comparison between, for example, healthy and diseased tissues. Briefly, in an IHC experiment, the antigen of interest is localized by the binding of an antibody. The antibody-antigen interaction is then further visualized via chromogenic or fluorescent detection.

The IHC protocol contains many steps that may require optimization to ensure specific antibody binding and optimal visualization of the target protein. As the IHC protocol contains many different variable factors, it can be challenging to find the best working conditions to obtain strong and specific staining. 

Step-by-step IHC protocol

Immunohistochemistry using slide-mounted paraffin sections

All steps in the following protocol are carried out at room temperature unless stated otherwise.  Recipes for all solutions highlighted  bold  are included at the end of the protocol.

1.  Deparaffinizing and rehydration:

• Immerse slides in xylene for 10 minutes. Repeat this step again in fresh xylene  for 10 minutes. (If required, repeat a third time in fresh xylene for another 10 minutes.)

• Rehydrate sections by sequentially incubating with 100%, 95%, 80% and 60%  ethanol for 5 minutes each.

• Rinse sections with distilled water three times for 3 minutes each.

 2.  Antigen retrieval (optional):

• Transfer slides to a microwave-safe container and cover with  Citrate buffer  or Tris-EDTA (TE) buffer.

• Heat in the microwave on medium power for 10 minutes.

• Allow slides to cool in the Citrate buffer or Tris-EDTA (TE) buffer  for approximately  35 minutes.

3. Proteintech antibody incubation:

• Rinse slides three times with 1x TBST  for 3 minutes each.

• Incubate slides with 3% H2O2  solution (diluted in distilled water) for 10 minutes  to quench endogenous peroxidase activity.

• Rinse slides three times with 1x TBST  for 3 minutes each, then rinse slides three  times with distilled water for 3 minutes each.

• Prepare 5% normal blocking serum in 1x TBST. The serum should be derived from  the same species in which the secondary antibody was raised. Block the sections  for 1 hour. (Alternatively, use 5% BSA in 1x TBST  for blocking if the corresponding  serum is not available.)

• Incubate sections with primary antibody diluted in 1x TBST  for 1 hour, or  overnight at 4°C; the optimal antibody dilution ratio should be pre-determined  by experimentation. Set up negative controls by omitting the primary antibody  incubation step for one slide per each experimental condition.

• Following primary antibody incubation rinse slides three times with 1x TBST  for  3 minutes each.

4. Signal detection

• Apply sufficient peroxidase labeled polymer and incubate for 30 minutes.

• Rinse slides three times with 1x TBST for 3 minutes each.

• Prepare an appropriate volume of substrate solution prior to use by mixing  one drop of Liquid DAB plus chromogen immediately with 1 ml of substrate  buffer.

• Apply the substrate carefully and incubate for 5–10 minutes till a brown  color develops.

• Rinse sections gently with sufficient distilled water.

5.  Hematoxylin counterstaining (optional):

• To stain nuclei, immerse slides in a bath of hematoxylin for 3 minutes.

• Rinse slides gently with distilled water.

• Transfer slides into a 1% HCl, 99% ethanol solution for 10 seconds; transfer  to distilled water immediately

6. Dehydration and mounting:

• Immerse slides sequentially into 60%, 80%, 95% and 100% ethanol baths  for 5 minutes each.

• Immerse slides in xylene for 5 minutes. Repeat this step again in fresh xylene  for 5 minutes.

• Mount the section with sufficient mounting media and cover with a cover slip.

• Air-dry in a well-ventilated area (e.g. fume hood).

Solutions

For more IHC technical support contact us via LiveChat.

Immunofluorescence Introduction

Immunofluorescence (IF) staining is a widely used technique in biological research and clinical diagnostics. IF utilizes fluorescent-labeled antibodies to detect specific target antigens. Followed by imaging, it is a very direct technique as you can visualize results. Although it is a well-established tool, multiple factors have to be considered and various optimization steps have to be taken to ensure successful staining.

Step by step IF protocol

Immunostaining Cultured cells

All steps in this protocol are performed at room temperature unless otherwise indicated. For optimum staining, incubations should be carried out on a slow-moving rotary shaker unless the cell line being  used is delicate (e.g. neuronal cells).  Recipes for solutions highlighted bold are provided following the protocol.

1. Fixation and permeabilization:

• Aspirate medium, wash cells seeded on clean glass cover slips briefly with  1X PBS.

• Fix the cells with 4% paraformaldehyde made fresh in 1X PBS  for 10 minutes.

• Rinse cover slips with 1X PBS  3 times for 3 minutes each.

• Permeabilize with 0.2% Triton X-100 made in 1X PBS for 5 minutes. Rinse cover slips 3 times with 1X PBS for 3 minutes each.

2.  Blocking:

• Prepare a blocking solution of 5% normal serum in 1X PBS.  Select serum from the same species in which the secondary antibody was raised e.g. if the secondary antibody is goat anti-mouse, then goat serum should be selected for the blocking  solution. Incubate the cells with the blocking solution for 1 hour (Alternatively, use 1% BSA in 1X PBS for blocking if the corresponding serum is not available.)

3.  Antibody incubations:

• Aspirate the blocking solution and apply primary antibody diluted in antibody dilution buffer. Set aside one cover slipper experimental condition for a negative control and incubate in antibody dilution buffer minus the primary antibody.  Leave these incubations for 1 hour, or, alternatively, incubate overnight at 4°C.

  Please Note: If incubating overnight, take measures to ensure the cover slips do not dry out.

• Wash cover slips with 1X PBS 3 times for 3 minutes each.

• Apply an appropriate fluorophore-conjugated secondary antibody diluted in  antibody dilution buffer  to the coverslips and incubate for 1 hour in a moist,  dark environment.

  Please Note: It is imperative that cover slips be kept in dark conditions as much  as possible after the addition of fluorescent reagents.

• Wash cover slips with 1X PBS 3 times for 3 minutes each.

4.  Mounting and visualization:

• Mount cover slips on microscope slides with Hydromount (National Diagnostics) containing DAPI (if desired) for nuclear staining.

• Examine slides under a fluorescence microscope.

Solutions

For more IF technical support download the step-by-step protocol guides below or contact us via LiveChat.

Immunoprecipitation Introduction

Immunoprecipitation (IP) is an affinity purification technique. IP purifies an antigen using a specific antibody that is immobilized to a solid matrix. IP is one of the most commonly used methods for isolation of proteins out of cell or tissue lysates. 

Step-by-step IP protocol

Keep samples as cool as possible by carrying out the steps below on ice or in a 4°C cold room.  Recipes for all solutions highlighted  bold  are included at the end of the protocol.

Sample preparation

• Cells may be lysed using any standard cell lysis protocol compatible with your  starting material. See “Cell and Tissue Lysate Preparation” for Proteintech recommended lysis protocols.

For more IP technical support, contact us via LiveChat.

Western Blot Introduction

Western Blotting is an analytical Immunoblotting Technique to detect specific proteins in a cell extract or tissue homogenate. Western Blotting relies on the specific binding between the protein-of-interest and an antibody raised against this particular protein.

Step-by-step western blot protocol

All steps are carried out at room temperature unless otherwise indicated. Recipes for all solutions highlighted bold are included at the end of the protocol.

SDS PAGE

• Construct an SDS-PAGE gel according to the molecular weight (MW) of your  target protein(s). (Recommendations and gel recipes are presented at the end of  this protocol)

• Tip 1: Tris-tricine gels separate low MW proteins (<20 kDa) better than Tris-glycine gels.

  Blog: Tech Tips | In search of low molecular weight proteins

  Tip 2: Gradient polyacrylamide gels can produce sharper bands and they separate a broader range of MW sizes on one gel, such as 10–500 kDa.

• Prepare samples in microfuge tubes. Add   4X SDS sample buffer  so the total  protein amount is 30–50 μg per sample (according to the protein amount  measured by Bradford or BCA protein assay).

• Flick microfuge tubes to mix samples, and then heat to 95-100°C for 5 minutes.

• Set up electrophoresis apparatus and immerse in  1x running buffer. Remove  gel combs and cleanse wells of any residual stacking gel.

• Load samples and protein markers onto the gel using gel loading tips. Set electrophoresis power pack to 80V (through the stacking gel), before  increasing it to 120V when the protein front reaches the separation gel.

  Tip 3: Load generous volumes of sample for the first experiment and adjust as necessary  after assessing the initial target signal.

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Protein transfer

• PVDF membranes (or PSQ membranes with 0.22 μm micropores when MW  of target is <30 kDa) are strongly recommended. Soak membranes in methanol  for 30 seconds before moving to  transfer buffer. Soak the filter papers and  sponges in  transfer buffer  as well.

• Sequentially assemble the transfer constituents according to the illustration  on page 7 of this booklet and  ensure no bubbles lie between any of the layers. Apply semi-dry  or wet transfer systems according to the manufacturer’s instructions.

  Tip 4: If target MW is larger than 100 kDa, wet transfer at 4°C overnight is suggested  in place of a semi-dry method; moreover, we recommend adding 0.1% SDS to the wet transfer buffer to facilitate transfer.

  Tip 5: Gradient polyacrylamide gels can provide sharper bands and they separate a broader  range of MW sizes on one gel, such as 10–500 kDa.

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Immunoblotting

• After transfer, wash the membrane twice with distilled water, and using a pencil, mark bands of the MW ladder on the membrane. If desired, stain the membrane with commercial Ponceau red solution for 1 min to visualize protein bands, then  wash any Ponceau red staining with copious amounts of  1x TBST.

• Block with 1x TBST containing (2-5%) nonfat dry milk (or 1-5% BSA for the  detection of phospho-epitope antibodies) with constant rocking for 1 hour or overnight at 4°C.

• Dilute primary antibody in blocking solution with a starting dilution  ratio of 1:1000. (Optimal dilutions should be determined experimentally.) Incubate the membrane with primary antibody for 1 hour at room temperature,  or overnight at 4°C.

• Wash membrane three times with 1x TBST for 10 minutes each.

• Incubate the membrane with a suitable HRP-conjugated secondary antibody (recognizing the host species of the primary antibody), diluted at 1:5000–1:50000 in blocking solution. Incubate for 1 hour with constant rocking.

• Wash membrane three times with 1x TBST for 10 minutes each.

  Tip 6: Do not let the membrane dry at any stage of the blotting process.

  Tip 7: For preservation of the primary antibody solution over long incubations, 0.02% NaN₃ could be included in the antibody dilution buffer. Not suitable for use  with secondary antibody solutions.

Signal detection

• Prepare ECL substrate according to the manufacturer’s instructions.

• Incubate the membrane completely with substrate for 1–5 minutes (adjust time for more sensitive ECL substrates e.g. SuperSignal West Femto Chemiluminescent Substrate [Pierce]).

• Expose the membrane to autoradiography film in a dark room or read using a  chemiluminescence imaging system.

  Tip 8: Use multiple exposure lengths to determine the optimal exposure time.  Use fluorescent markers as a guide for blot-film orientation.

Line up the developed film in the correct orientation to the blot and mark the  bands of the MW ladder directly onto the film. It is also advised to add notes  such as lane content, film exposure time and ECL properties. 

Solutions

SDS PAGE Gel recipes 

For target proteins with MWs between 20 and 200 kDa, make a conventional SDS-PAGE gel using the following recipes in the table below. Select the percentage of gel you require using the MW of your target protein.

Tricine Gel Recipe

For target protein with MWs of less than 20 kDa, a tricine gel system will obtain  higher resolution and is highly recommended. Make three layers of tricine gels as  laid out in the following table and diagram. Apply specific tricine gel running buffer to the running system and perform transfer as usual.

Introduction to Affinity-Purification

Antibody purification is performed using antigen-coupled sepharose, meaning only those antibodies specific to the immunogen are purified from the serum.

Step-by-step affinity-purification protocol

Affinity purification of antibodies via protein coupled sepharose

Preparation of affinity matrix and coupling of protein to the Sepharose matrix:

• Dialyze 1 mg of fusion protein in coupling buffer overnight at 4°C.

• Calculate the amount of cyanogen bromide (CNBr)-activated Sepharose4B needed  for protein coupling. Usually, 1 g of CNBr activates 3.5 ml of Sepharose beads and 1 ml of activated Sepharose beads may absorb 5–10 mg of protein.

• Activate the Sepharose beads in 20–50 ml cold 1 mM HCl for 15 min at 4°C.

• Wash the beads with 1 mM HCl. In general, 1 g of Sepharose beads requires 200 ml  of HCl to wash.

• Incubate appropriate amounts of activated Sepharose beads with dialyzed fusion proteins for 2 hrs at room temperature or overnight at 4°C.

• Wash protein-coupled Sepharose matrix with 15ml  of coupling buffer.

• Add 5 ml 0.1 M Tris-HCl buffer (pH 8.0) or 1 M Ethanolaniba to block the  uncoupled sites on beads and let stand for 2 h at room temperature or overnight at 4°C.

Purification of antiserum via protein-coupled Sepharose:

• Wash the beads at least three cycles with acid and alkali buffer alternative. (0.1 M Acetic/Sodium Acetate, 0.5 M NaCl, pH4.0; 0.1 M Tris-HCl, 0.5 NaCl, pH 8.0).

• Incubate the beads with serum for 1–2 h at room temperature or overnight at  4°C.

• Collect the flow-through from the purification column and save it for ELISA testing.

• Wash the beads 3 times with 10 ml PBS buffer.

• Wash the column with 10 ml 150 mM NaCl-HCl (pH 5) solution.

• Elute the antibodies with 6 ml elution buffer and neutralize the solution with  saturated phosphate buffer. Usually, 1 ml of elution buffer requires 50–100 μl  of saturated phosphate buffer depending on the temperature.

• For short-term storage, keep the antibody solution at 4°C; for long-term storage,  keep the antibody in a 50% glycerol solution with 0.02% sodium azide at -20°C

Beads washing and recycling:

• Wash beads 3 times with 15 ml 0.01 M Tris-HCl (pH 7.5) buffer.

• Wash beads 3 times with 10 ml of PBS buffer.

• Add 2 ml PBS, 3 ml Glycerol with 0.02% sodium azide to the beads and store at -20°C for future use.

Buffers needed:

For more technical support regarding antibody purification download the step-by-step protocol guides below or contact us via LiveChat.

Introduction to Flow Cytometry

Flow Cytometry is a technology that measures and analyzes several physical characteristics of single cells. The cells flow in a fluid stream through a beam of laser light. Using Flow Cytometry the following cell characteristics can be determined: cell size, cell granularity, complexity and relative fluorescence intensity.

Step-by-step flow cytometry protocol

Flow Cytometry Intracellular and Membrane Staining Protocol

Cell fixation (for membrane protein):

• Suspend cells in 1x PBS buffer and wash them twice with 1x PBS buffer by centrifugation at 350-500 x g for 5 min each time. Discard the supernatant.

• Re-suspend the cells in 1 ml of 1x PBS buffer briefly.

• Fix the cells in a final concentration of 4% formaldehyde (or paraformaldehyde)  for 20 min at room temperature.

• Wash the cells 3 times with 1x PBS buffer by centrifugation at 350-500 x g for 5 min  each time.

Cell fixation and Permeabilization (for intracellular protein):

• Permeabilize cells by adding 100% cold methanol slowly to pre-chilled  cells to a final concentration of 90% methanol before incubating for 30 min on ice.

• Alternatively, fix the cells in a final concentration of 4% formaldehyde  (or paraformaldehyde) for 20 min at room temperature. Then incubate the cells  in 0.1% Triton X-100 in 1x PBS buffer for 15 min at room temperature.

• Wash the cells 3 times with 1x PBS buffer by centrifugation at 350-500 x g for 5 min each time.

Immunostaining:

• Blocking: Incubate the cells with 3 ml blocking buffer for 1h at room temperature.

• Add primary antibody at an appropriate dilution and incubate for 15-45 min at  room temperature.

• Wash the cells 3 times with 1x PBS buffer by centrifugation at 350-500 x g for 5  min each time.

• Add diluted secondary antibody (enzyme or fluorescein conjugated or other types)  to the cells and incubate for 45 min at room temperature.

• Wash the cells 3 times with 1x PBS buffer by centrifugation at 350-500 x g for 5  min each time.

• Re-suspend the cells in 0.5 ml 1x PBS buffer and analyze the results on a flow cytometer. For DNA staining, re-suspend the cells in 0.5 ml of DNA dye instead;  incubate for at least 5 min at room temperature before analyzing the results on  a flow cytometer.

Buffers needed:

Introduction to Lysate Preparation

Cell lysis is the breaking down of the cell membrane and the separation of proteins from the non-soluble parts of the cell. Lysate buffers contain different detergents that help to release soluble proteins (Triton-X, Tween, SDS, CHAPS). Dependent on the location of the protein of interest, a different lysate buffer is needed to obtain a high yield and purity of the protein.

Cell and tissue lysate preparation

Recipes for all solutions highlighted  bold  are included at the end of the protocol.

Cultured cells:

Pre-cool a refrigerated centrifuge to 4°C. Pellet the cultured cells by centrifugation for 5 minutes at 1000 x g (approximately 2000 rpm) at 4°C. Wash 3 times with ice-cold 1X PBS  and then add chilled RIPA buffer  with  protease inhibitor. In general, add 100μl RIPA buffer for approximately every 10⁶   cells present in the pellet (count cells before centrifugation). Reduce the  volume of  RIPA buffer  accordingly if a higher protein concentration is required. Vortex to mix and keep on ice for 30 min, vortexing occasionally.

Tissues:

Dissect the tissue of interest and wash briefly with chilled 1X PBS  to remove  any blood if necessary, cut the tissue into smaller pieces whilst keeping it on ice. Transfer the tissue to a homogenizer and add RIPA buffer with protease inhibitor. In general, add 500μl RIPA buffer for approximately every 10 mg  of tissue. Homogenize thoroughly and keep the sample on ice for 30 min. Vortex occasionally.

Lysis and storage:

Sonicate the sample to break the cells or tissue up further and to shear DNA. Adjust sonication time to your type of sample: 1 min for cell lysates  and 2–5 min for tissue lysates at a power of about 180 watts (in rounds of 10 seconds sonication/10 seconds rest for each cycle). Keep the sample on ice  during the sonication.

Tip 2:   The addition of DNase for DNA digestion is not recommended as this introduces protein contamination from the enzyme.

• Centrifuge at 10,000 x g (approximately 9700 rpm for rotors of a 9.5 cm radius)  for 20 minutes at 4°C to pellet cell debris, and then transfer the supernatant to  a fresh microfuge tube without disturbing the pellet.

• Determine protein concentration of the lysate by Bradford or BCA protein assay.

• Samples can be frozen at -80°C for long-term storage, or be used for immediate Western blotting or immunoprecipitation.

• For Western blotting, mix sample with 4X SDS sample buffer to a final dilution of 1X. Heat the mixture to 95°C for 5 minutes before loading onto an SDS-PAGE gel.

Solutions:

Introduction

Inclusion bodies are aggregates of proteins. They can be found in the cell cytoplasm or nucleus. Inclusion bodies can be recovered from the cell lysates by centrifugation, extraction, and washing. 

Step-by-step protocol

Purification of proteins from inclusion bodies

Recipes for all solutions highlighted bold are included at the end of the protocol.

a. Suspend the cell pellet (from 1 L culture) in 30–35 ml of PBST buffer.

b. Sonicate cells in an ice-bath at 200 W for 6 min.

c. Centrifuge cell lysate for approximately 13 min at 8000 rpm, 4°C. Discard the supernatant.

d. Re-suspend the pellets in 5 ml TNMFX-2M Urea buffer before transferring to a 10 ml centrifuge tube.

e. Sonicate the solution in an ice-bath at 200 W for 1 min.

f. Add an additional 5 ml TNMFX-2M Urea to the tube. Rotate for 30 min at 4°C.

g. Centrifuge for 20 min at 4000 rpm, 4°C. Discard the supernatant.

h. Repeat steps d-g.

i. Re-suspend the pellets in 5 ml of TNMFX-0.1% Triton-X100.

j. Sonicate the solution in an ice-bath at 200 W for 1 min.

k. Add an additional 5 ml TNMFX-0.1% Triton-X100 to the tube. Rotate for 30 min at 4°C.

l. Centrifuge for 20 min at 4000 rpm, 4°C. Discard the supernatant.

m. Repeat steps i-l.

n. Vortex and wash the pellets with 2x volumes of dH₂O. Centrifuge at 1000 rpm for 2 min.

o. Repeat washing until the supernatant becomes clear. Collect the pellets.

p. Dissolve the proteins depending on intended application:

- For immunization, dissolve in 1.5x volumes of 8 M urea (pH 8).

-  For antibody purification, incubate in 2x volumes of PBS with 2% Sarkosyl overnight at 4°C. Collect the supernatant by centrifugation at 1000 rpm for 7 min.

Solutions

FlexAble Antibody Labeling Kit Introduction

Flexable is anovel antibody labeling kit that uses an affinity linker to conjugate fluorchromes, enzymes, and molecules in any buffer condition.

What's in the FlexAble Antibody Labeling Kit box?

• FlexLinker

• FlexQuencher

• FlexBuffer

2-Step-Protocol

Standard Workflow

Before you begin, equilibrate all reagents to room temperature.

Procedure:  

1. Combine 0.5µg of primary antibody with 1µL of FlexLinker. Add FlexBuffer to bring total volume to 8µL.

2. Mix gently and incubate for 5 minutes at room temperature in the dark.

3. Add 2µL of FlexQuencher.

4. Mix gently and incubate for 5 minutes at room temperature in the dark.

5. The antibody is now ready to be used.

Scale up proportionately to label larger quantities of primary antibody 

Multiplexing

Step 1-4 see "Standard Workflow"

• Primary antibodies of different species can be combined in the same tube and and labeled simultaneously using corresponding species-specific FlexLinkers. Scale up the reagent volumes appropriately based on the amount of primary antibody to be labeled.

• For primary antibodies of the same species, label each antibody individually. Once the primary antibodies are labeled, combine and use in your desired application.

• Antibodies labeled with the FlexAble kit can be used  alongside pre-conjugated primary antibodies.

For more FlexAble Antibody Labeling Kit technical support contact us via LiveChat.