Azure BiosystemsHigh-quality and affordable imaging systems, qPCR systems, and reagents.
Azure BiosystemsQuantitative protein identification using Azure Imaging Systems.
Multiplex fluorescent Western blot imaged with an Azure Imager using Cy3 and Cy5
Fluorescent Western blotting has revolutionized the field of protein analysis by providing a highly sensitive and specific method for detecting and quantifying proteins in complex mixtures. Azure Imaging Systems are a state-of-the-art platforms that combines cutting-edge technology with user-friendly software to provide accurate and reliable results in a streamlined workflow. In this protocol, we describe the steps required to perform a fluorescent Western blot using an Azure Imaging System. This protocol is suitable for researchers of all levels and will guide you through each stage of the process, from sample preparation to image analysis. With this protocol, you will be able to confidently generate high-quality data for your research.
Material / Reagent
Recommended
SDS-PAGE Gel Electrophoresis System
Transfer Cell System
Power Supply
SDS-PAGE Gel
User-provided
Sample Protein
User-provided
Protein Dye
User-provided
Molecular Weight Protein Ladder
User-provided
Running Buffer
User-provided
Transfer Buffer
Methanol, 100%
User-provided
Transfer Membrane
Forceps
Plastic (not metal) forceps strongly recommended
Blotting Paper
Rotary or Rocking Platform
Rocking is recommended over orbital shaking
Blocking Buffer
Washing Buffer
Primary Antibody
User-provided
Plastic Folder
Quenching Sheets
Ultimate Western Blot Imager
Fluorescent antibodies are attached to specific proteins on a low-fluorescence polyvinylidene fluoride (PVDF) membrane, allowing for their visualization and quantification through an imager with fluorescent compatibility such as the Azure 600 Imaging System.
Western blotting is a technique in which proteins are first separated by size through polyacrylamide gel electrophoresis (PAGE) and subsequently transferred to a membrane. From there, the membrane would previously be treated with chemiluminescent substrates to identify a protein of interest, but technological advances have allowed for a more advanced methodology in fluorescent labeling.
Fluorescence detection uses the combination of a primary antibody that specifically binds a protein of interest followed by a secondary antibody conjugated to a fluorescent dye molecule designed to bind to the specific species of immunoglobulin that the primary antibody was raised in. Through this combination, the protein of interest is bound to a fluorescent dye molecule so that it can be identified by an imager capable of fluorescent imaging. Therefore, a huge advantage of fluorescent labeling is the ability to identify multiple proteins of interest on the same membrane. A technique termed multiplexing involves using primary antibodies derived from different animal species to probe for multiple proteins of interest simultaneously. Through this, several proteins on the same membrane can be labeled and identified with species-specific, differently-colored secondary antibodies.
Identification and quantification of one or more proteins of interest through antibody-specific fluorescent tagging.
General guidelines are provided as a reference for experiment-dependent protocol optimizations.
1. Sample Preparation
Note: Sample preparation
method will differ
significantly depending
on the experiment.
a. Homogenize cell cultures while keeping cells at ice-cold temperatures to prevent
protein degradation.
b. Add ice-cold lysis buffer containing a protease inhibitor cocktail, as well as phosphatase inhibitor if working with phosphorylated targets to homogenized cells. Centrifuge to separate lysate supernatant from cell debris and collect the lysate supernatant only.
c. Measure protein concentration via a protein assay such as BCA or Bradford.
d. Add sample buffer to the lysate supernatant. Ideally, prepare lysate stocks that are at a final protein concentration of at least 1mg/mL. If reduction of disulfide bonds is desired, include a reducing agent such as DTT, β-mercaptoethanol or TCEP in the sample buffer.
e. Denature the sample by heating at 98°C for 5 minutes.
2. SDS-PAGE Gel Electrophoresis
a. Unpack the pre-cast SDS PAGE gel by unwrapping the gel, removing the tape at the bottom of the gel, and slowly removing the plastic comb from the gel lanes.
b. Place the unwrapped gel with the well’s open side facing inward on the Azure Aqua Quad Mini-Cell’s gel holder. If an odd number of gels are being run, use a plastic barrier to ensure each gel holder can contain liquid. Push the sides of the gel holder inwards to secure the gels and barriers into place.
c. Fill the middle of each gel holder with 1x Tris-Glycine SDS PAGE Running Buffer and check for leaks. If buffer leaks out of the bottom of the gel holder, readjust the sides of the gel holder to ensure that both gels or barriers are sealed securely.
d. Once the middle of each gel holder is filled to the top, fill the body of the Azure Aqua with 1x Running Buffer up to either the “2 Gel” or “4 Gel” lines based on the number of gels being run, rounded up to the nearest multiple of two.
e. Once the gels are secured and the Aqua filled with running buffer, load the denatured/reduced samples and the molecular weight marker into the gel lane.
f. Set up the Azure Aqua Quad Mini-Cell by connecting the red and black power cable from the lid to the Azure Aqua Power Supply. Place the lid on the Quad Mini-Cell by securely attaching the color-coordinated electrodes to the correct colors on the gel holder. Make sure the color of the electrodes on the gel holder matches the labels on the side of the Quad Mini-Cell as well.
g. After connecting the electronics securely, turn on the Azure Aqua Power Supply with the switch on the back-right of the instrument.
h. Once the cable is secured on both ends, press the “Run” option on the Aqua Power Supply to begin the run. When voltage is applied, bubbles should immediately begin to rise in the Running Buffer within the gel holder from the wire at the bottom.
3. Membrane Transfer
Note: The following process will only be detailed for one gel.
a. Gather a blot incubation tray, one transfer membrane, two blotting papers, plastic tweezers, a container large enough to fully submerse the stack in buffer, gel roller, and one transfer cassette from the red-and-black transfer case within the Azure Aqua Transfer Cell.
b. Equilibrate the PVDF membrane in 1X Transfer Buffer. First, place enough 200 proof Methanol in a blot incubation tray to completely submerge the blot.
Gently agitate for 15 seconds to ensure that the membrane is completely wet. Next, decant the methanol and add ~50mL of high purity water to the membrane. Incubate with rocking for 5 minutes. Decant the water and add ~25mL of 1X Transfer Buffer to the membrane. Incubate with rocking for at least 5 minutes.
c. Open the transfer cassette by sliding the white lock along its track and rotating upwards. Separate the two porous sponges within and place them on opposite sides in a large container containing 1X Transfer Buffer.
d. Using the forceps, fully immerse four blotting papers in transfer buffer and place two on each sponge within the transfer cassette.
e. Remove the gel from its casing by cracking open the casing with a designated metal or hard plastic tool.
f. Place the gel centered on the soaked blotting paper that is on the black side of the transfer cassette.
g. Place the PVDF membrane that has been equilibrated in 1X Transfer Buffer onto the gel. Ensure that the gel and membrane are exactly the same size and aligned for the best transfer.
h. Close the transfer cassette so that, from the black side to the clear side, the order should now be: outer sponge, soaked blotting papers, SDS-PAGE gel, equilibrated transfer membrane, soaked blotting papers, outer sponge.
i. Place the transfer cassette back into the red-and-black transfer case in the Azure
Aqua Transfer Cell such that the clear side of the cassette is towards the red side of the case. Additionally, place a frozen ice pack into the Transfer Cell to maintain a low temperature during transfer.
j. Fill the Azure Aqua Transfer Cell with ice-cold Transfer Buffer to the top line labeled “Blotting” and connect it to the Azure Aqua Power Supply.
k. Set the power supply to 55V and run for 35 minutes.
l. After the transfer process is finished, unplug the Aqua Transfer Cell from the power supply and turn the power supply off.
m. Remove the transfer cassette and open it. Using the forceps, carefully remove the transfer membrane and place it in an Incubation Tray.
n. The rest of the contents of the cassette with the exception of the two external sponges can be discarded in the appropriate disposal now. Wash the sponges thoroughly with high purity water then air dry for next usage.
4. Blocking and Staining
a. Transfer the blot to an incubation tray containing ~25mL of high purity water after the transfer is complete and incubate for 5 minutes with rocking.
b. Discard the water and add 10mL of Azure Fluorescent Blot Blocking Buffer.
d. While the blot is being blocked, prepare the primary antibody solution. Transfer 10mL of Fluorescent blocking buffer to a 15mL conical tube, then add primary antibody. Typical primary antibody dilutions range from 1:1000–1:5000.
e. After blocking, discard the Blocking Buffer in the container, while being very careful not to pour the transfer membrane out as well.
g. Wash the blot three times.
h. During the washing step, prepare a 1:50,000 dilution of secondary antibody solution. Transfer 10mL of Fluorescent blocking buffer to a 15mL Falcon tube then add 2μL of 1:10 diluted secondary antibody-HRP conjugate at 1 mg/mL.
i. After the third wash, decant the washing solution and apply the diluted secondary antibody-fluorophore conjugate.
j. Cover and rock under mild agitation for one 30 minutes.
k. Wash the blot three times.
6. Imaging
Note: The Azure 600 Imaging System is referenced in this protocol, but any Azure Imager model that is equal to or greater than the Azure 280 Imaging System will work with this workflow for chemiluminescence.
a. (Optional, but recommended) Prior to imaging, place the treated membrane on a background Quenching Sheet to reduce background autofluorescence.
b. Place the membrane inside the Azure 600 Imaging System on the included Black Chemi Tray.
c. Turn on the Azure 600 by using the power switch on the back-right, then the green power button on the front.
e. Select the “Fluorescent Blot” option and select the fluorescent dyes on the left that
match the Secondary Antibodies used as well as the pseudo-colors they will be
visualized as.
f. Choose between a system-calculated auto exposure with “Auto Image” or input
specific desired settings in “Manually Image.”
g. After the image is taken, it will automatically appear in the Gallery tab. From here, contrast settings can be altered in “Adjustments,” the image can be saved in multiple image formats to the imager’s D drive or an external drive, as well as other options detailed in the Azure Imaging System User Manual.
Images can then be quantitated through Azure Biosystems’ state-of-the-art analysis software AzureSpot Pro or other image analysis programs. Fluorescent quantification excels at identifying sample quantities through relative fluorescent signal intensities. Additionally, multiplexing allows for fluorescent quantifications of different proteins of interest within the same sample simultaneously
AzureSpot Pro combines several powerful analysis tools into one convenient and easy to use package. Designed to guide you through the analysis process, it is an easy-to-use Western blot image analysis software that makes complex and customized analysis simple. Try out AzureSpot Pro free by downloading a free trial.
Fluorescent Western blotting allows for more versatile labeling for proteins that have been transferred to a membrane. Unlike chemiluminescent labeling, fluorescent Western blotting allows for multiplexing several proteins of interests with individual fluorescent dye colors. In order to image each protein, the Azure 600 Imaging System is designed to detect multiple fluorescent dyes in both RGB visible and near-Infrared channels.
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