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Download the Protein Blotting Guide
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In fluorescence, a high-energy photon (ℎVex) excites a fluorophore, causing it to leave the ground state (S0) and enter a higher energy state (S’1). Some of this energy dissipates, allowing the fluorophore to enter a relaxed excited state (S1). A photon of light is emitted (ℎVem), returning the fluorophore to the ground state. The emitted photon is of a lower energy
(longer wavelength) due to the dissipation of energy while in the excited state.
When using fluorescence detection, consider the following optical characteristics of the fluorophores to optimize the signal:
Quantum yield — efficiency of photon emission after absorption of a photon. Processes that return the fluorophore to the ground state but do not result in the emission of a fluorescence photon lower the quantum yield.Fluorop hores with higher quantum yields are generally brighter
Extinction coefficient — measure of how well a fluorophore absorbs light at a specific wavelength. Since absorbance depends on path length and concentration (Beer’s Law), the extinction coefficient is usually expressed in cm–1 M–1. As with quantum yield, fluorophores with higher extinction coefficients are usually brighter
Stokes shift — difference in the maximum excitation and emission wavelengths of a fluorophore. Since some energy is dissipated while the fluorophore is in the excited state, emitted photons are of lower energy (longer wavelength) than the light used for excitation. Larger Stokes shifts minimize overlap between the excitation and emission wavelengths, increasing the detected signal
Excitation and emission spectra — excitation spectra are plots of the fluorescence intensity of a fluorophore over the range of excitation wavelengths; emission spectra show the emission wavelengths of the fluorescing molecule. Choose fluorophores that can be excited by the light source in the imager and that have emission spectra that can be captured by the instrument. When performing multiplex western blots, choose fluorophores with minimally overlapping spectra to avoid channel crosstalk
:: Posted by American Biotechnologist on 01-30-2012
The most common method for analyzing protein expression levels is western blotting with detetion of a single protein target, using horseradish peroxidase-conjugated or alkaline phosphatase-conjugated antibody probes combined with colorimetric or chemiluminescent detection. While these methods work well for studying a single target, they are unsuitable for anlayzing multiple targets at the same time, particularly if the target proteins are of unknown or similar sizes. For analysis of multiple targets, the blot is typically stripped and reprobed for additional targets of interest. Reprobing is time consuming, and often some of the target protein on the blot is lost as a result of the stripping procedure. If one protein is removed to a greater of lesser extent relative to another protein, the ability to quantitate the relative amounts of diffferent proteins of interest is compromised.
In this technical note, you will be introduced to fluorescent western blotting detection which is superior to traditional western blotting when trying to analyze multiple proteins.
fast and quantitative detection of multiple proteins in a single experiment
sensitivity compared to chemiluminescent detection
linear dynamic range up to 10 times greater than that of chemiluminescent detection
fewer experimental steps than chemiluminescent detection
no substrate requirement, and therefore no risk of exhausting the substrate and causing a “dead zone” in the blot
the ability to visualize and quantitate both phosphorylated and non-phosphorylated forms of individual proteins
The technical note is divided into three sections to help those who are new to fluorescent western blot detection quickly generate reliable and reproducible results.
:: Posted by American Biotechnologist on 01-19-2012
So you’ve isolated your protein, ran them on a gel and now you’re ready to transfer them to a membrane to begin western blotting. Sounds simple, right? Not so fast. Don’t forget to equilibrate your gel prior to beginning your transfer. Gel equilibration generally involves rinsing the gel in diH2O and soaking it in transfer buffer for approximately 15 min. While it may sound simple, (and it truly is), it is a step that might make the difference between an ugly blot and one that is publication worthy.
Below are some points to consider about gel equilibration:
Gel equilibration removes contaminating electrophoresis buffer salts. If not removed, these salts increase the conductivity of the transfer buffer and the amount of heat generated during transfer.
Equilibration also allows the gel to adjust to its final size prior to electrophoretic transfer. Gels shrink or swell to various degrees in the transfer buffer depending on the acrylamide percentage and the buffer composition.
Equilibration is not necessary (i) when the same buffer is used for both electrophoresis and transfer (for example, native gel transfers), or (ii) when using rapid semi-dry transfer systems such as the Trans-Blot® Turbo™ system (consult the user manual for the system you are using).
:: Posted by American Biotechnologist on 01-12-2012
So you think that protein transfer for western blotting is a piece of cake? Consider these important tips before proceeding:
Use only high-quality, analytical grade methanol. Impure methanol can increase transfer buffer conductivity and yield a poor transfer.
In many cases, ethanol can be substituted for methanol in the transfer buffer with minimal impact on transfer efficiency. Check this using your samples.
Do not reuse transfer buffer since the buffer will likely lose its ability to maintain a stable pH during transfer.
Do not dilute transfer buffers below their recommended levels since this decreases their buffering capacity.
Do not adjust the pH of transfer buffers unless specifically indicated. Adjusting the pH of transfer buffers can result in increased buffer conductivity, manifested by higher initial current output and decreased resistance.
Increasing SDS in the transfer buffer increases protein transfer from the gel but decreases binding of the protein to nitrocellulose membrane. PVDF membrane can be substituted for nitrocellulose when SDS is used in the transfer buffer.
Addition of SDS increases the relative current, power, and heating during transfer, and may also affect antigenicity of some proteins.
Increasing methanol in the transfer buffer decreases protein transfer from the gel and increases binding of the protein to nitrocellulose membrane.
:: Posted by American Biotechnologist on 12-21-2011
Despite the difficulties associated with measuring in-situ protein-protein interactions in neural networks, Dr. Akira Chiba of the University of Miami recently announced that his team has embarked on a project to develop a protein interaction map within brain cells. Why have these studies been so difficult to perform until now and what does Dr. Chiba have that will make him successful? The answer lies in the small size of neural proteins and the technical limitations associated with even the highest resolution microscope.
Using a a custom- built 3D FLIM (fluorescent lifetime imaging microscopy), Chiba’s team has been able to spatially and temporally quantify fluorescently tagged protein-protein interactions in genetically modified fruit flies.
According to Chiba, “collaborating fluorescent chemistry, laser optics and artificial intelligence, my team is working in the ‘jungle’ of the molecules of life within the living cells. This is a new kind of ecology played out at the scale of nanometers—creating a sense of deja vu 80 years after the birth of modern ecology.”