This work was supported by grants from the National Institutes of Health and the George E. Hewitt Foundation fellowship (A.P.).

Gel electrophoresis
Technical note: Before starting the procedure, have your protein samples ready.
During this first step, the proteins in the sample are separated according to their molecular weight using denaturing polyacrylamide gel electrophoresis (PAGE). The NuPAGE&reg; LDS Sample Buffer loaded with Lithium Dodecyl Sulfate (LDS) maintains polypeptides in a denatured state once the protein sample has been heated at 70&deg;C for 10 minutes. A strong reducing agent is used in conjunction to remove secondary and tertiary structure (DTT, to break disulfide bonds). In addition, sample proteins become covered in the negatively charged LDS and therefore move through the acrylamide mesh of the gel toward the positively charged electrode. This allows their separation according to molecular weight (measured in kilo Daltons, kDa).
Detailed step-by-step protocols for the sample preparation and the PAGE procedure can be found on the Invitrogen website2, or in the NuPAGE technical guide3.
Technical tips
<ol>
<li>In the Invitrogen NuPAGE Bis-Tris discontinous buffer system, the electrophoretic mobility of the proteins and the subsequent separation range of the gel is dependent on two factors: i) the acrylamid concentration of the gel (with greater acrylamide concentration resulting in better resolution of lower molecular weight proteins) and ii) the trailing ion of the running buffer, MOPS or MES. To choose the appropriate combination for the separation range that you wish to achieve, refer to the Gel Migration chart1. The thickness of the gel and number of wells will depend on the volume and the quantity of samples that you plan to load, respectively.</li>
<li>When you load your samples into wells of the gel, at least one lane is reserved for a molecular weight marker (or ladder). These protein standards are commercially available from several companies and typically consist of a mixture of stained proteins having defined molecular weights, so as to form visible bands that allow you to follow the migration progress. Pick a range of molecular weights that is compatible with your gel resolution.</li>
<li>To achieve a nice and even migration pattern we recommend loading ALL the wells of your gel with a similar volume of sample or 1X LDS sample buffer.</li>
</ol>
Transfer
Technical Note: Before beginning the transfer step, prepare the transfer buffer (1X with 10% methanol) and pre-cool to 4&deg;C in a cold room.
In order to make the proteins accessible to antibody detection, they are transfered by electroblotting from the gel onto a nitrocellulose membrane. Protein binding is based upon hydrophobic interactions, as well as charge interactions between the membrane and protein.
(Polyvinylidene fluoride (PVDF) membrane can also be used as an alternative. In this case, the PVDF membrane needs to be pre-wet in methanol at least 30 seconds before use.)
A detailed step-by-step protocol of the transfer procedure can be found in reference 4 if you use the Bio-Rad Mini Trans-Blot Electrophoretic Transfer Cell, or in references 2-3 if your lab is equipped with the Invitrogen&rsquo;s XCell II&trade; Blot Module.
As a result of this process, the proteins are exposed on a thin surface layer and ready for detection. The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by the reversible Ponceau S dye membrane staining.
Ponceau S staining procedure (optional):
<ol>
<li>After transferring the proteins, place the membrane in an incubation tray (proteins facing up).</li>
<li>Add enough Ponceau S Staining to cover the membrane and incubate at least 30 seconds with gentle agitation.</li>
<li>Rinse membrane with distilled water until the background is clear.</li>
<li>Destain the membrane with running distilled water for 2-3 minutes.</li>
<li>Place the membrane in blocking solution (see below).</li>
</ol>
Technical tips:
<ol>
<li>We recommend labeling your membrane with a pencil before protein transfer in order to denote the side in which the proteins were exposed and the orientation of your samples.</li>
<li>Both Ponceau S staining and the molecular weight markers may be used to cut the membrane after transfer for probing the resulting parts with different antibodies </li>
</ol>
Immunodetection:
Technical Note: This immunodetection procedure is provided as a guideline only. Optimization may be required for each antibody; specific information can be found in most data-sheets of the commercially available antibodies (e.g., working dilution, incubation time, etc.).
During this last process, the target protein will be detected using a specific antibody and will appear as a band on the film. The position of the band is dependent of the molecular weight of the target protein, whereas the band intensity depends on the amount of target protein present.
Typically, this is achieved after three substeps:
<ol>
<li>Blocking: To avoid non-specific interactions of the antibody with the membrane (the excess space on the membrane is covered with a dilute solution of a generic protein).</li>
<li>Probing: The protein of interest is detected by a specific <a title="Primary antibody" href="http://en.wikipedia.org/wiki/Primary_antibody">primary antibody</a>. After the unbound primary antibody is washed away, the membrane is exposed to a different antibody linked to the reporter <a title="Enzyme" href="http://en.wikipedia.org/wiki/Enzyme">enzyme</a>, <a title="Horseradish peroxidase" href="http://en.wikipedia.org/wiki/Horseradish_peroxidase">horseradish peroxidase</a> (HRP). This secondary antibody is directed against the species-specific portion of the primary antibody (e.g. an anti-rabbit secondary antibody will bind to any rabbit-sourced primary antibody). </li>
<li>Detection: A chemiluminescent agent is used as a substrate that will luminesce when exposed to the HRP on the secondary antibody. This reaction produces luminescence in place and in proportion to the amount of probed protein. The light is then detected by photographic film.</li>
</ol>
A generic step-by-step procedure is provided below. Other detailed procedures can be found in the ECL Plus Western Blotting Detection Reagents instruction manual5 or in most data-sheet accompanying the commercially available antibodies. Incubation time, antibody dilution, and blocking and wash solutions have to be empirically optimized for each antibody.
Immunodetection procedure:
<ol>
<li>Block the non-specific binding by incubating with enough volume of PBS 1% casein blocking solution to cover the entire membrane. Place on a rocker for at least 30 min at room temperature or overnight at 4&deg;C. </li>
<li>Incubate the membrane with your primary antibody (specific for the protein of interest). The primary antibody should be diluted in blocking solution. For most antibodies, complete binding is achieved after 1-2 hours incubation at room temperature under gentle agitation. Alternatively, the incubation step can be performed overnight at 4&deg;C to increase the signal to noise ratio. </li>
<li>Remove the solution (discard or keep at 4&deg;C to -20&deg;C for repeated use) and quickly wash the membrane once. Then 3X10 minutes with PBS, 0.05% Tween 20 (PBST) at room temperature with vigorous shaking.</li>
<li>Incubate the membrane with a HRP-coupled secondary antibody diluted in blocking solution. Choose an antibody which recognizes the IgG portion of the species where the primary antibody was raised. The incubation is performed for 1 hour at room temperature under gentle agitation.</li>
<li>Discard the solution and quickly wash the membrane once. Then 3X10 minutes with PBST at room temperature with vigorous shaking.</li>
<li>Proceed to the chemiluminescent detection according to the manufacturer instruction. We use the ECL Plus Western Blotting Detection Reagents from GE Healthcare (see reference 5 for specific instructions).</li>
<li>Place your membrane in saran wrap or a pouch and place inside an autoradiography cassette. Make sure to drain any excess liquid and remove all air bubbles.</li>
<li>Expose the photographic film to membrane in a dark room. Start with a 1 minute exposure time and adjust according to desired signal intensity.</li>
</ol>
Technical tips:
<ol>
<li>Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein. We typically use 1% casein, but <a title="Bovine serum albumin" href="http://en.wikipedia.org/wiki/Bovine_serum_albumin">bovine serum albumin</a> (~2% BSA) or non-fat dry milk (~5%) can be used as an alternative.</li>
<li>Tris-Buffered Saline (TBS) can be used throughout the procedure instead of PBS. We recommend using TBS if you plan to probe your membrane with phosphospecific antibody.</li>
<li>The use of a glow-in-the-dark sticker (e.g., Stratagene&rsquo;s Glogos&reg; II Autorad Markers) tapped in the bottom of the autoradiography cassette will help you to align your film and your membrane during the analysis step.</li>
</ol>
Analysis
Now that you have exposed your film, you will realize that in practical terms, not all Westerns reveal protein as one nice single band. Additional bands may also appear due to the non-specific binding of both primary and secondary antibodies. This background signal can be reduced by optimizing the immunodetection procedure. In addition, an appropriate control (e.g., untransfected cells, siRNA-treated cells, etc ) will be useful to determine the specificity of your antibody and the exact location of the target protein on the membrane.
After marking on the film the position of the stained protein standard bands from the membrane, plot the log of each molecular weight of the protein standards (y-axis) against their corresponding relative mobility (x-axis). Relative mobility (Rf) is the term used for the ratio of the distance the protein has moved from its point of origin (top of the gel) relative to the distance the tracking dye or a low molecular weight marker has moved (the gel front). Determine the regression line of the standard curve to obtain values for slope and y-intercept. The unknown molecular weight (size) of your target protein is estimated using its Rf and the following modified equation:
log molecular weight = (slope)(mobility or Rf of the target protein) + y-intercept
(see reference 6 for detailed instructions).
Expression level approximations are taken by comparing the band intensity of the target protein to that of a structural protein (e.g., tubulin or actin) or a housekeeping gene product such as GAPDH. This so called "loading control" should not change between samples and is revealed using a specific primary antibody. The image may be further analyzed by densitometry to evaluate the relative amount of protein staining and quantify the results in terms of optical density.

<ol>
	<li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gel+Migration+chart:+.">Gel Migration chart: .</a> , .</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complete+western-blot+protocol+from+the+Invitrogen+website.">Complete western-blot protocol from the Invitrogen website.</a> , .</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NuPAGE+technical+guide.">NuPAGE technical guide.</a> , .</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Instruction+Manual,+Mini+Trans-Blot+Cell.">Instruction Manual, Mini Trans-Blot Cell.</a> , .</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ECL+Plus+Western+blotting+detection+reagents+instruction.">ECL Plus Western blotting detection reagents instruction.</a> , .</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimation+of+protein+molecular+weights+by+SDS+gel+electrophoresis,+GE+Healthcare+Website.">Estimation of protein molecular weights by SDS gel electrophoresis, GE Healthcare Website.</a> , .</li><li>Yeromin, A. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+identification+of+the+CRAC+channel+by+altered+ion+selectivity+in+a+mutant+of+Orai.">Molecular identification of the CRAC channel by altered ion selectivity in a mutant of Orai.</a> Nature. 443, 226-229 (2006).</li><li>Zhang, S. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Store-dependent+and+-independent+modes+regulating+CRAC+channel+activity+of+human+Orai1+and+Orai3.">Store-dependent and -independent modes regulating CRAC channel activity of human Orai1 and Orai3.</a> J. Biol. Chem. 283, 17662-17671 (2008).</li><li>Penna, , et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+CRAC+channel+consists+of+a+tetramer+formed+by+Stim-induced+dimerization+of+Orai+dimers.">The CRAC channel consists of a tetramer formed by Stim-induced dimerization of Orai dimers.</a> Nature. , .</li></ol>

The procedure presented here uses a Bis-Tris gel with MES running buffer as an example of denaturing PAGE using the Invitrogen NuPAGE Novex electrophoresis system. In addition, this pre-cast gel system allows protein separation under denaturing or non-denaturing conditions as well as accomodates a wide range of molecular weights (from 1-200 kDa for the Bis-Tris gels to 36-400 kDa for the Tris-Acetate gels). Depending on your experiment, you may choose to follow only part of the western-blotting technique described here a...

western blotting using the invitrogen nupage novex bis tris minigels

Western Blotting (or immunoblotting) is a standard laboratory procedure allowing investigators to verify the expression of a protein, determine the relative amount of the protein present in different samples, and analyze the results of co-immunoprecipitation experiments. In this method, a target protein is detected with a specific primary antibody in a given sample of tissue homogenate or extract. Protein separation according to molecular weight is achieved using denaturing SDS-PAGE. After transfer to a membrane, the target protein is probed with a specific primary antibody and detected by chemiluminescence.
Since its first description, the western-blotting technique has undergone several improvements, including pre-cast gels and user-friendly equipment. In our laboratory, we have chosen to use the commercially available NuPAGE electrophoresis system from Invitrogen. It is an innovative neutral pH, discontinuous SDS-PAGE, pre-cast mini-gel system. This system presents several advantages over the traditional Laemmli technique including: i) a longer shelf life of the pre-cast gels ranging from 8 months to 1 year; ii) a broad separation range of molecular weights from 1 to 400 kDa depending of the type of gel used; and iii) greater versatility (range of acrylamide percentage, the type of gel, and the ionic composition of the running buffer).
The procedure described in this video article utilizes the Bis-Tris discontinuous buffer system with 4-12% Bis-Tris gradient gels and MES running buffer, as an illustration of how to perform a western-blot using the Invitrogen NuPAGE electrophoresis system. In our laboratory, we have obtained good and reproducible results for various biochemical applications using this western-blotting method.

Watch this Scientific Journal Video about Western Blotting Using the Invitrogen NuPage Novex Bis Tris MiniGels at JoVE.com

Western Blotting Using the Invitrogen NuPage Novex Bis Tris MiniGels

This technical article describes a standard western-blotting procedure using the commercially available NuPAGE electrophoresis Mini-Gel system from Invitrogen.

Western Blotting (or immunoblotting) is a standard laboratory procedure allowing investigators to verify the expression of a protein, determine the ...

western-blotting-using-the-invitrogen-nupage-novex-bis-tris-minigels

Research

JoVE Journal

Biology

37.2K vues.  University of California, Irvine (UCI). Cet article technique décrit un standard occidental buvard procédure en utilisant l'électrophorèse commercialement disponibles NuPAGE Mini-Gel du système d'Invitrogen.

Vidéo: Western Blotting Using the Invitrogen NuPage Novex Bis Tris MiniGels

Western Blotting: Sample Preparation to Detection

Western blotting is an analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target protein.

Western blotting is an analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract.

Western blotting is an analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract. It uses gel ...

Solid Phase Synthesis of a Functionalized Bis-Peptide Using "Safety Catch" Methodology

In 1962, R.B. Merrifield published the first procedure using solid-phase peptide synthesis as a novel route to efficiently synthesize peptides. This technique quickly proved advantageous over its solution-phase predecessor in both time and labor. Improvements concerning the nature of solid support, the protecting groups employed and the coupling methods employed over the last five decades have only increased the usefulness of Merrifield's original system. Today, use of a Boc-based protection and base/nucleophile cleavable resin strategy or Fmoc-based protection and acidic cleavable resin strategy, pioneered by R.C. Sheppard, are most commonly used for the synthesis of peptides1.
Inspired by Merrifield's solid supported strategy, we have developed a Boc/tert-butyl solid-phase synthesis strategy for the assembly of functionalized bis-peptides2, which is described herein. The use of solid-phase synthesis compared to solution-phase methodology is not only advantageous in both time and labor as described by Merrifield1, but also allows greater ease in the synthesis of bis-peptide libraries. The synthesis that we demonstrate here incorporates a final cleavage stage that uses a two-step "safety catch" mechanism to release the functionalized bis-peptide from the resin by diketopiperazine formation.
Bis-peptides are rigid, spiro-ladder oligomers of bis-amino acids that are able to position functionality in a predictable and designable way, controlled by the type and stereochemistry of the monomeric units and the connectivity between each monomer. Each bis-amino acid is a stereochemically pure, cyclic scaffold that contains two amino acids (a carboxylic acid with an &alpha;-amine)3,4. Our laboratory is currently investigating the potential of functional bis-peptides across a wide variety of fields including catalysis, protein-protein interactions and nanomaterials.

Solid Phase Synthesis of a Functionalized Bis-Peptide Using &quot;Safety Catch&quot; Methodology

The efficient solid-phase peptide synthesis of a functionalized bis-peptide trimer utilizing a "safety catch" cleavage procedure from HMBA resin is described.

In 1962, R.B. Merrifield published the first procedure using solid-phase peptide synthesis as a novel route to efficiently synthesize peptides. This ...

Using High Resolution Computed Tomography to Visualize the Three Dimensional Structure and Function of Plant Vasculature

High resolution x-ray computed tomography (HRCT) is a non-destructive diagnostic imaging technique with sub-micron resolution capability that is now being used to evaluate the structure and function of plant xylem network in three dimensions (3D) (e.g. Brodersen et al. 2010; 2011; 2012a,b). HRCT imaging is based on the same principles as medical CT systems, but a high intensity synchrotron x-ray source results in higher spatial resolution and decreased image acquisition time. Here, we demonstrate in detail how synchrotron-based HRCT (performed at the Advanced Light Source-LBNL Berkeley, CA, USA) in combination with Avizo software (VSG Inc., Burlington, MA, USA) is being used to explore plant xylem in excised tissue and living plants. This new imaging tool allows users to move beyond traditional static, 2D light or electron micrographs and study samples using virtual serial sections in any plane. An infinite number of slices in any orientation can be made on the same sample, a feature that is physically impossible using traditional microscopy methods.
Results demonstrate that HRCT can be applied to both herbaceous and woody plant species, and a range of plant organs (i.e. leaves, petioles, stems, trunks, roots). Figures presented here help demonstrate both a range of representative plant vascular anatomy and the type of detail extracted from HRCT datasets, including scans for coast redwood (Sequoia sempervirens), walnut (Juglans spp.), oak (Quercus spp.), and maple (Acer spp.) tree saplings to sunflowers (Helianthus annuus), grapevines (Vitis spp.), and ferns (Pteridium aquilinum and Woodwardia fimbriata). Excised and dried samples from woody species are easiest to scan and typically yield the best images. However, recent improvements (i.e. more rapid scans and sample stabilization) have made it possible to use this visualization technique on green tissues (e.g. petioles) and in living plants. On occasion some shrinkage of hydrated green plant tissues will cause images to blur and methods to avoid these issues are described. These recent advances with HRCT provide promising new insights into plant vascular function.

High resolution x-ray computed tomography (HRCT) is a non-destructive diagnostic imaging technique that can be used to study the structure and function of plant vasculature in 3D. We demonstrate how HRCT facilitates exploration of xylem networks across a wide range of plant tissues and species.

High resolution x-ray computed tomography (HRCT) is a non-destructive diagnostic imaging technique with sub-micron resolution capability that is now being ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

Differentiation of Newborn Mouse Skin Derived Stem Cells into Germ-like Cells In vitro

Studying germ cell formation and differentiation has traditionally been very difficult due to low cell numbers and their location deep within developing embryos. The availability of a "closed" in vitro based system could prove invaluable for our understanding of gametogenesis. The formation of oocyte-like cells (OLCs) from somatic stem cells, isolated from newborn mouse skin, has been demonstrated and can be visualized in this video protocol. The resulting OLCs express various markers consistent with oocytes such as Oct4 , Vasa , Bmp15, and Scp3. However, they remain unable to undergo maturation or fertilization due to a failure to complete meiosis. This protocol will provide a system that is useful for studying the early stage formation and differentiation of germ cells into more mature gametes. During early differentiation the number of cells expressing Oct4 (potential germ-like cells) reaches ~5%, however currently the formation of OLCs remains relatively inefficient. The protocol is relatively straight forward though special care should be taken to ensure the starting cell population is healthy and at an early passage.

Differentiation of Newborn Mouse Skin Derived Stem Cells into Germ-like Cells In vitro

In recent years the formation of germ cells using in vitro culture methods has been demonstrated using several different types of somatic stem cells. This article will visually present the differentiation of newborn mouse derived stem cells to early oocyte-like cells.

Studying  germ cell formation and differentiation has traditionally been very difficult  due to low cell numbers and their location deep within developing ...

V3 Stain-free Workflow for a Practical, Convenient, and Reliable Total Protein Loading Control in Western Blotting

The western blot is a very useful and widely adopted lab technique, but its execution is challenging. The workflow is often characterized as a &#34;black box&#34; because an experimentalist does not know if it has been performed successfully until the last of several steps. Moreover, the quality of western blot data is sometimes challenged due to a lack of effective quality control tools in place throughout the western blotting process. Here we describe the V3 western workflow, which applies stain-free technology to address the major concerns associated with the traditional western blot protocol. This workflow allows researchers: 1) to run a gel in about 20-30 min; 2) to visualize sample separation quality within 5 min after the gel run; 3) to transfer proteins in 3-10 min; 4) to verify transfer efficiency quantitatively; and most importantly 5) to validate changes in the level of the protein of interest using total protein loading control. This novel approach eliminates the need of stripping and reprobing the blot for housekeeping proteins such as &#946;-actin, &#946;-tubulin, GAPDH, etc. The V3 stain-free workflow makes the western blot process faster, transparent, more quantitative&#160;and reliable.

V3 workflow is a western blot procedure using stain-free gels. The stain-free technology allows researchers to visualize protein separation quality, to verify the transfer efficiency, and most importantly, to validate the change in the protein of interest using total protein quantification as a reliable loading control.

The western blot is a very useful and widely adopted lab technique, but its execution is challenging. The workflow is often characterized as a &#34;black ...

The Fastest Western in Town: A Contemporary Twist on the Classic Western Blot Analysis

The Western blot techniques that were originally established in the late 1970s are still actively utilized today. However, this traditional method of Western blotting has several drawbacks that include low quality resolution, spurious bands, decreased sensitivity, and poor protein integrity. Recent advances have drastically improved numerous aspects of the standard Western blot protocol to produce higher qualitative and quantitative data. The Bis-Tris gel system, an alternative to the conventional Laemmli system, generates better protein separation and resolution, maintains protein integrity, and reduces electrophoresis to a 35 min run time. Moreover, the iBlot dry blotting system, dramatically improves the efficacy and speed of protein transfer to the membrane in 7 min, which is in contrast to the traditional protein transfer methods that are often more inefficient with lengthy transfer times. In combination with these highly innovative modifications, protein detection using infrared fluorescent imaging results in higher-quality, more accurate and consistent data compared to the standard Western blotting technique of chemiluminescence. This technology can simultaneously detect two different antigens on the same membrane by utilizing two-color near-infrared dyes that are visualized in different fluorescent channels. Furthermore, the linearity and broad dynamic range of fluorescent imaging allows for the precise quantification of both strong and weak protein bands. Thus, this protocol describes the key improvements to the classic Western blotting method, in which these advancements significantly increase the quality of data while greatly reducing the performance time of this experiment.

This protocol explores the latest advancements in performing Western blot analyses.&#160;These novel modifications employ a Bis-Tris gel system with a 35&#160;min electrophoresis run time, a 7&#160;min dry blotting transfer system, and infrared fluorescent protein detection and imaging that generates higher resolution, quality, sensitivity, and improved accuracy of Western data.

The Western blot techniques that were originally established in the late 1970s are still actively utilized today. However, this traditional method of ...

A Guide to Modern Quantitative Fluorescent Western Blotting with Troubleshooting Strategies

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins within complex biological samples. Since then, western blotting methodology has become a common component of the molecular biologists experimental repertoire. A cursory search of PubMed using the term &#8220;western blot&#8221; suggests that in excess of two hundred and twenty thousand published manuscripts have made use of this technique by the year 2014. Importantly, the last ten years have seen technical imaging advances coupled with the development of sensitive fluorescent labels which have improved sensitivity and yielded even greater ranges of linear detection. The result is a now truly Quantifiable Fluorescence based Western Blot (QFWB) that allows biologists to carry out comparative expression analysis with greater sensitivity and accuracy than ever before. Many &#8220;optimized&#8221; western blotting methodologies exist and are utilized in different laboratories. These often prove difficult to implement due to the requirement of subtle but undocumented procedural amendments. This protocol provides a comprehensive description of an established and robust QFWB method, complete with troubleshooting strategies.

The advancement of western blotting using fluorescence has allowed detection of subtle changes in protein expression enabling quantitative analyses. Here we describe a robust methodology for detection of a range of proteins across a variety of species and tissue types. A strategy to overcome common technical problems is also provided.

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins ...

Method for the Assessment of Effects of a Range of Wavelengths and Intensities of Red/near-infrared Light Therapy on Oxidative Stress In Vitro

Red/near-infrared light therapy (R/NIR-LT), delivered by laser or light emitting diode (LED), improves functional and morphological outcomes in a range of central nervous system injuries in vivo, possibly by reducing oxidative stress. However, effects of R/NIR-LT on oxidative stress have been shown to vary depending on wavelength or intensity of irradiation. Studies comparing treatment parameters are lacking, due to absence of commercially available devices that deliver multiple wavelengths or intensities, suitable for high through-put in vitro optimization studies. This protocol describes a technique for delivery of light at a range of wavelengths and intensities to optimize therapeutic doses required for a given injury model. We hypothesized that a method of delivering light, in which wavelength and intensity parameters could easily be altered, could facilitate determination of an optimal dose of R/NIR-LT for reducing reactive oxygen species (ROS) in vitro.
Non-coherent Xenon light was filtered through narrow-band interference filters to deliver varying wavelengths (center wavelengths of 440, 550, 670 and 810nm) and fluences (8.5 x 10-3 to 3.8 x 10-1 J/cm2) of light to cultured cells. Light output from the apparatus was calibrated to emit therapeutically relevant, equal quantal doses of light at each wavelength. Reactive species were detected in glutamate stressed cells treated with the light, using DCFH-DA and H2O2 sensitive fluorescent dyes.&#160;
We successfully delivered light at a range of physiologically and therapeutically relevant wavelengths and intensities, to cultured cells exposed to glutamate as a model of CNS injury. While the fluences of R/NIR-LT used in the current study did not exert an effect on ROS generated by the cultured cells, the method of light delivery is applicable to other systems including isolated mitochondria or more physiologically relevant organotypic slice culture models, and could be used to assess effects on a range of outcome measures of oxidative metabolism.

Method for the Assessment of Effects of a Range of Wavelengths and Intensities of Red/near-infrared Light Therapy on Oxidative Stress In Vitro

Non-coherent Xenon light was passed through narrow-band interference and neutral density filters to deliver light of varying wavelength and intensity to cultured cells. This protocol was used to assess the effects of red/near-infrared light therapy on production of reactive species in vitro: no effects were observed using the tested parameters.

Red/near-infrared light therapy (R/NIR-LT), delivered by laser or light emitting diode (LED), improves functional and morphological outcomes in a range of ...

Imaging the Intracellular Trafficking of APP with Photoactivatable GFP

Beta-amyloid (A&#946;) is the major constituent of senile plaques found in the brains of Alzheimer&#8217;s disease patients. A&#946; is derived from the sequential cleavage of Amyloid Precursor Protein (APP) by &#946; and &#947;-secretases. Despite the importance of A&#946; to AD pathology, the subcellular localization of these cleavages is not well established. Work in our laboratory and others implicate the endosomal/lysosomal system in APP processing after internalization from the cell surface. However, the intracellular trafficking of APP is relatively understudied.
While cell-surface proteins are amendable to many labeling techniques, there are no simple methods for following the trafficking of membrane proteins from the Golgi. To this end, we created APP constructs that were tagged with photo-activatable GFP (paGFP) at the C-terminus. After synthesis, paGFP has low basal fluorescence, but it can be stimulated with 413 nm light to produce a strong, stable green fluorescence. By using the Golgi marker Galactosyl transferase coupled to Cyan Fluorescent Protein (GalT-CFP) as a target, we are able to accurately photoactivate APP in the trans-Golgi network. Photo-activated APP-paGFP can then be followed as it traffics to downstream compartments identified with fluorescently tagged compartment marker proteins for the early endosome (Rab5), the late endosome (Rab9) and the lysosome (LAMP1). Furthermore, using inhibitors to APP processing including chloroquine or the &#947;-secretase inhibitor L685, 458, we are able to perform pulse-chase experiments to examine the processing of APP in single cells.
We find that a large fraction of APP moves rapidly to the lysosome without appearing at the cell surface, and is then cleared from the lysosome by secretase-like cleavages. This technique demonstrates the utility of paGFP for following the trafficking and processing of intracellular proteins from the Golgi to downstream compartments.

While the transport of cell surface proteins is relatively easily studied, visualizing the trafficking of intracellular proteins is much more difficult. Here, we use constructs incorporating photoactivatable GFP and demonstrate a method to accurately follow the amyloid precursor protein from the Golgi apparatus to down-stream compartments and follow its clearance.

Beta-amyloid (A&#946;) is the major constituent of senile plaques found in the brains of Alzheimer&#8217;s disease patients. A&#946; is derived from the ...