Advancing Plant-inspired Materials at University of Georgia with the Azure 200

Categories
Customer Spotlight Imaging

Customer Spotlight: Thomas Curry, Ph.D. Candidate, Urbanowicz Lab, University of Georgia

PhD candidate Thomas Curry has been researching how understanding the chemistry of plant cell walls can lead to the development of environmentally friendly petroleum-based fuels and plastic alternatives for over four years. He and other lab members use the Azure 200 Imager in the Urbanowicz Lab. The Lab is headed by Assistant Professor Breeana Urbanowicz and part of the Complex Carbohydrate Research Center, in the Department of Biochemistry & Molecular Biology at the University of Georgia (UGA).

Plastic pollution is an enormous problem facing the world. The UN Environmental Programme simply states: our planet is choking on plastic1. With a substantial amount of plastic used only once before being discarded, 400 million tons of non-biodegradable plastic waste are produced each year. Such catastrophic pollution is leading scientists like Curry to ask pressing questions.

Gel image shown on Azure 200 imager
Curry uses the Azure 200 to image Coomassie-stained protein gels and SYBR Safe–stained DNA gels.

Curry explains one objective of the research community studying plant cell walls is to build a more sustainable future by addressing plastic waste and global warming by engineering alternatives to plastics and biofuels. “A major goal in our field is to replace non-renewable resources with those sourced from plants,” Curry says.

How plant cells could be the answer to pollution

Plant cells are surrounded by carbohydrate-based cell walls. The Urbanowicz Lab believes of these cell wall components are as an underutilized renewable resource and chooses to focus on achieving a complete understanding of the biochemistry of cell wall biosynthesis. There is a large amount of diversity among cell walls across the plant kingdom, meaning Curry and his lab mates must study a wide variety of plants. He works with proteins from trees to moss and everything in between.

“I am inspired by the passion of people I have met who care about solving these unprecedented issues. I am hopeful to see the work that I do have a tangible impact on the world.”

Work in the lab involves the classic plant model system, Arabidopsis thaliana, as well as poplar, switchgrass, duckweed, algae, and carbohydrates from a variety of food crops. Eventually, the knowledge gained may allowed for the engineering of other plants to produce biofuels and other biomaterials.

Curry’s main work is focused on reconstituting enzymatic pathways responsible for the synthesis of cell wall polysaccharides and biochemicals in vitro. Ultimately, his work may allow large-scale synthesis of cell wall components. These chemicals are currently too rare and expensive to produce by currently available technology.

A review of Curry’s recent published work on xylan structure and biosynthesis can be found here.

Imaging coomassie-stained protein gels and SYBR Safe-stained DNA gels using the Azure 200

Curry’s work involves recombinant expression of plant enzymes, an experimental workflow that requires running numerous DNA and protein gels to track expression. Because these proteins are often glycosylated, they frequently cannot be expressed successfully in bacterial expression systems. Instead, they are expressed in human cell culture.

In addition to appreciating the color imaging capacity the Azure 200 has brought to their documentation, the system has another customization level that Curry has grown to love. He jokes that the instrument has become a favorite in the lab because of its “ability to change the background to funny pictures of each other.” Shown below is Curry next to the lab’s imager, with an image of his PI as the background. The Azure 200 is also able to image, edit, and store data.

Thomas Curry, brunette male, wearing University of Georgia hoodie, standing next to Azure 200 Imager
PhD Candidate Thomas Curry pictured next to the Azure 200. In addition to being field upgradeable, this gel doc is also capable of acute customization, including the ability to change the imager's background.

A special combination of basic research and practical applications keeps Curry interested in his line of research work. “The process of optimizing and troubleshooting the reactions is like a puzzle,” he says. “It’s a great feeling when it all comes together.”

See what the Urbanowicz Lab is up to and stay updated on their research into cell wall biosynthetic pathways on their website.

Gel Imaging Made Simple

The Azure 200 Imager used by Curry at UGA is an upgradeable, simple, touchscreen-based gel documentation imaging system. It is designed for UV, color imaging, blue-excited DNA dyes, and Coomassie gel imaging. Field upgrades are available for chemiluminescent, RGB fluorescent and NIR fluorescent applications.

SOURCES

  1. “Visual Feature | Beat Plastic Pollution.” UNEP, https://www.unep.org/interactives/beat-plastic-pollution/. Accessed 7 October 2023.

10 Top Tips for Converting Western Blots from Chemiluminescence to Multiplex Fluorescence

Categories
Fluorescence imaging Multiplex Western Blotting

Chemiluminescence is the most familiar method of detection for Western blotting and offers great sensitivity; however, many scientific questions and experimental designs require the additional information provided by fluorescent Western blotting. Fluorescent detection provides precise quantitation and visualization of similarly sized proteins within the same sample. If you want to convert from a chemiluminescent Western blot protocol to a multiplex fluorescent Western blot protocol, you’ll need to carefully consider and optimize your current protocol for the best results. To ensure this change is done effectively, first review this standard chemiluminescent Western blot protocol before beginning to pre-optimize a fluorescent protocol.

In this blog post, we share our ten top tips to help you successfully convert from chemiluminescence to multiplex fluorescent Western blots (shown below).

Digital image of 3-color fluorescent multiplexed Western blot
Digital image of 3-color fluorescent multiplexed Western blot using Azure Biosytems c600 imager. Lanes (from left to right) loaded with 1, 2, 5, 10, 20 µg HeLa cell lysate. Probed for tubulin (top), beta actin (middle) and GAPDH (bottom).

Table of Contents

Tip 1: Dilute your ladder

With a fluorescent protocol, only a small amount of ladder is needed. Many brands of the molecular weight ladder need to be diluted 1:10 in loading buffer. This will result in a very faint ladder on the blot, which may cause alarm at first. Know that this is expected and required for high quality fluorescent Western images.

Tip 2: Get rid of the dye front

The dye front can cause background fluorescence. Because of this, be sure to either let the dye front run off the gel during electrophoresis, or cut the dye front off the gel before transferring to the membrane.

Tip 3: Optimize transfer conditions

The type of transfer used can affect the outcome of a multiplex fluorescent blot. Using a wet transfer is ideal as it allows for optimization and quantitative analysis  due to their high customizability. To ensure complete transfer of a broad range of proteins, the time, temperature, voltage, and buffer can be varied based on the protein of interest. With the semi-dry transfer method, the transfer time cannot be extended to allow for more protein transfer and has the potential to dry out due to the limited amount of buffer used. This means that with a semi-dry transfer, proteins at either extreme of the size range will have difficulty transferring.

  • Quick Tip: Pre-soak the low fluorescence PVDF membrane in methanol to activate the membrane for transfer.

PVDF membranes are best for fluorescent Western blots. To ensure background fluorescence is not an issue, use low fluorescence PVDF membranes. These ready-to-use PVDF membranes are conveniently available in three pre-cut sizes for you to choose from.

Preparing to transfer an SDS-PAGE gel to a membrane for a Western blot.
Preparing to transfer an SDS-PAGE gel to a membrane for a Western blot using ready-to-use PVDF membranes from Azure Biosystems.

Once the transfer is complete, do not write with ink or pencil on the membrane; the graphite and ink will bleed and autofluoresce. After the transfer, do not expose the membrane to any container that has ever been exposed to Coomassie stain. Only use clean, dedicated Western blot trays. 

Tip 4: Take the guesswork out of blocking

Blocking fluorescent Westerns can be tricky. Initially, using a dedicated fluorescent blot blocking buffer, like this one, makes the transition to fluorescence from chemiluminescence much simpler and easier.  We highly recommend using Azure Fluorescent Blot Blocking Buffer, as it stabilizes fluorescent signals and comes ready to use. Once you have optimized other parts of your protocol, then you can test milk and other blocking buffers.

Read more: Getting Rid of the Noise: Western Blot Blocking

Tip 5: Skip Ponceau staining

Ponceau stain can increase background fluorescence. Since our end goal is a beautiful fluorescent Western blot, avoid any reagents because using them could increase background fluorescence.

  • Quick Tip: After blocking, never do a Ponceau stain on the membrane.

Tip 6: Ensure the primary antibodies are optimized

Before transitioning to a multiplex fluorescent protocol, evaluate how your current Western results look. Which proteins do you intend to multiplex together? Are the current antibodies you are using for the different proteins resulting in clean, sharp bands with low background? If not, optimizing the primary antibodies is necessary.

Direct antibody labelling allows the use of antibodies from the same species or of the same isotype and reduces the potential for cross reactivity.

Read more: Antibody Labeling in the Lab

Tip 7: Use the right wash buffer

The differences in image quality caused by wash buffer choice alone can be dramatic. Using a buffer that is specific for fluorescent Westerns is ideal here. Check out this fluorescent wash buffer. It’s perfectly optimized for use with fluorescent secondary antibodies.

See the difference using a fluorescent wash buffer can make for yourself by preparing two blots in parallel using different types of buffer. We also show this in the comparison images below.

Comparison of using fluorescent Washing Buffer vs. insufficient wash
Azure Fluorescent Blot Washing Buffer is optimized for use with both visible and near-infrared fluorescent Western blots. It is specially formulated to reduce non-specific binding of fluorescent secondary antibodies to produce clean blots with high signal-to-noise ratios.

If you don’t have this premade buffer readily available, using TBST with 0.1% Tween20 is an acceptable substitute.

  • Quick Tip: The final wash step should be performed with either TBS or PBS with no detergent. After the final wash, dry the membrane by air drying on a quenching sheet in a dark cabinet or drawer. Dipping the membrane in methanol can help it dry more quickly.

Tip 8: Optimize your proteins for fluorescent detection

Typically, proteins that require less than 3 minute exposures with film can be detected reasonably well with fluorescence. If you are currently using a strong ECL substrate with greater than 5 minute exposures to detect the protein of interest, you will likely need to optimize your protein fluorescence detection to get optimal results.

Optimizing your proteins is easy to do with fluorescent Western blotting kits, such as this one. It is a full kit with everything you need to perform ten two-color fluorescent Western blots.

AzureSpectra Fluorescent Western Blotting Kits with Fluorescent Block Each kit contains enough materials for 10 fluorescent Western blots: 9x7cm Low Fluorescence PVDF Membranes, 10 membranes 10x Fluorescent Blot Washing Buffer, 250mL 1x Fluorescent Blot Blocking Buffer, 300mL Background Quenching Sheets, 2 sheets 2 Fluorescent Secondary Antibodies, 40μL each
Fluorescent Western Blotting Kit from Azure contains enough materials and reagents for 10 fluorescent Western blots.

Tip 9: Protect from light

Once the fluorescent secondary antibodies have been added to the membrane, protecting the membrane from light is crucial. During washes and imaging, keep exposure to light to an absolute minimum to ensure the strongest fluorescent signal.

AzureSpectra secondary antibodies are labeled with fluorophores that emit light in visible and near-infrared wavelengths. AzureSpectra 490, 550, 650, 700 and 800 secondary antibodies offer unparalleled sensitivity and performance for immunoblotting applications when used in conjunction with Azure Imaging SystemsDue to low background autofluorescence in the near-infrared region, AzureSpectra 700 and 800 secondary antibodies can produce higher signal-to-noise ratios.

Tip 10: Choose the right fluorescent channel

A good rule is to detect the most abundant proteins with the IR700 channel You can use the IR800 channel to detect less abundant proteins. If both proteins being evaluated require less than a one-minute exposure on film, either channel may be used to detect either protein.

Choosing the correct fluorescent channel on the Azure 600 Imager.
The Azure 600 is the only system that offers two channel, laser-based IR and chemiluminescent detection, with the speed and sensitivity of film. It has the ability to image visible fluorescent dyes, standard EtBr and protein gels, and laser excitation for quantitative Western blot imaging in the NIR.

The Azure 600 is an imager that offers laser technology with two IR detection channels, enabling you to image more than one protein in an assay. It provides accurate and fast chemiluminescent detection, as well as the sensitivity, dynamic range, and linearity needed for quantitative blot analysis.

Use an imager with multiplex capabilities

Fluorescent Western blots are visualized using an imager, like the Azure Sapphire FLUsing an imager with multiple channels bypasses the need to strip and re-probe membranes. Newer imaging systems have sophisticated detectors with the ability to exhibit a broader dynamic range than film and therefore avoid signal saturation issues.

Azure Sapphire FL Biomolecular Imager with lid open
The Sapphire FL Biomolecular Imager is capable of high-resolution imaging and wide depth of field enable many sample types, including arrays, microarrays, Western blots, tissue slides, and small animals.

The Sapphire FL can detect up to four proteins on the same blot and overlapping bands can be quantified. This saves you money, reagents, and reduces your total imaging time.

We are committed to supporting your research and ensuring a seamless transition to multiplex fluorescence in your Western blotting experiments. By following these best practices and keeping these ten fluorescent multiplex tips in mind, we hope you’ll find that converting your existing chemiluminescent Western blot protocol into a multiplex fluorescent Western blot protocol can be both simple and successful. If you encounter any difficulties or have questions, please reach out to our team for further assistance.

Genetic Variants Impacting Cardiovascular Disease Revealed using Sapphire

Categories
Imaging Publication Spotlight

Researchers from the University of Puerto Rico employed the Azure Sapphire Biomolecular Imager to  explore the possible links between DNA variants and cardiovascular disease through their impact on DNA binding. Peña-Martínez et al examined the relationship between NKX2-5 (a crucial transcription factor) and its DNA binding abilities. Utilizing predictive modeling and experimental assays, the team pinpointed specific non-coding variants that significantly modified NKX2-5’s affinity for DNA binding. These discoveries offer compelling evidence that disturbances in transcription factor binding sites could contribute to cardiovascular disease.

Since the release of this publication, the first generation Sapphire used has been succeeded by the new Sapphire FL, which was designed to be the flexible choice in bringing precise quantitation of nucleic acids and proteins. Learn more about this new imager.

Disease variants and the non-coding genome

A staggering 98% of our genome is composed of non-coding DNA. Through innovative genome-wide association studies (GWAS), scientists have determined that non-coding DNA harbors crucial regulatory elements (or cis-regulatory elements (CREs)), including promoters and enhancers, which play pivotal roles in orchestrating gene expression.

Tiny alterations in the DNA sequence are called single nucleotide polymorphisms (SNPs). SNPs are often situated in close proximity to genes. They influence the function of regulatory DNA-binding proteins, such as transcription factors, affecting gene activity and, ultimately, vital biological processes.

NKX2-5 and heart development

NKX2-5 is a key player in heart development. Previous research has demonstrated that mutations within its DNA-binding domain (DBD), specifically the homeodomain, can disrupt its regulatory function, leading to cardiovascular diseases like congenital heart diseases (CHDs). Interestingly enough, the majority of genetic variants associated with cardiovascular diseases are found not in the genes themselves, but rather in the CREs that function as binding sites for transcription factors involved in heart development.

The Peña-Martínez’s team at the University of Puerto Rico conducted a study to explore the influence of cardiovascular-associated SNPs on NKX2-5’s DNA binding abilities.

Since this paper was published, the Sapphire has been succeeded by the new Sapphire FL

Designed for flexible choice in detection chemistry and samples, the Sapphire FL brings precise quantitation of nucleic acids and proteins
Scientist changing optical modules on the new Azure Sapphire FL
The Sapphire FL Biomolecular Imager is capable of high-resolution imaging and wide depth of field enable many sample types, including arrays, microarrays, Western blots, tissue slides, and small animals.

Narrowing down non-coding SNPs for in vitro validation

Using a predictive model, the researchers identified over 8000 SNPs that were predicted to affect NKX2-5 DNA binding. After cross-referencing disease and quantitative trait-associated SNPs from the GWAS catalog, the five SNPs with the greatest potential magnitude were selected to test in vitro.  These SNPs are associated with traits that impact cardiovascular health, such as hemoglobin levels.

Non-coding mutations affect NKX2-5 binding

To study the effects of the SNPs on DNA binding, the researchers created a recombinant NKX2-5 homeodomain (the area of the protein responsible for DNA binding).

They created six test sequences, each composed of a 40bp genomic sequence with either a control nucleotide or one of the SNPs of interest at the center. To enable visualization, each sequence was conjugated with an IRDye® 700 fluorophore. An electrophoretic mobility shift assay (EMSA) was used to assess DNA binding affinity between the recombinant NKX2-5 homeodomain and test DNA sequences.

EMSA gels used to evaluated the effect of mutations

The researchers used the images acquired with the Sapphire to evaluate the effect of each mutation on the binding affinity of the recombinant NKX2-5 homeodomain. In Figure 3, the lower band represents free DNA, while the upper band shows bound DNA.

The researchers found that all five variants significantly altered NKX2-5’s affinity for binding DNA. These findings suggest disruptions in transcription factor binding sites may play a role in the development of cardiovascular diseases and thereby provides a mechanistic connection between genome variants and disease.

EMSA gel captured using Azure Sapphire Biomolececluar Imager
Figure 3 from Peña-Martínez et al. (2023) showing a representative EMSA gel used to validate the binding of the rs6121514 variant. The remaining panels are binding curves for all five of the variants. The Azure Sapphire Biomolecular Imager was used to capture images of the EMSA shown here. Licensed under CC BY 4.0.

While fluorescent labels were used here, traditional EMSA employs radioactive-labeled DNA which requires phosphorimaging for analysis. The Sapphire FL offers a distinct advantage by supporting both phosphor imaging and IR detection, allowing capture of any type of EMSA. This versatility provides flexibility and convenience throughout the experimentation process.

Designed for ultimate flexibility, the Sapphire FL allows for precise detection of proteins and nucleic acids. If you’re eager to explore the possibilities of this groundbreaking system for your research, schedule a demo today, and experience what the future of biomolecular imaging has to offer.

Have you published with an Azure instrument?

We’d love to read it! Email your publication to us and we’ll send you something for sharing.

SOURCE

  1. Peña-Martínez, Edwin G, et al. “Disease-associated non-coding variants alter NKX2-5 DNA-binding affinity.” Biochimica et Biophysica Acta (BBA) – Gene Regulatory Mechanisms,, vol. 1866, no. 1, 2023, pp. 1-5. Science direct, https://doi.org/10.1016/j.bbagrm.2023.194906.

Azure c600 used to detect fluorescence-based proanthocyanidins in situ

Categories
Fluorescence imaging Publication Spotlight

Proanthocyanidins (PAs) are found in plants and have a role in protecting plants against herbivores and fungal pathogens. They are also used in many industrial applications due to the importance of PAs’ function and dynamics in the plant cell walls; however, there is a lack of sufficient methods to analyze them in planta. A research team at the Swedish University of Agricultural Sciences found 4-dimethylaminocinnamaldehyde (DMACA) can be used as a PA-specific fluorescent dye to localize PAs in plant cells1. The Azure c600 Western blot imager served an important role in the study, as the team used it to visualize and validate using DMACA to localize PAs in leaves. Prior to this study, how PAs incorporate into plant cell walls or what their functions under stress conditions were, was unknown.

Since the release of this publication, the cSeries Imaging Systems have been succeeded by the new Azure Imaging Systems. The upgraded systems are high performance instruments capable of NIR fluorescence, visible fluorescence, and chemiluminescence.

Fluorescent detection method for PAs

To understand the dynamics of PAs in cell walls, high-resolution microscopy is required, along with fluorescent dyes that can identify PAs. Common methods for detecting PAs in plants requires maceration of the plant tissue for processing, ruling out in situ methods. Chowdhury et al set out to identify a PA-specific fluorescent dye that could be used for in situ analysis of PAs. DMACA has been used before in light microscopy to identify PAs as its reaction with PA causes a blue precipitate, but the use of DMACA as a PA-specific fluorophore for high-resolution microscopy had not been evaluated. Chowdhury and team explored this possibility.

More Reading: Problems With Ponceau Stain? Consider Alternative Total Protein Stains for Fluorescent Western Blots

Investigating PA-specific fluorogenic properties of DMACA

Using commercially available PAs and isolated PAs from poplar roots, the researchers tested if DMACA would fluoresce in a PA-specific manner. Their results showed classic DMACA excitation and emission was only observed in samples containing PAs. With the confirmation that DMACA is PA-specific and fluoresces, they began to look for any potential interference from proteins found in the plant cell wall.

The researchers investigated whether DMACA fluoresces in the presence of isolated cell wall polymers, such as cellulose. To evaluate this, they implemented the fluorescence spot test (FST) method. PAs and cell wall polysaccharides were mixed with the DMACA reagent and before being added to a PVDF membrane. An Azure c600 was used to acquire images using three RGB channels: Cy2, Cy3, and Cy5 (Figure 2).

The Ultimate Western Blot Imaging System

The Azure 600 offers laser technology with two IR detection channels enabling you to image more than one protein in an assay. It provides accurate and fast chemiluminescent detection, as well as the sensitivity, dynamic range, and linearity needed for quantitative blot analysis.
Scientist choosing settings on Azure 600
The Azure 600 is the only system that offers two channel laser based IR detection, chemiluminescent detection with the speed and sensitivity of film, and the ability to image visible fluorescent dyes, standard EtBr and protein gels.

Chowhury et al confirmed their previous notion of the existence of a PA-dependent shift from Cy3 to Cy5 with DMACA, where the fluorescence intensity was dependent on PA concentration. Using the semi-quantitative FST method, we now know plant polymers do not interfere with PA-specific DMACA fluorescence in the Cy 5 channel. Chlorophyll, however, proved troublesome, since the emission range of PA-specific DMACA overlaps with the chlorophyll’s autofluorescence.

Results of the fluorescence spot test using Azure c600 Western blot imager
Figure 2 from Chowhury et al 2022 representing the results of the fluorescence spot test (FST) method. The DMACA fluorescence with different PA species and cell wall polysaccharides at various dilutions. Fluorescence was detected with the Azure c600 imager.

After using ethanol to remove chlorophyll, the researchers used the Azure c600 to examine the blue precipitate resulting from DMACA-PA interaction in leaves under both white light and fluorescence (Figure 3). Overall, the fluorescence corresponded to the areas with the blue precipitate, successfully indicating DMACA could be used to detect PAs in planta.

Bright light and fluorescent images of leaves exposed to DMACA
Figure 3 from Chowdhury et al. 2022. The bright light and fluorescent images of leaves exposed to DMACA captured using Azure c600. The blue precipitate shows DMACA interacting with PAs (D) and the red fluorescent areas (E) overlap these blue areas.

PA-specific DMACA used to study PAs in situ in plant roots

Chowdhury and team then looked at DMACA for use in fluorescence microscopy for in situ analysis of PAs in root tissue. Under bright light microscopy, the DMACA-PA blue precipitate reaction method was used once more to locate the PAs exact location in the root tip. The team found fluorescence overlapped with blue precipitate areas, indicating the presence of PAs in the roots. In the brightfield images, their findings showed fluorescence was more prominent than the blue coloration; DMACA fluorescence is a sensitive way to visualize cell-wall bound PAs. The team concluded PA-specific DMACA signal both is highly stable and, while it can be useful in leaves, more suitable for use in roots.

Upon proving the ability of DMACA to serve as a fluorophore specific for PAs, further investigation of the co-localization of PAs with cell wall polymers was needed. PAs overlapped significantly with other cell wall polymers in older portions of the root, but were absent from the newly formed root tip. The results from this paper indicate PAs are incorporated later in development.

Have you published with an Azure instrument?

We’d love to read it! Email your publication to us and we’ll send you something for sharing.

What are PAs and why are they important?

PAs are commonly found in woody plants, such as grapes, and forest trees. They are an end product of the flavonoid biosynthetic pathway and act as plant pigments and may prevent cancer2. PAs are involved in plant defense and protect plants by acting against leaf-eating herbivores, fungal pathogens, and UV damage.

Takeaways from this study

In this groundbreaking study, Chowdhury et al was able to characterize the fluorogenic properties of DMACA as a PA-specific fluorophore and validate its use in fluorescence microscopy as they studied PAs in situ and in planta. The team identified a novel tool which can be used to study PAs and their dynamics at the subcellular level. This study opens the door for further study of PAs in plants.

SOURCES

  1. Chowdhury J, Ferdous J, Lihavainen J, Albrectsen BR, Lundberg-Felten J. Fluorogenic properties of 4-dimethylaminocinnamaldehyde (DMACA) enable high resolution imaging of cell-wall-bound proanthocyanidins in plant root tissues. Front Plant Sci. 2023 Jan 16;13:1060804. doi: 10.3389/fpls.2022.1060804. PMID: 36726681; PMCID: PMC9884812.
  2. “Proanthocyanidins – Health Encyclopedia – University of Rochester Medical Center.” URMC, https://www.urmc.rochester.edu/encyclopedia/content.aspx?contenttypeid=19&contentid=proanthocyanidins. Accessed 11 March 2023.

Electrophoresis 101: the Difference between Running and Transfer Buffer

Categories
Electrophoresis

SDS-PAGE vs. Western blotting- what's the difference?

SDS-PAGE (Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis) is a commonly used technique for the separation of proteins based on their size. In SDS-PAGE, smaller proteins migrating more quickly through the gel inside of a transfer cell. The percentage of polyacrylamide in the gel determines how easily proteins of various sizes can move through the gel, with higher percentage gels having a tighter gel matrix better for resolving smaller proteins.

Western blotting is a commonly used method for detecting and analyzing the separated proteins, after being separated using SDS-PAGE.

The role of buffers in Western blotting

An important aspect of Western blotting is the use of buffers, which play a crucial role in the success of the experiment. Two types of buffers are commonly used in Western blotting: running buffer and transfer buffer. Understanding the difference (Table 1) between the two and what their specific functions are is crucial to a successful Western blotting experiment.

Table of contents

What's the difference between running buffer and transfer buffer?

For conducting nucleic acid or protein electrophoretic separations, running buffer is an essential, as it provides the required solutions.

Transfer buffer is a solution used in the process of Western blotting. It is used to transfer the separated proteins from the gel to a solid support, such as a nitrocellulose or PVDF (Polyvinylidene Fluoride) membrane. The transfer step allows for the detection and analysis of specific proteins of interest. Keep reading for ingredients and guidelines on how to use each type of buffer.

Table 1. The difference between running and transfer buffer

What it isWhen it's used
Running bufferRunning buffer is a solution used in electrophoresis, specifically in SDS-PAGE.Running buffer's main purpose is to create an electric field that allows for the movement of proteins through the gel during the electrophoresis process.
Transfer bufferTransfer buffer serves as a conductive medium to facilitate the transfer process. It is designed to preserve a consistent pH level, aiding the elution of proteins from the gel, as well as their binding to the membrane.Transfer buffer is used in the process of Western blotting and electrophoretic transfer.

How do you make running buffer?

Running buffer is made up of a mixture of SDS and a salt, such as Tris-HCl, which helps to maintain the pH and conductivity of the solution. The concentration of SDS in the running buffer is usually around 1% and helps to uniformly denature and negatively charge the proteins, allowing for their separation based on size and not charge1.

To make the running buffer, mix the following ingredients in distilled water:

  • 10X

    SDS running buffer concentrate (e.g. 1 M Tris-HCI, 0.1 M SDS)
  • 50 mM

    Tris-HCI (pH 8.3)
  • 1%

    SDS
  • Distilled water to make the final volume of the solution

pH variations can affect the separation of proteins, which is why the pH of the running buffer should be checked and adjusted to 8.3 before use. The running buffer should be pre-heated to 70-75°C to solubilize the SDS when first making it, before cooling it down to room temperature.

Can I reuse running buffer?

Reusing running buffer is generally not recommended because it can affect the accuracy of protein separation in SDS-PAGE. Running buffer becomes contaminated with protein fragments and other debris after each electrophoresis run, which can interfere with subsequent runs and lead to unreliable results. Additionally, the SDS and other components in the buffer can degrade over time, affecting its effectiveness.

For these reasons, we recommend preparing fresh running buffer for each experiment to ensure your results are accurate and consistent. It is possible to prepare and store a larger batch for a short period of time (e.g. a few days) in a refrigerated environment; however, you should still replace it regularly to save time.

How do you make transfer buffer?

Before jumping into mixing your own, you should know that transfer buffer can easily be purchased. The Azure transfer buffer is a proprietary transfer buffer that provides improved detection of low-abundance and post-translationally modified proteins. We offers free samples of it if you find yourself wanting to eliminate the time it takes to mix your own. High-efficiency protein transfer and increased protein retention on the membrane add up to more sensitive Western blots.

However, the answer to how to make transfer buffer is simple: it typically contains a combination of salt, such as Tris-HCl, and a reducing agent, such as methanol, to help maintain the stability of the proteins during transfer by dissociating SDS from the proteins. Methanol is also important for helping the proteins adhere to the membrane. Unlike running buffer, transfer buffer does not contain SDS, as its role is to preserve the proteins and not to denature them. The pH of the transfer buffer is usually around 7-8 to ensure optimal protein transfer and stability.

Transfer buffer should be prepared fresh for each experiment, because its effectiveness can be affected by long-term storage or repeated use. To ensure consistency and reliability, purchasing transfer buffer is a great idea. Using pre-made transfer buffer provides improved detection of low-abundance and post-translationally modified proteins. The combination of high-efficiency protein transfer and increased protein retention on the membrane amount to more sensitive Western blots. Using pre-made transfer buffer allows you to transfer proteins in less than 20 minutes vs. several hours required with traditional, homemade transfer buffers.

Mix your own transfer buffer using this recipe:

  • 50 nm

    Tris-HCI (pH 8.0)
  • 20%

    Methanol
  • 0.1%

    SDS
  • Distilled water to make the final volume of the solution

For additional buffer recipes, check out the app note Wet or Dry?

How many times can transfer buffer be used?

The number of times transfer buffer can be reused depends on several factors, including the composition of the buffer,  the size and stability of the proteins being transferred, and the conditions of storage and use. In general, reuse of transfer buffer is not recommended, as the efficiency of protein transfer can decrease with repeated use.

Proteins can become trapped in the gel or in the pores of the transfer membrane, reducing the overall efficiency of transfer. Additionally, the reducing agent in the transfer buffer can degrade over time, affecting its ability to preserve the proteins during transfer resulting in inaccurate results. The presence of contaminants or impurities in the buffer can also reduce its effectiveness.

For these reasons, we suggest you prepare fresh transfer buffer for each experiment to ensure your results are accurate and reliable. If cost or availability is a concern, transfer buffer can be stored for a short period of time (e.g., a few days) in a refrigerated environment, but it should still be replaced regularly to ensure optimal results.

Quick, 1-hour or transfers overnight at lower voltages

The Azure Aqua Transfer Cell is able to keep gels cool through a compatible ice pack in the buffer chamber. You can also place the entire gel apparatus in a cold room while it runs.
Loaded gel in electrophoresis for SDS-PAGE
Protein samples loaded into a gel inside an Azure Aqua Vertical Gel Running System

We hope after reading this article you are able to easily distinguish the difference between transfer and running buffers. If you still have questions regarding the steps of Western blotting, fill out the form below and someone from our team will assist. Additional resources on SDS page can be found below as well. Cheers!

Additional resources for gel electrophoresis and SDS-PAGE:

SOURCE
  1. Gavini K, Parameshwaran K. Western Blot. [Updated 2022 Apr 28]. In: StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing; 2022 Jan-. Available from: https://www.ncbi.nlm.nih.gov/books/NBK542290/

When to use ELISA vs Western blot

Categories
Immunoassay Protein Assays Western Blotting

ELISA (Enzyme-linked immunosorbent assay) and Western blotting are two of the most commonly used techniques in molecular biology for the detection and quantification of proteins. Although both techniques are based on the principle of binding specific antibodies to target proteins, they have different applications and advantages.

By weighing the pros and cons of both ELISA and Western blotting for protein analysis, you can make a more informed decision about which method is best for your experiment. Both are important techniques for detecting and quantifying proteins, but each technique has its own unique advantages and disadvantages.

In this blog post, we will discuss the key differences between ELISA vs Western blotting to help you decide when it is best to use each method.

Table of contents

What's the difference between ELISA and Western blot?

ELISAs are immunoassays carried out in 96-well plates as a simple, rapid, and highly sensitive method for measuring the presence of a specific protein in a sample. ELISAs are often used for qualitative and quantitative analysis of proteins in various sample types, such as serum, cell culture supernatants, and urine.

Western blotting is a powerful technique for the detection and characterization of specific proteins in a sample. The basic principle of Western blotting involves separating proteins in a sample using electrophoresis, transferring them to a nitrocellulose or PVDF membrane, and then probing the membrane with an antibody specific to the protein of interest. If the target protein is present, the antibody will bind to the protein. A secondary antibody allows for detection through one of two main methods: chemiluminescence or fluorescenceLet’s review the cases where you would use either application.

When to use ELISA for protein analysis

Now that we have covered the instances where you might find using ELISAs more beneficial over Western blots, let’s briefly discuss the disadvantages using ELISA may pose.

ELISAs have limited multiplexing capability; traditionally, only one protein can be targeted in each ELISA. This caveat makes it difficult for ELISA to simultaneously analyze multiple proteins in a single sample. In this case, we suggest using multiplex fluorescent Western blots. Using a fluorescent scanner like the Sapphire FL, you will save precious time (and $$$) on each multiplex fluorescent Western blot. Don’t just take our word for it, compare the costs here.

Table 1. Advantages of measuring levels of specific proteins by ELISA

Use ELISA when you...
Are looking to detect very small amounts of target proteins, or rare or low-abundance proteins. A major advantage of ELISA is its high sensitivity.
Need to measure the exact amount of target protein in a sample. ELISA is a valuable tool for determining protein concentration or monitoring changes in protein levels over time.​
Are in a time crunch in the lab. ELISAs are rapid, easy to perform, and can be done relatively quickly with minimal sample preparation. It is a convenient choice for routine assays.​

When to use Western blotting for protein analysis

Since ELISA is a high-throughput, sensitive, and simple assay, it’s ideal for detecting low-abundance proteins in large numbers of samples. So, when should you use Western blotting for protein analysis? Western blotting is a highly specific technique that is capable of resolving proteins into different molecular weight ranges, making it useful to detect possible protein modifications and subtle differences between experimental conditions.

Table 2. Advantages of using Western blotting over ELISA

Use Western blotting when you...
Need to detect a wide range of proteins. Western blots are valuable tools for analyzing complex mixtures, such as cell lysates. To take your analysis one step further, fluorescent multiplexing will allow you to detect multiple targets at once. The Azure 400 imager allows you to simultaneously detect 3 target proteins image using three-color fluorescence.​
Want to identify specific proteins. Western blots can be used to specifically identify the presence of a target protein in a sample, even when the protein is present at low levels, or in a complex mixture.​
Want to confirm the correct protein is being measured. Western blots can confirm proteins through a number of factors, such as protein size. It can also reveal if there are protein modifications, impurities, etc.

Table 2 shows the instances where you would use Western blotting over an ELISA test; however, keep in mind that unlike ELISA, Western blotting is a multi-step process that can take from several hours to overnight to perform. Western blotting is also less sensitive than ELISA. If you’re working with low-abundance proteins, stick with ELISA.

Want to run a Western blot but not sure where to start? Check out this checklist for everything you need to run a successful Western blot.

AdvantagesDisadvantages
Detect a wide range of proteins: Western blots are valuable tools for analyzing complex mixtures, such as cell lysates. Multiplexing using fluorescence allows for the detection of multiple targets in one experiment. Using the Azure 400 allows you to simultaneously detect 3 target proteins image using three-color fluorescence.Longer process with more steps involved: Western blotting is a multi-step process that can take several hours (often overnight) to perform
Ideal for identifying specific proteins: Western blots can be used to specifically identify the presence of a target protein in a sample, even when it is present at low levels or in a complex mixture.Less sensitive than ELISA: Western blotting is less sensitive than ELISA, making it more difficult to detect low-abundance proteins.
Gives additional information about the protein: A Western blot can confirm the correct protein is being measured by using a number of factors such as protein size. It can also reveal if there are protein modifications, impurities, etc.

How does an ELISA test work?

The basic principle of ELISA involves coating a solid substrate, such as a microplate, with an antibody that specifically binds the target protein. The sample potentially containing the target protein is then added to the coated microplate and allowed to incubate. If the target protein is present, it will bind to the antibody and can be detected using a second antibody conjugated to an enzyme that can cause a color change when a specific substrate is added. This allows for the detection of protein in the sample with a plate reader, like the Ao Absorbance Microplate ReaderThe amount of signal produced is directly proportional to the amount of target protein in the sample.

Azure Biosystems Ao Absorbance Microplate Plate Reader is used for ELISA, Bradford Assays, and more
Trust your data. With the Ao Microplate Reader you will not sacrifice speed for accuracy. With a read speed of <6 seconds, and an accuracy of <0.005 ± 1% (0-3) OD, you will quickly get your results and know they are correct.

The Ao conveniently comes with eight filters: five standard filters (405, 450, 492, 595, and 630 nm), and three filters you can personalize for your needs. We give you the ability to choose your own filters to always ensure you have what your assays require.

Western blot detection methods

Western blots can be visualized using chemiluminescence or fluorescence. In chemiluminescent Western blotting (example shown below), an HRP-conjugated secondary antibody binds to the primary antibody. When exposed to a detection substrate, like ECL, light is produced.

Light can be detected two ways:

  1. Using a chemiluminescent imager (like the chemiSOLO), or
  2. In the darkroom using film.
Chemiluminescent Western Blot imaged with Azure Imager
Chemiluminescent Western blot imaged with Azure Imager

For fluorescent Western blots, you will use secondary antibodies that are directly conjugated to fluorescent dyes. Fluorescent Western blots are typically only visualized using digital imagers, such as the Azure 400.

Multiplex fluorescent Western blot from Azure Biosystems imager
Digital image of 4-color multiplex Western Blot. Using distinct fluorescent and near-infrared targeting antibodies can detect each wavelength and merge them into a four-color multiplex image. No background noise or bleeding between channels.

The newer generation of imaging systems use sophisticated cameras that exhibit a broader dynamic range than film, thus avoiding the signal saturation problems that limit the dynamic range of film. For example, the Azure 600 imager comes standard with a 9.1MP camera which provides high-resolution imaging perfect for publications.

Simultaneously Detect Multiple Proteins

Using lasers for NIR fluorescent imaging sets the Azure Imagers apart from competitors. Our Imagers are the only ones on the market to use lasers.
Two scientists looking at multiplex fluorescent Western blot on Azure 600 Western blot imager

Choosing either method based on the needs of your experiment

When choosing between ELISA and Western blotting, consider the specific needs of your experiment and the questions you are trying to answer. Both techniques are valuable tools in your laboratory arsenal. Selecting the right one will depend on the specific requirements of your experiment. Out of ELISA and Western blot, which method do you most commonly use?

If you find yourself now in need of either a microplate reader or a digital imager, you’re in luck. Azure Biosystems is the leading supplier for life science systems. Check out this page to request pricing on the products mentioned in this post, the Ao Microplate Reader or Azure 400 Imaging System. If you’re unsure about what you need for protein analysis, fill out the form below and a product expert will be in touch. Our experts are available to help you choose the right system. Cheers for now.

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Azure 600 helps evaluate fusion protein affinity for SARS-CoV-2 spike protein variants

Categories
COVID-19 Imaging Publication Spotlight

In a recent study from Case Western Reserve University, Matthews et al. developed a laboratory-scale production process to better evaluate fusion proteins that can neutralize emergent severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2) variants using an Azure 600 to evaluate fusion protein affinity for spike protein variants. It is the only system that offers two channel laser based IR detection, chemiluminescent detection with the speed and sensitivity of film, and the ability to image visible fluorescent dyes, standard EtBr and protein gels.

The initial countermeasures against SARS-CoV-2 infection included vaccines, small molecule drugs, and neutralizing monoclonal antibodies. Since then, SARS-CoV-2 variants have emerged that could evade these initial countermeasures. Fusion proteins have been suggested as an ideal alternative that is not affected by variants. This work can help with fusion protein design and improving purification strategies to better study potential fusion protein treatment options.

Introduction to SARS-CoV-2, spike protein, ACE2, and treatment options

The spike (S) protein of SARS-CoV-2 infects human cells by binding to angiotensin enzyme 2 (ACE2)-expressing host cells. This leads to activation of the S protein and membrane fusion as the virus enters the cell. The ability of the S protein to bind to ACE2 is key for the virus’s pathology. The receptor binding domain (RBD) of the S protein is involved in the overall infectivity, immune evasion, and resulting transmissibility of SARS-CoV-2. Due to the high affinity of the RBD of the S protein for ACE2, a possible strategy for counteracting the virus is to use soluble ACE2 as a decoy. Ideally, ACE2 would compete for the S protein binding and thus sequester the virus to prevent it from entering cells.

More Reading: Sapphire Biomolecular Imager used in Investigation for Potential COVID-19 Nasal Vaccine

In vitro system for ACE2 fusion proteins

Previous research has that ACE2 fused with an IgG Fc region (ACE2-Fc) works best at sequestering the SARS-CoV-2 virus, but further characterization is still needed, such as determining the binding affinity for each SARS-CoV-2 variant.

Having an in vitro system to create and test ACE2-Fc proteins against SARS-CoV-2 is important to determine the best countermeasures against current and emerging variants. Matthews et. Al. (1) approached this by generating a cell line that stably expresses ACE2-Fc. The research team qualified the process for obtaining purified ACE2 fusion proteins, and created a system that allows researchers to test a hypothesis: can emerging variants that evade current countermeasures be neutralized by ACE2 fusion proteins?

Figure 1 from Matthews et al. showing upstream production of ACE2-Fc and ACE2(NN)-Fc from CHO cells. PLOS ONE 17(12): e0278294. To confirm the production of the fusion proteins from the CHO cells, immunoblots were used. Images were acquired with the Azure 600.

Creating a cell line to stably express ACE2 fusion proteins

ACE2 is part of the renin/angiotensin system, where angiotensin II regulates the cardiovascular system. ACE2 is shed into the plasma in its catalytically active form to regulate angiotensin II (2). The researchers engineered a mutant ACE2-Fc that would reduce potential effects of having an elevation of ACE2 when used as a decoy fusion protein. To abolish the enzymatic activity while retaining the binding capability to SARS-CoV-2, they mutated two residues (ACE2-(NN)-Fc).

In order to assess the binding affinity of these ACE2 fusion proteins with the S protein, researchers created a Chinese Hamster Ovary (CHO) cell line that stably expressed both ACE2-Fc and ACE2(NN)-Fc. They then confirmed the production of ACE2 fusion proteins using immunoblotting on the Azure 600 (Figure 1). Once confirmed, a laboratory-scaled protein production was performed. After purifying these recombinant proteins, Matthews et al. evaluated their stability under thermal stress, used mass spectrometry to look at their N-glycan profiles, and examined binding activity to the S protein and the different variants.

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Characterizing the ACE2 fusion proteins

Protein stability is important for patient safety and must be maintained during the manufacturing process. Matthews et al. evaluated protein stability following different thermal-stress conditions and did not find any changes in stability at 4°C or 25°C after 1 day; however, on day 7 at 25°C, the protein was barely detectable. It is important to note the sensitivity of the ACE2-Fc proteins to thermal-induced stress discovered as this information is important for future storage and handling protocols.

Another important characterization of these fusion proteins was the binding affinity for the spike protein variants. The results found demonstrated that both ACE2-Fc and ACE2(NN)-Fc fusion proteins are active in binding to S proteins. Interestingly, they have greater affinity for alpha, beta and delta S protein variants compared to the parental and omicron variants.

Takeaways

Though a clinical model for evaluating ACE2-Fc proteins as a therapeutic against SARS-CoV-2 does not currently exist, this study provides data supporting this option in the future. Matthews et al have provided data that could aid in the development of ACE2-Fc fusion proteins as a potential countermeasure against SARS-CoV-2. Furthermore, the information found can be used to support new ACE2 fusion protein designs, purification methods, and formulation studies.

Used by Case Western University

Azure 600 Western blot Imaging system
The Azure 600 is one of the five imaging systems in the Azure Imaging Systems lineup. It improves data quality imaging with infrared dyes and offers signal stability. Low background fluorescence imaging with NIR dyes allows researchers to study multiple proteins in a blot, even if those proteins overlap in molecular weight.

SOURCES

  1. Matthews AM, Biel TG, Ortega-Rodriguez U, Falkowski VM, Bush X, Faison T, et al. (2022) SARS-CoV-2 spike protein variant binding affinity to an angiotensin-converting enzyme 2 fusion glycoproteins. PLoS ONE 17(12): e0278294. https://doi.org/10.1371/journal.pone.0278294
  2. Turner AJ. ACE2 Cell Biology, Regulation, and Physiological Functions. The Protective Arm of the Renin Angiotensin System (RAS). 2015:185–9. doi: 10.1016/B978-0-12-801364-9.00025-0. Epub 2015 Apr 24. PMCID: PMC7149539. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7149539/

Gel Electrophoresis Steps

Categories
Electrophoresis SDS-PAGE Western Blotting

While the specific steps of gel electrophoresis may differ somewhat between running DNA/RNA gels and protein gels, overall, the steps are the same. In this post, we’ll go over the five steps of gel electrophoresis.

Table of contents

What is gel electrophoresis?

Gel electrophoresis is a method used in the lab to separate DNA, RNA, or proteins from one another. The molecules of interest are forced through a porous gel by an electrical current, with one end of the gel being positively charged and one end being negatively charged. This results in negatively charged molecules, like DNA and RNA, traveling toward the positive end of the gel. Since proteins can have a variety of charges, they must be neutralized using sodium dodecyl sulfate (SDS) to ensure the molecule separation is not affected by charge but is only due to size. SDS also denatures proteins, preventing variations in molecule shape from affecting migration patterns.

Due to the size of the pores in the gel, larger molecules do not travel as far as smaller molecules, allowing for size separation to occur. In the end, the separated molecules can be visualized as bands1.

What do you need for gel electrophoresis?

First, let’s go over everything you need for gel electrophoresis. While there are different types of gel electrophoresis, the same tools are required for each type2What you’ll need to begin gel electrophoresis:

  • Gel box

    The gel boxes differ depending on the type of gels being run. The optimal choice for DNA and RNA separation using agarose gels is horizontal gel electrophoresis. Vertical gel box systems, like the Azure Aqua Quad Mini Cell (shown below), are the best for separating proteins using polyacrylamide gels. The Aqua Quad Mini-Cell is designed for running 1–4 precast or handcast gels (cassette size 10cm x 8cm). It features locking side fasteners that provide a tight seal to ensure rapid and easy electrophoresis.

Loaded gel in electrophoresis for SDS-PAGE
Protein samples loaded into a gel inside an Azure Aqua Vertical Gel Running System
  • Gel

    Either pre-cast or hand-cast, with wells at the top of the gel for the samples to be loaded into prior to their migration through the gel. The gel is submerged in running buffer containing salt ions which conduct the electrical current throughout.

  • Running buffer

    Research the type of running buffer you will need and prepare it ahead of time. Buffers can often be purchased; however, they are also often made in bulk in the lab.

  • Power supply

    A power supply, like the Azure Aqua Power Supply (shown below), provides the electrical current through cables that connect to the positive and negative terminals of the gel box. It is equipped with four power connections that allow you to run multiple experiments at once, and can control constant power, current, or voltage.

Azure Aqua
The Azure Aqua Power Supply is a universal power supply that is designed for powering electrophoresis and transfer modules.
  • Visualization system

    Because DNA and proteins cannot be identified with the naked eye, there must also be a way to visualize them after separation. After transferring the gel to a membrane, use an Azure Imager, or another digital imaging system for further analysis.

Azure Imaging Systems allow you to stop wasting money on film. These modular, multichannel imagers are capable of UV, color imaging, blue-excited DNA dyes, silver-stain, Coomassie gel, fluorescence imaging, and more. Azure Imagers are come equipped with visible fluorescence, visible light, and UV excitation channels and are fully upgradeable to access a wide breadth of applications.

Compare each model in the Azure Imaging System lineup by clicking here.

Scientist choosing settings on Azure 600
The Azure 600 is the only system that offers two channel laser based IR detection, chemiluminescent detection with the speed and sensitivity of film, and the ability to image visible fluorescent dyes, standard EtBr and protein gels.

For quick, simple confirmation of the presence of the bands, a handheld UV light or light box can also be used. To detect individual proteins, antibodies specific to the proteins of interest must be used. Antibodies can be designed to be detected by either chemiluminescence or fluorescence. For chemiluminescence, the protein bands can be observed using a digital imager, or with film. Fluorescence signal detection requires an imager.

How long does gel electrophoresis take?

The run time for gel electrophoresis can vary anywhere from 45 to 90 minutes. The specific time needed to run a gel depends on a variety of factors, including:

  • the degree of separation desired,
  • the voltage applied, or
  • the gel’s composition.

Gel electrophoresis steps

Gel electrophoresis is a pretty straightforward process that involves preparing the samples in loading buffer, loading the gel box with running buffer, pipetting the samples into the wells, actually running the gel, and finally, visualizing your proteins.

STEP 1: Prepare samples

Samples will differ dramatically by individual experiment, but they all must be processed similarly prior to gel electrophoresis. To begin gel electrophoresis, you will mix your samples with a loading buffer. Loading buffer contains both dye, as a visual indicator while loading and running the sample, and glycerol, to increase the density of the samples. Increasing sample density promotes sinking to the bottom of the wells during loading, preventing the otherwise light samples from quickly diffusing out of the wells during loading.

  • Quick Tip: Consider how these samples may be presented in a future figure for presentation or publication.

    For example, if a sample may need to be cut out of an image, it is advisable to load that sample on the end to prevent compromising the integrity of the image. We put together a full list of publication requirements from the most popular scientific journals, like nature, PLOS, and MDPI to help you out.

STEP 2: Prepare gel and buffer

Gels can be purchased already made (pre-cast) or they can be made in the lab (hand cast).  In preparing the gel, there are a number of factors to consider, including the gel composition, the percentage of the gel (this will affect pore size and thus separation resolution), the number of wells needed, and the size of those wells.

  • Quick Tip: Buying pre-cast gels for can save time and ensure consistency of results by removing the inherent variation that comes with making gels by hand in the lab.

Choose the type of running buffer you will need and prepare it ahead of time. Buffers can often be purchased, though these are often made in bulk in the lab. When you’re ready to load, remove the comb from the gel. Fill the gel box with the running buffer and place the gel into the box so that it is covered by the running buffer.

STEP 3: Load and pipette samples

Before loading the samples, decide on the ideal order of the samples on the gel. Using a pipette, carefully add samples to individual wells in the gel. Additionally, a ladder with specific size markers needs to be added to one of the wells as a reference for downstream analysis.

Samples loaded into individual wells in the gel.
Visual showing DNA samples loaded into the gel wells. To load the samples into the wells, you will use a pipette and pipette tips. Later, an electric current will be applied to pull the DNA through the gel. (Created with BioRender.com)

STEP 4: Electrophoresis (Run the gel)

Once the samples are loaded, place the lid on the gel box, plug the cords into the power supply, and run the gel with electrophoresis. The voltage and time required will need to be adjusted based on each lab’s specific experiment.

Representation showing the two main methods of gel electrophoresis: agarose (horizontal) or polyacrylamide (vertical).
Visual representation showing the two main methods of gel electrophoresis: agarose/horizontal (left) or polyacrylamide/vertical (right). Each type requires the same key components: a gel with wells for the sample, buffer, and an electric current. Once samples are loaded into the wells of the gel, the power source supplies an electric current that moves the molecules (DNA or protein) through the gel. (Created with BioRender.com)

STEP 5: Visualize and document bands

Azure chemisolo next to a hand using a mobile device to connect
A unique web browser interface allows the chemiSOLO to be controlled by phone, tablet, or PC, without the need to install any additional software.

When the steps for gel electrophoresis is complete, the resulting bands of DNA, RNA, or protein need to be visualized. For DNA gels, a DNA stain added to the gel allows visualization when placed under UV light. DNA and RNA blots require additional steps prior to visualization. Proteins can be visualized in the gel (such as with two-dimensional difference gel electrophoresis or 2D- DIGE); however, more often, they need to be transferred from the gel to a membrane for further analysis. Digital imagers, like the Azure Imagers, allow for both the visualization and documentation of results in one, swift step.

Azure Biosystems offers a range of imagers capable of imaging gels stained with Coomassie, silver stain, and more. Imagers able to image under white light (epi or trans-illumination) include the new chemiSOLOIt’s the first personal Western blot imager of its kind on the market! This personal Western blot imager is able to easily and quickly image chemiluminescent Western blots without additional software downloads.

Stained gels and blots can be imaged on both laser- and CCD-based fluorescent imaging systems using total protein stains, like AzureRed (shown below) or Azure TotalStain Q. AzureRed is a quantitative, fluorescent protein stain for total protein normalization in blots and total protein detection in gels. It is fully compatible with downstream Western blotting or mass spectrometry. Azure TotalStain Q can be used to see all proteins on the membrane.

Overlay of four channels. Western blot stained with total protein stain, AzureRed, probed for three proteins of interest without a destaining step, scanned with Azure Sapphire Biomolecular Imager
AzureRed is imaged simultaneously with three proteins of interest. The gel was loaded with dilutions of HeLa cell lysate. After transfer, the blot was stained with AzureRed and then probed for tubulin, ß-actin, and GAPDH without a destaining step. The blot was scanned with each of the four lasers of the Sapphire Biomolecular Imager. In this overlay of the four channels, total protein (AzureRed stain) is shown in gray; tubulin in red, ß-actin in blue, and GAPDH in green.

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Different types of gel electrophoresis

Most gels used for electrophoresis are made from either agarose or polyacrylamide. DNA and RNA are separated horizontally via agarose gels, while proteins are separated vertically using polyacrylamide gels.

Longer run times are required for higher degrees of separation. While using a higher voltage can reduce the run time, if the voltage is too high, the gel can start to melt and create fuzzy or irregular bands. Gel composition affects how much voltage can be applied; a higher voltage may cause a low percentage gel to melt, where a higher percentage gel could withstand the higher voltage.

Check out these troubleshooting resources for gel electrophoresis and SDS-PAGE

SOURCES

  1. Electrophoresis. (2022, December 8). National Human Genome Research Institute. Retrieved December 15, 2022, from https://www.genome.gov/genetics-glossary/Electrophoresis
  2. DNA Gel Electrophoresis Equipment. (2019, September 11). LabXchange. Retrieved December 15, 2022, from https://www.labxchange.org/library/pathway/lx-pathway:33b08759-5d13-4128-8867-68428a8d1081/items/lx-pb:33b08759-5d13-4128-8867-68428a8d1081:html:ca030dca?source=%2Flibrary%2Fclusters%2Flx-cluster%3Aabe

Azure Imagers used to better understand the inhibitory mechanism of gut-derived colibactin production

Categories
Fluorescence imaging Publication Spotlight

The gut microbiota is made up of the trillions of microorganisms that colonize the human gut and have a significant impact on human health through their secreted products. These microbes can be commensal or pathogenic, and some have been connected to the development of colorectal cancer. Colibactin is a common genotoxin produced in the gut by bacteria, such as E. coli, and colibactin-mediated DNA damage appears to play a role in colorectal cancer development. The Azure Sapphire and the Azure 300 imagers helped identify a small molecule inhibitor that prevents bacterial genotoxin production in a recent study by Volpe et. al1 at Harvard University.

Since the release of this publication, the Azure Sapphire has been succeeded by the new Azure Sapphire FL, which was designed to be the flexible choice in bringing precise quantitation of nucleic acids and proteins. Learn more about the new Sapphire.

Sapphire Helps Assess Inhibitor’s Specificity

Colibactin is a known genotoxic bacterial product produced by a non-ribosomal peptide synthetase called polyketide synthase and encoded by the gene pks. This pks gene is carried by many Escherichia coli strains (pks+ E. coli).

To determine the inhibitors’ mechanism of action, the researchers initially examined the structure of ClbP and found the inhibitors mimicked some intermediates in the hydrolysis of precolibactin.

The researchers tested the effectiveness of Inhibitor 3, the most potent of the four inhibitors, against pks+ E. coli. Using metabolomics, this inhibitor was observed to be able to block the colibactin biosynthesis while only minimally disrupting other metabolic functions.

Volpe et. al used an activity-based protein profile (ABPP) to find additional targets of their four inhibitors. In this gel-based assay, small molecules that bind to a target protein are detected by the target protein’s decreased ability to bind to a nonspecific fluorophosphonate (FP) probe compound. Using the Azure Sapphire to image the gels, no measurable differences were found (Image 4). This demonstrates a lack of additional inhibitor targets and their specificity to ClbP.

Western blots from Azure Sapphire used by Volpe et al
Image 4 from Volpe et. al. (2022) examining the efficiency of their ClbP inhibitors. The Azure Sapphire Biomolecular Imager was used in Figure 4C to examine secondary targets of the inhibitors via a ABPP assay.

Designed for flexible choice in detection chemistry and samples

The new Sapphire FL brings precise quantitation of nucleic acids and proteins. As the first system on the market of its kind to allow user-interchangeable filter modules, the Sapphire FL offers a broad range of excitation and emission wavelengths.
Scientist changing optical modules on the new Azure Sapphire FL
Pick the modules that support your research. Changing the optical modules on the new Sapphire FL is simple and easy. The unique mechanism makes selecting lasers to match your dyes finally possible. Easily swap lasers, filters, and/or entire optical modules in under two minutes to suit the needs of your experiment.

Examining the Impact of Inhibitor 3 on Genotoxic Effects of Colibactin using the Azure 300

The researchers assessed whether Inhibitor 3 could inhibit the genotoxic effects of colibactin on human cells. They exposed HeLa cells to a pks+ strain of E. coli, and treated the cells with Inhibitor 3. The results indicated Inhibitor 3 is able to inhibit the genotoxic effects of colibactin in human cells, as determined by the number of cells experiencing cell-cycle arrest post treatment. It suppressed the DNA alkylating activity caused by colibactin comparable to what is observed with a genetic deletion of clbP.

Volpe et al also looked at the effects on the colibactin-induced DNA damage response. FANCD2 is a protein known to be monoubiquinated (FANCD2-Ub) in response to stalled replication forks. Previous studies show when cells are missing FANCD2, there is an increased sensitivity to colibactin3. When HeLa cells exposed to pks+ E. coli were treated with Inhibitor 3, the ubiquitination of FANCD2 was prevented, which was observed using Western blot imaged on the Azure 300.

Western blot from Volpe et al imaged with Azure 300 imager
Figure 5C of a Western blot imaged using an Azure 300 from the abstract of Volpe et. al. This represents the basic dynamic between E. coli, colibactin, and DNA damage, as well as where the created ClbP inhibitors function.

This same effect on the DNA damage response was not seen in the presence of other DNA damaging agent. This indicates Inhibitor 3 is specific to the colibactin biosynthetic pathway and not merely the DNA damage response; it ultimately inhibits the genotoxicity caused by colibactin.

Come see the light! Say goodbye to the darkroom

The Azure 300 offers the simplicity, speed and sensitivity of film detection, with better resolution and more quantitative results. The Azure 300 replaces a darkroom with film, while providing accurate and fast chemiluminescent detection, as well as the sensitivity, dynamic range, and linearity needed for quantitative blot analysis.
Azure 300 chemiluminescent Western blot imager

Designing and Selecting Potential Inhibitors

Considering the proposed structure of colibactin and knowing colibactin-activating peptidase ClbP is involved in its biosynthesis, the team designed and characterized a series of inhibitors that target ClbP2.

ClbP recognizes a motif not commonly found in substrates other than colibactin. so choosing it as a target reduces the change of secondary, off-target effects from its inhibition. The researchers found four inhibitors with potential using enzymatic activity assays.

Genotoxic Bacterial Products and the Gut Microbiome

Colibactin-mediated DNA damage appears to play a role in colorectal cancer development. Additionally, colorectal cancer patients are more frequently reported to have pks+ E.coli and mouse models show an increased tumor load when colonized with pks+ E. coli. While there is a strong correlation between colibactin and colorectal cancer, markers of colibactin mutations have been observed in normal patient samples as well. This indicates the exact timing and duration of exposure to colibactin is likely an important factor influencing colorectal cancer risk and is still poorly understood.

It would be ideal to study the effects of colibactin in the natural gut environment; however, the complexity of such an environment would make it difficult to pinpoint colibactin-specific effects. Genetic manipulations of colibactin could affect the expression or functions of other genes, including structural components, and could confound the results. Since timing and exposure to colibactin is likely important in colorectal cancer development, these variables need to be accounted for in a study of this relationship.

Volpe et. al. reasoned a compound that could specifically inhibit colibactin production would allow for the complex microbiota to remain intact while assessing the specific effects of colibactin itself.

Effects of the Inhibitor on a Complex Microbial Community

Considering the inhibitors would be used in the context of the gut microbiota, the researchers examined how Inhibitor 3 would affect other members of this complex microbe community and if these conditions affected the inhibitor’s effectiveness.

These results showed in a complex microbe environment, Inhibitor 3 would likely be able to maintain its efficacy and not damage other key members of the gut microbiota.

The Findings

By targeting the biosynthetic pathway, Inhibitor 3 abrogates colibactin and completely blocks the genotoxic effects, usually observed when mammalian cells are exposed to colibactin in culture. This precise control presents an opportunity to study natural products secreted in complex microbial communities and determine potential therapeutic strategies.

Are you staying up to date on the scientific discoveries of your peers? Check out our full list of publications by visiting our Publications Database to browse publications and pre-prints using Azure Imagers and reagents. If you’re interested in speaking with an expert about any of the products Azure Biosystems provides, please send fill out the form on this page.

Have you published with an Azure instrument?

We’d love to read it! Email your publication to us and we’ll send you something for sharing.

SOURCES

  1. Volpe, M.R., Velilla, J.A., Daniel-Ivad, M. et al. A small molecule inhibitor prevents gut bacterial genotoxin production. Nat Chem Biol (2022). https://doi.org/10.1038/s41589-022-01147-8
  2. Dubois, D., Baron, O., Cougnoux, A. al. ClbP Is a Prototype of a Peptidase Subgroup Involved in Biosynthesis of Nonribosomal Peptides. Journal of Biological Chemistry (2011).
  3. Bossuet-Greif, N., Vignard, J., Taieb, F., Mirey, G., Dubois, D., Petit, C., Oswald, E., & Nougayrède, J.-P. . The colibactin genotoxin generates DNA Interstrand Cross-Links in infected cells. MBio (2018). https://doi.org/10.1128/mbio.02393-17
  4.  

Azure 300 Introduces Young Scientists to Western Blotting

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Customer Spotlight Western Blotting

Customer Spotlight: Sarah Boylan, Director of The Applied Science & Engineering Program at St. Paul’s School in New Hampshire

At St. Paul’s School, a boarding school burrowed in New Hampshire, a robust and life-changing STEM program for students called The Applied Science & Engineering Program (ASEP) provides students with an opportunity to engage in real scientific research. The program’s director, Sarah Boylan, M.S., wants to ensure students have the skills needed to facilitate quality research, which means they need access to the best equipment and resources, including the Azure 300 Imager.   

The need for additional resources

When Boylan became Director of ASEP, she inherited lab equipment with the capability to do cell culture and other basic assays, like PCR. From her own experiences doing research at the Harvard Stem Cell Institute, Boylan wanted to expand the resources of the lab to broaden the research capabilities for the students. Since then, her purchases have expanded the applications ASEP students are able to run. Her initial purchases included a Nanodrop, microplate reader, and eventually, a qPCR machine. The students were able to visualize basic DNA gels, but their previous instruments were not able to image Western blots.

Chemiluminescent Western blot imaged on Azure 300 by ASEP students at St. Paul's School
Representative results generated by a student in the ASEP program using the Azure 300 Imager. Western blot reflects TcpA and Hcp Expression of V. cholerae samples grown in AKI and LB at 2hr, 4hr, and 6hr time points.

A great match with the Azure 300

Because some of the research projects included gene expression analysis, Boylan realized her students needed to have data from both qPCR and Western blots to ensure accuracy. After careful consideration, she eventually chose the Azure 300 Imager. Boylan saw the Azure 300 as the best choice for Western blot imaging at ASEP due to its affordability and ease of use. The added functionality to upgrade later on as their research capabilities expand was taken into consideration during the decision making process as well. It was clear that the Azure 300 Imager was the ideal option for ASEP.

Since acquiring the Azure 300 Imager early last year, the ASEP students have already begun implementing Western blots into their research projects. One student is examining siRNA and will use Westerns to determine the most effective concentration of siRNA to knock down a gene for cancer immunotherapy drugs. Another student is working with a cholera lab and will be using Westerns to look at two proteins involved in cholera pathogenesis.

Azure 300 imager
The Azure 300 is a multichannel, multimodal imager, with visible fluorescence, visible light, and UV excitation channels. The system is fully upgradeable to a fluorescent Western Blot Imaging System.

Paving a research path at St. Paul's

These young scientists recognize that troubleshooting is an incredibly important piece of the scientific process. When gels run incorrectly or their Western blots are blank, they must learn how to identify and rectify the issues. Boylan encourages collaboration amongst the students. If one student is learning a new application, such as Western blotting or qPCR, their peers will often observe in preparation for future experiments. 

ASEP gives students the opportunity to be independent learners and thinkers, while requiring them to take full responsibility for their learning. Boylan notes that while many people may think high school students aren’t adept enough to perform some applications, she’s seen first-hand just how capable her students can be with the right resources. Some students even opt to come in during free blocks to study new material. Boylan’s goal is to ensure her students get a feel for what real research is like, including the highs and lows that come with it. Scientific research is not easy, and ASEP is designed to prepare them for that reality. Students are taught that failure is part of the process. Perseverance, commitment, and dedication will serve them in life and in their careers. 

ASEP & student life

Students begin preparing for ASEP during their junior year by securing a summer research externship at a university or company in a field of interest of their choice. After they have secured a position, they may apply to ASEP. The program is very selective and only accepts about 12 students each year. During the program, students spend the spring term of their junior year preparing for their upcoming externship.  

In the fall, students return to St. Paul’s School, where they continue their research on campus and work towards a senior capstone proposal. Some students will do an extension of their summer research and choose to work with the externship lab in an ongoing collaboration. Under Boylaa’s guidance, the young researchers determine what skills they need to ensure the best chances of success. Boylan essentially acts as a Principal Investigator overseeing twelve different research projects 

The future of science

Moving forward, Boylan hopes ASEP continues to prepare the next generation of scientists by instilling in them a love of science as well as providing a life-changing opportunity to experience real-life research. She hopes to show students and other scientists that there are more options for researchers than the rigors of academia.

Ready to expand your research horizons? Check out the capabilities of the Azure 300 Imager and discover how it can enhance your research by clicking here.

To learn more about the Applied Science and Engineering Program (ASEP) at St. Paul’s Boarding School and their research, go to https://asep.sps.edu/.