Deeper understanding of coordinated DNA polymerase and helicase activities during DNA replication achieved with help from Sapphire

Imaging Publication Spotlight

In recent work published in Nature Communications, Xu et al outlined a mechanism for polymerase-helicase coupling based on structural and biochemical studies. Their work examined a yeast replisome using cryo-electron microscopy and found a dynamic mechanism in which the polymerase engages and disengages from the helicase. The authors examined the effect of helicase-polymerase coupling on the enzymatic activities of each, using assays whose output was measured by phosphor imaging on an Azure Sapphire Biomolecular Imager.

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.

Phosphor imaging is a method to detect radioactive material in applications such as NorthernSouthern, or Western blotting, or in radiolabeled tissue sections.

Learn more >

During cell division, DNA must be replicated accurately while maintaining any epigenetic information marked on the parent DNA. DNA replication is carried out by the replisome, a multi-protein complex comprised of a DNA helicase that unwinds the DNA, at least two DNA polymerases that synthesize the new DNA strands, and numerous additional protein factors. For accurate and efficient DNA replication, the helicase and DNA polymerase activities must be coordinated, but how this coordination occurs is not well understood.

How cryo-electron microscopy structures led to models for DNA translocation

In eukaryotes, the DNA helicase is a multi-subunit complex made up of a hexameric helicase shaped like a ring (referred to as MCM) combined with Cdc45 and the GINS complex, together called the CMG complex. Xu et al studied a yeast replisome consisting of the CMG complex, the leading strand polymerase Polε, and accessory factors bound to a DNA replication fork. Cryo-electron microscopy structures revealed Pole cycling on and off the MCM ring as DNA translocated through the helicase. Based on these results, the authors propose a model that may synthesize two opposing models for DNA translocation through the helicase that have been put forward based on prior sets of data. These earlier models state that either

  1. ssDNA threads around the MCM pore in a symmetrical, rotary mechanism, or
  2. that the ssDNA moves through the pore in a nonsymmetric, “inch worm” fashion.

The new experiments suggest that the MCM subunits remain in a planar ring, not an open spiral, and that DNA threads through in a rotary fashion, involving only some ATPase domains per cycle.

What affects the activity of either enzyme?

The authors also examined whether the coupling of helicase and Polε affected the activity of either enzyme (Figure 6). Previous studies have shown that Polε coupling with CMG can cause the helicase to stall at barriers on the DNA fork. Xu et al assayed the helicase activity of the reconstituted yeast replicase in vitro. Components of the assay were separated by native page and 32P-labeled products detected by phosphor imaging on the Sapphire. The results showed that, in this system, the helicase could unwind DNA in the absence of Polε. Adding Polε had little effect on helicase activity, causing only slight suppression. The authors state their structure suggests Polε stabilizes the replisome in a state such that it can stall while fork barriers are removed or repaired.

Phosphor imaging gels using Sapphire Imager from Azure Biosystems
Figure 6 from Xu et al, Synergism between CMG helicase and leading strand DNA polymerase at replication fork, showing helicase (b,c) and replication (g) assays, analyzed by imaging of gels by phosphor imaging on the Sapphire Biomolecular Imager. Licensed under CC BY 4.0.

DNA polymerase activity was also assessed in an in vitro replication assay: 32P-labeled products were separated on a 1% alkaline agarose gel and detected by phosphor imaging on the Sapphire. A mutation in Polε that disrupts coupling with MCM prevented the synthesis of the expected DNA product, suggesting that coupling is required for Polε to carry out leading strand DNA replication.

Since this paper was published, the Sapphire has been succeed by the 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 Azure 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.

What a flexible pole hinge region may allow Polε to do

In conclusion, this recent work illuminates how Polε activity is synchronized with DNA translocation by CMG. The authors find that the interaction is dynamic, with Polε cycling on and off of the MCM ring. In their proposed model, a flexible Polε hinge region may allow Pole to flip out of the way to allow for repair of DNA damage or for histone transfer to maintain epigenetic information. Future experiments can investigate the potential role of Polε-MCM coupling in epigenetic inheritance and in replication fork stalling to allow DNA repair.

More about system used in this study

The Sapphire FL is a fluorescent imaging system that includes phosphor imaging among its multiple available imaging applications. Its ability to phosphor image comes in handy especially for applications such as electrophoretic mobility shift assays (EMSA), enzyme assays, and in-vivo imaging. In addition to phosphor imaging, the Sapphire FL can carry out white light, multiplex fluorescence, bioluminescence, and chemiluminescence imaging of a wide variety of sample types, from membranes to slides to model organisms. Learn more about the Sapphire FL here.

  1. Xu Z, Feng J, Yu D, et al. Synergism between CMG helicase and leading strand DNA polymerase at replication fork. Nat Commun. 2023;14:5849.

Sapphire FL Nominated for Best New Life Sciences Product of 2023

Press Releases

Dublin, Calif. – March 7, 2024 – Azure Biosystems is thrilled to announce that the Sapphire FL Biomolecular Imager is a nominee for the SelectScience® 2024 Scientists’ Choice Awards Best New Life Sciences Product of 2023. The Scientists’ Choice Awards honor the best new laboratory products from the previous year. The Sapphire FL is one of 11 products on the short list for the prestigious award. We invite scientists and clinical professionals worldwide to vote from now until March 25 for the Sapphire FL.

The Sapphire FL is a high-resolution imaging system with resolution adjustable from 1000 micron to 5 micron, and includes customizable, user-changeable laser and filter modules that allow detection of almost any fluorophore and easy adaptation to changing imaging needs.

“We are beyond excited the Sapphire FL has been nominated for the Best Life Sciences Product of 2023,” said Lisa Isailovic, Vice President of Marketing at Azure . “We are proud of its ability to support our customer base by meeting their diverse imaging needs. It was designed to allow research-driven experimental design and put an end to research dictated or limited by laboratory instrumentation. We’re proud to say it does exactly that, and more.”

Voting for the Sapphire FL to win the SelectScience® 2024 Scientists' Choice Award for Best New Life Sciences Product is now open!. Voting closes March 25, 2024. Click here to vote now!

The Sapphire FL is the only biomolecular scanner in its class that spans the spectrum from UV to near-infrared (NIR) wavelengths, facilitating multiplex experiments. In addition to fluorescence imaging, the Sapphire FL can conduct white-light imaging and phosphor imaging, as well as chemiluminescence imaging with the optional Chemiluminescence Module.

The Sapphire FL also supports a wide range of sample types, from flat samples such as gels, blots, or slides, to samples with depth up to 4 cm including culture dishes, plants, and small model organisms. In vivo imaging of mice is possible with five built-in anesthesia ports. The focal plane is adjustable, and it is possible to collect Z-stacks and GIFs of thicker samples.

With such flexibility in imaging mode and sample type, the Sapphire FL supports a large variety of imaging applications. Azure Biosystems has published application and technical notes for using the Sapphire FL for phosphor imaging, in-cell Westerns, tumor tissue imaging, in vivo imaging of mouse tumors and more. 

Other notes describe how to take advantage of the multiplex capabilities of the Sapphire FL to conduct live/dead (AO/PI) assays on cultured cells and to increase assay efficiency with four-color multiplex detection.  There is also a searchable database of recent publications using the Sapphire FL in which new applications of the instrument can always be found.

>Learn more about the Sapphire FL and its capabilities here.

About Azure Biosystems

Azure Biosystems Inc. is an innovative life science platform company that designs, develops and markets state-of-the-art instruments, including the Azure Imaging Systems and the Azure Cielo Real-time PCR system. Our experienced team has applied their technical and market knowledge to develop industry standard-setting 2nd and 3rd generation imaging systems for life sciences.


Fluorescent protein gel assays help characterize snake venom toxins and evaluate potential therapeutics

Fluorescence imaging Protein Assays Publication Spotlight

In recent work, Bittenbinder et al developed new assays to study the proteolytic activity of snake venom and evaluate potential inhibitors. The assays take advantage of fluorescently labeled ECM proteins to characterize the proteolytic profiles of different snake venoms, assess kinetics and inhibition of proteolysis, and identify the proteins responsible for proteolysis. Fluorescent proteins were separated on SDS-PAGE and detected using the Azure 400 Imaging System (AZI400-01).

New assay to characterize proteolytic activities in venom

The authors from the Naturalis Biodiversity Center developed an assay to characterize the proteolytic activities present in the venom of eight medically relevant snakes against six ECM substrates (gelatin, collagen, elastin, fibronectin, laminin, and hyaluronic acid). For this assay, dye-quenched (DQ) fluorescently labeled ECM substrates were incubated with each snake venom. The DQ substrates are highly labeled such that their fluorescence is quenched until they are digested; once the proteins are digested, the resulting peptides fluoresce brightly. These labeled substrates are often used to study proteolysis by following the increase of fluorescence in the reaction.

In the present work, the authors ran the digestion reaction products on a gel to separate the products and visually assess which snake venoms were able to digest each ECM component. The gels were imaged on the Azure 400 using 472 nm or 524 nm excitation, depending on the fluorescent label (Figure 1D).

Fluorescently labeled substrates using Azure 400
Panel from Figure 1D from Bittenbinder et al (2023). Monitoring snake venom-induced extracellular matrix degradation and identifying proteolytically active venom toxins using fluorescently labeled substrates. Licensed under CC BY 4.0. The figure shows the ability of eight snake venom samples to degrade fluo-hyaluronic acid. Activity across the different types of snake venoms ranges from very high (D. polylepis, DePo) to very low (N. naja, NaNa).

The assay provides a powerful way to scan multiple venoms for activity against multiple ECM targets. By stopping the reactions at various times, the authors were able to follow the kinetics of substrate degradation. In addition, the assay allowed them to study whether known protease inhibitors could block the degradation of ECM substrates, which could be applied to identifying or evaluating novel compounds for inhibitory activity.

A new fluo-zymography assay

A second type of assay, a new fluo-zymography assay (Figure 5), was developed to identify the proteins in the venom responsible for degrading each substrate. In conventional zymography assays, gelatin or collagen was included in SDS-PAGE gels. Venom proteins were run in the gels, then incubated in activity buffer overnight.

Figure 5 from Bittenbinder et al (2023). Monitoring snake venom-induced extracellular matrix degradation and identifying proteolytically active venom toxins using fluorescently labeled substrates. Licensed under CC BY 4.0. Figure 5 shows a fluo-zymography assay developed by the authors. Eight snake venom samples were separated on an SDS-PAGE gel containing DQ-gelatin (fluorescently labeled and quenched gelatin). After overnight incubation in activity buffer, fluorescent bands reveal the location of proteins that have proteolytic activity against gelatin. The authors cut out these bands and analyzed by LC-MS/MS to identify the toxin protein in each band.

Gels were stained with Coomassie Brilliant Blue. Proteins that degraded the gelatin or collagen appeared as clear bands in a dark background. The authors’ new fluo-zymography assay included fluorescently labeled (quenched) gelatin or collagen in the gel matrix. Bands representing proteins that degraded gelatin or collagen are brightly fluorescent (see second figure). Gels were again imaged using the Azure 400.

The fluo-zymography assay has an advantage over conventional zymography in requiring a lower substrate (gelatin or collagen) concentration, which could conserve substrate or make the assay possible when substrate amounts are limited.

What this research means for the future of potential therapeutics

The authors conclude that these new assays allow visualization and comparison of the proteolytic activity of snake venoms. The fluo-zymography assay offers a new, sensitive way, in combination with proteomics, to identify proteins in venom with proteolytic activity. The new assays could help identify and characterize both the toxins responsible for snakebite morbidity as well as new therapeutics to treat snake bite toxicity.

Background on the impact of snake bites

Snake bites kill as many as 137,880 people per year worldwide. About three times more people experience permanent disability from non-mortal bites1. Snake bite morbidity and mortality are attributable to the toxins in snake venom, most of which are proteins. These snake venom toxins cause paralysis, kidney failure, and fatal hemorrhage, as well as tissue damage, leading to permanent disability and amputation. In fact, three of the four major classes of snake venom, including snake venom serine proteases, cause tissue damage. Targets for toxin proteases include the proteins that make up the extracellular matrix (ECM). The ECM provides structural support to tissue and is important for cell viability; degrading the ECM leads to tissue damage and bleeding.

Used in this study: the Azure 400 Imager

Azure 400 Visible fluorescent imaging system
The Azure 400 is capable of three-channel visible fluorescence detection, which enables sensitive multiplex detection of Western blots, fluorescent biomolecules and Cy2/Cy3/Cy5 or similar fluorochromes. This fluorescent imager allows users to simultaneously image and quantify up to three different targets.

The Azure 400 is a fluorescent imager that provides high-resolution imaging is ideal for publication purposes, as well as higher pixel well cap for higher dynamic range.

It is a flexible fluorescent imager that enables three-color fluorescent detection for dyes in the visible range. With the Azure 400, you can simultaneously image and quantify up to three different targets. This fluorescent imager is capable of three-channel visible fluorescence detection, and enables sensitive multiplex detection of Western blots, fluorescent biomolecules and Cy2/Cy3/Cy5 or similar fluorochromes.

More research done with the Azure 400

Ready to learn more about how easy Western blotting is by using an Azure Imager?

Set up a free virtual demo with the Azure Imaging Systems! We'd love to meet with you and your lab.
Two scientists looking at multiplex fluorescent Western blot on Azure 600 Western blot imager
Revolutionizing the way you Western blot! Azure Imagers are high performance Western blot imaging systems capable of NIR fluorescence, visible fluorescence, and chemiluminescence.


  1. Snakebite envenoming. The World Health Organization website. Published September 12, 2023. Accessed February 5, 2024.
  2. Bittenbinder MA, Bergkamp ND, Slagboom J, et al. Monitoring snake venom-induced extracellular matrix degradation and identifying proteolytically active venom toxins using fluorescently labeled substrates. Biology. 2023;12(6):765.

Assessing the response of Asian seabass (Lates calcarifer) to vaccination against Streptococcus iniae using ELISA

Immunoassay Publication Spotlight

Vaccination is an important tool to protect farmed animals against contagious diseases. Though common in livestock and domestic animals, vaccination of cultured fish has been more limited1. A recent publication from Tinh et al at the Asian Institute of Technology in Thailand investigated the ability of young Asian seabass (Lates calcarifer, Bloch 1790) to mount an immune response to a vaccine to the common pathogen Streptococcus iniae2. ELISAs performed using the Ao Absorbance Microplate Reader (Catalog # AC3000) from Azure Biosystems were an integral part of the work.

ELISA stands for enzyme-linked immunosorbent assay. The ELISA assay is a research method for quantifying proteins or antigens in an unknown solution, or in medicine as a diagnostic tool.

Learn more about ELISA >

Lates calcarifer and its response to the bacterium Streptococcus iniae

The Asian seabass is a commercially significant species for aquaculture in the coastal waters of Southeast Asia. Like other fish species, this cultured fish species is subject to disease outbreaks from bacterial and viral infections. A significant pathogen of cultured Asian seabass is the bacterium Streptococcus iniae. S. iniae can infect other fish and mammals, such as humans and dolphins, causing substantial mortality rates, skin lesions, and systemic septicemia. To protect the fish (and subsequently human fish farmers) from S. iniae, vaccination is necessary.

Vaccination would ideally take place early in the fish lifespan, before exposure to the pathogen. However, much remains unknown about timing of the development of the adaptive immune system in young Asian seabass and whether young fish can respond to vaccination.B

Studying the immune response to S. iniae

In recent work, Vinh et al investigated whether young Asian seabass could mount an immune response to a vaccine against S. iniae. Fish were vaccinated at 35 or 42 days after hatching (dph) using a heat-killed S. iniae vaccine delivered through immersion immunization (the vaccine was added to the water in which the fish were kept). The immune response of the vaccinated fish was assessed by measuring fish production of IgM by ELISA, and by measuring the expression of several immune-related genes.

The ELISA experiments assessed the amount of antibodies the fish produced that targeted an S. iniae antigen. Plates were coated with antigen and then incubated with antibody extract prepared from the immunized fish at predetermined times after immunization. Six fish were assessed at zero, seven, and 14 days after immunization. Fish antibody was bound to the plate and detected using an anti-Asian seabass primary antibody and a goat-anti-mouse secondary antibody conjugated to HRP. The amount of bound secondary antibody was determined by measuring the absorbance of a chromogenic substrate (3,3′,5,5′-tetramethylbenzidine (TMB)) at 450nm on the Ao Absorbance Microplate Reader.

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 study's findings

The ELISA experiments demonstrated that immunization resulted in a significant increase in production of antibodies targeting S. iniae in the young Asian seabass. Of the fish that were 35 dph, four of six fish generated antibodies to S. iniae antigen 14 days after immunization. Of the fish that were 42 dph, two out of 6 had antibodies to S. iniae antigen seven days after immunization, and three out of six had antibodies 14 days after immunization.

The gene expression experiments similarly demonstrated that expression of relevant genes increased in the young fish.  Expression changes were detected 1 day after immunization in the 42 dph fish compared to seven days in the 35 dph fish. No assays were performed between one and seven days, so any expression changes during that period could not be determined.

The authors conclude that early vaccination of Asian seabass is feasible and 35 dph Asian seabass can acquire immunity against a bacterial pathogen. The immersion immunization approach was successful with these young fish, showing this approach is more practical when used with young fish.

Used in this study: the Ao Absorbance Microplate Reader from Azure Biosystems

The Ao Absorbance Microplate Reader used by the researchers at the Asian Institute of Technology includes an 8-position filter wheel for the versatility to measure absorbance of many common assays, including TMB (as in the recent work discussed here), Lowry assays, Bradford assays, and more. This plate reader includes a built-in shaker with speed selection, single and dual-read modes, and analysis software to make performance and interpretation of ELISAs and other 96-well plate-based assays fast and easy. Learn more about the Ao Absorbance Microplate Reader by clicking here.

  1. Cain K. Vaccines may be the biggest tool in the fish health toolbox. Aquaculture North Ameriican website. Published March 22, 2021. Accessed January 19, 2024.
  2. Vinh NT, Dong HT, Lan NGT, et al. Immunological response of 35 and 42 days old Asian seabass (Lates calcarifer, Bloch 1790) fry following immersion immunization with Streptococcus iniae heat-killed vaccine. Fish Shellfish Immunol. 2023;138:108802.

2024 Scientific Publication Requirements for Western Blots and Gels

Western Blotting

You’ve worked hard on your research, so when it’s time to submit your work for publication, don’t forget to check the publication requirements for Western blots for each journal. With over 3,000 publications worldwide featuring our products, you could say our users know a thing or two about publishing. The imagers and systems from Azure Biosystems deliver the quality journals are asking for, but image capture is only the beginning. When capturing images of gels and Western blots, it is imperative to record relevant information to ensure an image meets the requirements for journal publication.

Autoradiograph of an SDS-PAGE gel using phosphorimaging from Azure Sapphire
Figure 1. The above figure was published in the Journal of Cell Biology and shows an analysis of phosphorylation levels on wild-type and mutant Dam1 complexes. (A) Autoradiograph of an SDS-PAGE gel (8–14%). (B) Coomassie blue–stained SDS-PAGE gel (8–14%). Licensed under CC BY 4.0.

Journal-specific publication requirements for Western blots and gels:

You should be aware of specific requirements for Western blots and gels when capturing images and preparing figures. Each journal will provide general guidelines for file types, file size, resolution, and color mode which apply to all images, including gels and Western blots. A list of general guidelines for publishing Western blots and gels in popular journals, such as ElsevierNatureScience, and Wiley are outlined below:

General publication requirements for Western blots and gels

Unfortunately, publication requirements for Western blots (Figure 1) and gels are not always uniform. Some journals recommend formats other journals will not accept, so always check the guidelines of the specific journal you plan to submit before preparing your draft for review! Knowing this, you can plan your experiments so you don’t need to cut apart the image of your gel or blot and reassemble it to create a figure. Avoid cutting and cropping your images more than necessary. Make sure internal controls are included on every gel or blot. Multiplex fluorescent Western blots are an excellent way to image your control with one or more targets on the same blot.

Some journals accept a variety of image types in the submission stage, but impose strict formatting rules once the manuscript is accepted. At the submission stage, some journals simply require figures be legible and provided in a format that is compatible with a broad range of operating systems and visualization programs. Others recommend figures be in the preferred publication format throughout all stages of the submission process.

Once a paper is accepted for publication, the requirements for Western blots and gels become much more specific and vary from journal to journal. In general the following guidelines will help ensure your image is publication ready:1

  • Capture images with at least 300 dpi and at least 190 mm wide.

    You can always shrink an image if it is too large, but you cannot increase the size of an image and many journals explicitly ban "upsampling." Using the Azure 600 Western blot imager makes capturing and saving large files easy.

  • Save a raw version of the image with no manipulations, including brightness and contrast.

  • Keep a record of the settings used to capture the image (resolution, exposure time, etc.)..

  • For modified versions of the image, keep a record of all manipulations that were performed to achieve that version of the image (brightness adjustments, channels overlaid, etc.).

Two scientists working on Azure 600
The Azure 600 uses a 9.1MP camera to provide high resolution imaging perfect for publications. Change the sample to optics distance using adjustable height shelf for enhanced detection. Zoom into the area of interest with ROI imaging to reduce background.

Best practices for preparing gel and Western blot images for publication

Creating figures from images of gels and Western blots (Figure 2) often presents a challenge. Authors must balance presenting the most relevant lanes and regions of a gel or blot with providing an accurate representation of the “big picture” of the experiment. Careful attention in the experimental design phase can help simplify this process. When planning the layout of the gel, think about how the data will be presented in an eventual figure and arrange samples in a logical manner. Do not include extraneous or unrelated samples in between the samples you plan to compare.

Western blots captured using Azure c600, published in nature
Figure 2. This figure was published in nature and shows Western blots captured using Azure c600 from Pascini et al. Immunoblotting evaluating the tissue-specific expression of PAI-1 in the transgenic mosquito.

Whenever possible, comparisons should only be made between samples run on the same gel. Internal controls, housekeeping protein standards, or total protein stains should always be processed on the same gel as the experimental samples. Therefore, in the planning phase, consider ways to optimize the amount of data to be generated from running just one gel. If one gel is not possible, make sure to include a control on each gel.

A recent publication by Kroon et al examined published Western blot figures and found a majority are cropped, missing essential information about the methods, and do not supply the original images as supplementary information. This reading provides recommendations to make Western blot figures more informative and reproducible, such as minimizing cropping and including (and labeling) molecular weight markers in all images.

Two scientists smiling holding Azure pub mugs
Are you getting ready to submit your paper for publication? Congratulations! We want to recognize your hard work. If you have published using an Azure Imager, post your publication on social media using the hashtag #ImagedbyAzure and tag us, we'll send you a pub mug to show off!

Editors at the Journal of Cell Biology published an article in 2004 addressing the temptation to alter or “beautify” images and describing acceptable and unacceptable manipulations of digital images. It provided an overview of the guidelines for blot and gel images that had been published to date by a variety of journals. Adjustments applied evenly across the entire image such as adjustments to brightness, contrast, or color balance are generally acceptable; however, it is always preferable to use an image that does not require such adjustments. For example, if your bands of interest in your Western blot are faint, it is better to take a longer exposure for publication rather than choose to digitally adjust the faint image to increase perceived band strength.

quantitative western blot basics


Get a quick overview of the steps you can take to ensure your Western blots are quantitative. This free guide also includes a troubleshooting section and tear-out quantitative Western blotting checklist.

Nonlinear adjustments to an image should be avoided if you’re submitting to a journal. If they are used, carefully document the adjustments that were made. Some journals will require that these adjustments be described in the methods or figure legends. It is never acceptable to digitally alter the data in an image of a gel or blot; do not adjust contrast to hide background or faint bands. Those “nonspecific” bands may indicate your Western blotting conditions were not ideal and you need to change your blocking buffer or adjust your antibody concentrations. Alternatively, such bands maybe contain data whose importance will only become clear in the future. Maybe that “extra” band is actually an isoform of your protein, or a cleavage product.

Always save the original images used to make a figure. Some journals will request original images during the review process. Some journals, like the Nature portfolio journals in the life sciences, require original, unprocessed images of gels and Western blots used in figures be published as Supplementary Information (Figure 3).

Journals vary in how they prefer to receive figures in initial submissions. Always look up the specific requirements for the journal to which you are submitting, including required naming conventions for figure and image files. In summary, remember to capture high-resolution images (at least 300 dpi) and to carefully record your imaging settings. If you adjust an image, keep track of exactly what changes you made and always maintain a copy of the original raw image.

If you’re looking for an imager that’s reliable and provides sharp, clear images with every scan, look no further than a system from Azure Biosystems. Browse this list of available systems and choose the system that works best for your studies, like the Azure Sapphire Biomolecular Imager. The Sapphire is a next generation NIR fluorescent scanner equipped with lasers that delivers unmatched flexibility and performance for phosphor imaging, Western blots, animal imaging, in-cell Westerns, and more. Learn more about the Sapphire by clicking here.

  1. Rattan, U.K.; Kumar, S.; Kumari, R.; Bharti, M.; Hallan, V. Homeobox 27, a Homeodomain Transcription Factor, Confers Tolerances to CMV by Associating with Cucumber Mosaic Virus 2b Protein. Pathogens 2022, 11, 788. pathogens11070788

When to use Wet, Semi-Dry and Dry Transfers for Western Blots

Transfers Western Blotting

Choosing the right transfer method can determine your Western blot’s success. The first step to Western blotting is separating the proteins in a sample by size, using a denaturing process called gel electrophoresis.

How does Western blot transfer work?

In Western blotting, after the electrophoresis step, is the transfer step. The transfer step is important to Western blotting because it moves the separated proteins from the gel onto a solid support matrix using a membrane (either nitrocellulose or polyvinylidene difluoride (PVDF)). This is where the blot will form.

Cases where you would not need to transfer the gel

Sometimes, protein separation is not required, so your sample may be directly added to the membrane using an approach called “dot blotting.”

Three types of Western blot transfers

Your choice of transfer comes down to whether you need quantitative information from your blot, time and cost, or if your protein is finicky and requires you to customize transfer conditions (Table 1). So, wet or dry – which is the best transfer method for your Western blot? Let’s go over when to use each transfer method to best prepare you for your next Western blot procedure.

Table of contents

Table 1. When to know when to choose which Western blot transfer method

ScenerioWhich transfer method to use
You want to gain quantitative information from your Western blotWet transfer
Wet transfers allow you to customize the time, temperature, voltage, or buffer to best suit your protein of interest.
Saving time and reagents is equally importantSemi-dry transfer
In a semi-dry transfer, the only buffer used is the one that saturates the stack components (Figure 2).
You're short on timeDry transfer
Dry transfers typically only take around ten minutes to finish. They are also the least quantitative Western blot method.

How wet transfers for Western blotting work

If you want to gain quantitative information from your Western blot, you should do a wet transfer using a tank. Wet transfers are performed in a tank filled with transfer buffer (Figure 1). Most transfer systems, like the Azure Aqua Transfer Tank, have room for two precast or handcast gels. Tanks are routinely used in wet labs for Western blot transfers.

Azure Aqua during Western blotting experiment
The Azure Aqua Transfer Cell is used for transferring two mini gels to membranes for Western blotting experiments (wet transfer). The transfer cassettes and electrode core are colored for directionality which makes transferring easy and straightforward. You can perform a quick 1-hour transfer or you can transfer overnight at a lower voltage.

The transfer buffer used for wet transfer protocols is traditionally a Tris-glycine buffer, which contains methanol, but you may also use other buffers too. In certain cases, SDS may be added to the buffer to aid the transfer of large proteins. 

Wet transfer setup for electrophoresis
Figure 1. Wet transfer setup. A “stack” is built in which the gel is placed next to a membrane (nitrocellulose or PVDF), both of which have been pre-equilibrated in transfer buffer. Blotting papers and sponge, which have also been pre-soaked in transfer buffer, are added to the outside of the stack so the stack can be held firmly within a cassette that is suspended in the transfer buffer–filled tank. Electrodes on the cassette allow an electric current to be run through the stack so the proteins migrate from the gel to the membrane.
  • Quick Tip: The Transfer buffer from Azure Biosystems is specially is formulated to increase protein transfer and protein retention on the membrane for optimal sensitivity.


  • Wet transfers are highly customizable. The time, temperature, voltage, and buffer can be varied to suit the protein of interest and to achieve complete transfer of a broad range of proteins.
    • For example, longer transfer times may be used to allow larger proteins to fully migrate out of the gel, while shorter transfer times can prevent loss of low-molecular weight proteins that may otherwise migrate through the membrane entirely.
    • The voltage can also be reduced to slow the transfer process for an overnight run, or increased to complete the transfer in an hour or two.
  • Overall less expensive than other transfer methods
  • Can use multiple buffers to optimize transfers
  • Transfers broad molecular weight range at one time
  • Extended transfer is possible
  • Can be used for quantitative Westerns


  • One disadvantage of wet transfers is that heat is generated during the transfer. This can contribute to inconsistent transfers and to breakdown of the gel. To combat the effects of excess heat, wet transfers are often conducted in a cold room and/or with ice packs or cooling units in the tank.
  • Another disadvantage to wet transfers is they require a large volume of transfer buffer. If you regularly conduct a large number of transfers, reagent consumption can become an issue. Because of the larger volume of usage, you will also accumulate more hazardous waste. To conserve reagents, semi-dry transfer methods can be used.
  • Wet transfer can also take up to one hour. With some systems, you can also run transfers overnight at a low voltage.
  • Cooling mechanism and/or cold room space is required during transfer.

How semi-dry Western blot transfer methods work

In a semi-dry transfer, the only buffer used is the one that saturates the stack components. The membranes and gels will be placed between wet filter papers (a sandwich) that will be in direct contact with the electrodes (Figure 2). Semi-dry transfers should be your choice if saving time and reagents is your first priority.

Semi-dry transfer stack for electrophoresis
Figure 2. Semi-dry transfer setup. In a semi-dry transfer, the stack consists of gel and membrane placed between two pieces of filter paper, all equilibrated in transfer buffer. This stack is placed directly between two electrode plates.
  • Quick Tip: Transfer times cannot be extended for proteins that do not transfer completely with the standard protocol.

    The stack can dry out and the buffer capacity of the small amount of transfer buffer will be exhausted if transfer times are too long.


  • Overall easy to setup. Good for performing large numbers of blots where you are analyzing the same protein.
  • With semi-dry transfers, transfer times are reduced to about an hour, but may be as short as five  minutes with rapid semi-dry transfer protocols.
  • The hazardous waste from transfer buffer is minimized because only small volumes are used each time.


Semi-dry transfers run into difficulties at extreme ends of the protein size range. Large proteins may not transfer out of the gel quantitatively in the short transfer time available, while small proteins may transfer entirely through the membrane.

  • To compensate, you can use discontinuous buffer systems. With these systems, the two pieces of filter paper on either side of the stack are equilibrated in different buffers. For example, buffers can be chosen to help transfer difficult proteins out of the gel, and/or to improve retention of proteins in the membrane.
  • High intensity field strength may cause low molecular weight proteins to migrate through membrane.
  • Difficulty in transferring high (>120 kDa) molecular weight proteins
  • Not recommended for quantitative Western blots
  • Quick Tip: Even though discontinuous buffer systems can help improve transfer across a larger range of protein sizes, semi dry transfers are not recommended for quantitative Western blotting.

How “dry” Western blot transfer methods work

With dry transfer, the gel is placed between the preassembled stacks, that already contain the transfer membrane and proprietary buffer matrices you’ll need. Because of this convenience, if you are short on time, dry transfers are the best option.


  • Dry Western blot transfer systems do not use transfer buffer at all, which saves time.
  • Can be completed in as little as ten minutes when using newer models.


  • There is no opportunity to customize or optimize your solutions based on your protein of interest.
  • With dry transfers, preassembled stacks must be purchased ahead of time. This adds to reagent and consumable costs.

Remember: Western blot transfers are customizable!

Wet Western blot transfers are highly customizable and are recommended for quantitative Western blotting but consume a lot of reagents. Semi-dry Western blot transfers conserve time and reagents, but may not allow quantitative transfer for all proteins, especially those that are very small or very large. Dry Western blot transfers are the fastest of all and require no buffer preparation, but do not allow much room for optimization.

Looking for clear and consistent imaging results? Your search ends here

The Azure 600 uses a 9.1MP camera to provide high resolution imaging perfect for publications. Request a free demo of an Azure Imaging System, and say "Hello" to beautiful Western blots.
Two scientists working on Azure 600

In summary, the transfer method you choose will depend on several factors of your Western blotting experiment. One of the factors is how you balance the importance of speed and/or reducing reagent consumption vs needing quantitative transfers or to customize the protocol for a difficult protein. Check out the resources below if you’re still having trouble, or send us a message using the form on this page. Cheers for now!

Frequently Asked Questions about Western blot transfers

The reason for blocking the membrane before and during incubation of the primary antibody is due to the membranes being “sticky” to any protein. We “block” in order to cover up all parts of the membrane that don’t have protein on them, including any region in between lanes, etc.  When we add the antibodies, they will only bind to the protein of interest, not the blank membrane.  Blocking ensures we don’t get signal from primary antibodies sticking to the membrane. We will only get signal from primary antibodies binding to their protein of interest.

Want to try a new blocking buffer? We offer free samples!

The blocking agent may be protein or non-protein based. Some of the more commonly used blocking agents include: normal serum, Bovine serum albumin, non-fat, dry milk, Polyvinylpyrrolidone, or Tween 20. Read more about blocking agents in this application note.

There’s an application note for that! Check out our app note “Wet or Dry?” for a variety of buffer recipes.

Shop Western blotting accessories for electrophoresis

Quantitative Westerns: What is the Best Way to Normalize your Western blot?

Fluorescence imaging Multiplex Quantification Western Blotting

Far from being an “is-it-there-or-not” technique, modern digital detection instruments can make Western blotting reproducible and quantitative. By working within the linear dynamic range of your detection method and normalizing the data to control for variations in protein load and membrane transfer, you can get truly quantitative results.

But what is the best way to normalize protein levels for a Western blot? In the past, the gold standard normalization method was to use a housekeeping protein based on the assumption that the levels of these proteins are fairly consistent across experimental conditions and cell lines. However more recent studies have shown that this assumption is not always true1,2 leading to inaccurate measurements of relative protein abundance. Instead, quantitative Western blotting experts1,2 and the journals they publish in4 are recommending a new gold standard for normalization—normalizing to total protein detected in each lane, preferably by staining on the membrane.

Table of contents

Using Total Protein Stains for Normalization

With total protein normalization, instead of trying to find a protein that can represent the total amount of sample that transferred to the membrane, total protein is measured on the membrane directly. This value is then used as the denominator when normalizing.1-4 Many total protein stains used to stain gels and membranes are commercially available.1 Total protein stains provide a larger dynamic range and demonstrate lower variability and cleaner data than housekeeping proteins.1,2

Total protein normalization can also be much faster than using a housekeeping protein, especially for chemiluminescent Western blots. This is because the time it takes to stain the blot takes less time compared to stripping and reprobing. Ideally, total protein staining is conducted on the membrane, either before or after immunodetection.2 Using some stains, such as AzureRed Fluorescent Total Protein Stain, it is possible to stain the blot before immunodetection and then to image total protein simultaneously with the protein(s) of interest. For Western blots and gels, AzureRed is able to detect less than 1 ng of protein per band or spot and is non-toxic and biodegradable, for safe and simple disposal.

AzureRed showed superior correlation and a much broader dynamic range than the common housekeeping proteins.
AzureRed showed superior correlation and a much broader dynamic range than the common housekeeping proteins, such as GAPDH.

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.

AzureRed Fluorescent Total Protein Stain
Choose AzureRed Fluorescent Protein Stain for sensitive detection of total protein on 1D or 2D gels. AzureRed is as sensitive as silver stain and is compatible with downstream Western blotting, mass spectrometry, and Edman sequencing. It provides very low background and higher signal to noise than other fluorescent stains and can be imaged with UV or blue light excitation.

AzureRed allows you to stain 1D and 2D gels in less than 3 hours, with high sensitivity, low background, and no speckling. Stained gels and blots can be imaged on both the new Sapphire FL (or other laser-based systems) and the Azure Imaging Systems (or other CCD-based fluorescent imaging systems).

Advantages and disadvantages with using Housekeeping Proteins for Normalization

Housekeeping protein• Familiar and commonly used• Narrow, linear dynamic range • Abundance can vary with experimental conditions • Abundance may not be consistent between sample types • High variability • Must ensure housekeeping protein physically resolves from protein of interest on the gel
Total protein• Larger linear dynamic range • Low variability • Constant across sample types • No change with experimental conditions• Must ensure the total protein stain that's used is compatible with antibody binding and detection method

Table 1. Benefits and challenges of using a housekeeping protein vs. total protein for Western blotting

Inconsistent levels

The most significant drawback of using housekeeping proteins is their levels may not be consistent across samples and conditions.1,2 While it is possible to use a housekeeping protein for normalization, but you must first spend the time and effort to validate your choice. You may also need to examine multiple potential standards before you find one that is truly expressed at the same level across all of your samples and does not change across your experimental conditions.

High abundance

A second significant challenge associated with housekeeping proteins is their high abundance.1,3 If the housekeeping protein is present at a very high level in your sample, this limits the amount of sample you can load on the gel because you will need to keep the housekeeping protein within the linear range of detection and not saturate the signal for the housekeeping protein. This is particularly problematic if the protein of interest is not similarly highly expressed, because the two proteins will not be within the same linear range of detection.2,3

Generating primary and secondary antibodies from non-overlapping species is difficult

A third challenge to consider if you’re doing multiplex Western blots, such as comparing phosphorylated and non-phosphorylated forms of the same protein, is the complexity of generating primary and secondary antibodies from non-overlapping species.

Keep in mind it is always possible that detecting the housekeeping protein could interfere with detection of the protein of interest.1 Ideally, the housekeeping protein should be a different size than the protein of interest, so the two proteins are spatially resolved on the Western blot. This becomes increasingly difficult when an experiment examines multiple proteins of interest on the same Western blot.


>> ADDITIONAL READING: Multiplex fluorescent Western blotting

Analysis after using a total protein stain

Comparison of a traditional western blot workflow to a western blot workflow using AzureRed Total Protein Stain
Comparison of a traditional Western blot workflow to a Western blot workflow using AzureRed Total Protein Stain

The analysis workflow after image capture is essentially unchanged compared to using a housekeeping protein; the signal density for the entire lane or a large portion of the lane is used for normalization instead of the density for a single band.

Staining the membrane with a total protein stain provides an added quality control benefit, allowing verification that membrane transfer was complete and free of artifacts. With this very simple workflow, images for the protein(s) of interest and total protein are automatically aligned, avoiding the need resize and align images captured at different times.

While you’re here, check out this brochure for a complete overview of available total protein stain options. For more on how to perform accurate Western blot normalization using AzureRed Fluorescent Protein Stain, check out this application note. Cheers for now.

Additional blog posts regarding total protein:

Frequently Asked Questions

Total protein normalization (TPN) is used to quantify the abundance of the protein of interest, without having to rely on housekeeping genes. It is usually done by incubating the membrane with a total protein stain. Read more

TPN uses the entire protein content of each sample for normalization instead of relying on only a single housekeeping protein. You can see an example of total protein staining here.

AzureRed is a perfect choice for staining applications, including post-transfer staining to confirm uniform protein transfer from gel to membrane, and
staining quantitative Western blots as part of a TPN protocol. Read more

Azure offers a range of imaging systems includes several models that allow target protein detection to be multiplexed with TPN – with no need for dedicated precast gels or laborious stripping and re-probing. Instead, you simply treat your blots with TotalStain Q between protein transfer and blocking, and process them as you would normally. Read more

Shop Total Protein Stains


  1. Moritz CP. Tubulin or not tubulin: heading toward total protein staining as loading control in Western blots. Proteomics. 2017;17:1600189.
  2. Thacker JS et al. Total protein or high-abundance protein: which offers the best loading control for Western blotting? Anal Biochem. 2016;496:76-78.
  3. McDonough AA et al. Considerations when quantitating protein abundance by immunoblot. Am J Cell Physiol. 2015;308(6):C426-C433.
  4. Fosang AJ, Colbran RJ. Transparency is the key to quality. J Biol Chem. 2015;209(50):29692-29694.

Azure to Exhibit at 2023 AACR Annual Conference in Orlando, FL

Press Releases

Dublin, Calif. – April 3, 2023 ­– Azure Biosystems will be exhibiting at the 2023 Annual Meeting of the American Association for Cancer Research (AACR) in Orlando, FL at the Orange County Convention Center. This will be Azure’s first attendance at the event since 2019. The Azure team will be led by Tram Tran (Associate Marketing Manager), Vanna Sombatsaphay (Application Scientist, qPCR), and Justin Menko (Technical Product Specialist), who are excited to show meeting attendees Azure’s complete line of imaging and qPCR products. 

Azure Biosystems AACR 2023
In attendance for Azure at 2023 AACR will be Vanna Sombatsaphay (Application Scientist, qPCR), Justin Menko (Technical Product Specialist), and Tram Tran (Associate Marketing Manager).

Visit the team at booth #304 to see how easy gel imaging and densitometry can be with Azure’s brand-new personal chemiluminescence imager, the chemiSOLO. Also on display will be the new Sapphire FL Biomolecular Imager, a flexible imaging system with completely customizable and user-changeable optical modules. The Sapphire FL is compatible with nearly any imaging application, from gels and blots, to slides and small living animals, thanks to the addition of five anesthesia ports. Other flagship products from Azure that will be on display include the Cielo qPCR system, as well as the Azure Imaging Systems; all products will be available for in-person demos. The 2023 AACR Annual Meeting takes place April 14-19, 2023.

Is your AACR schedule quickly filling up? Schedule a meeting with someone from our team to make sure you don’t miss out! Schedule your meeting time here.

About Azure Biosystems

Azure Biosystems Inc. is an innovative life science platform company that designs, develops and markets state-of-the-art instruments, including the Azure Imaging Systems and the Azure Cielo Real-time PCR system. Azure Biosystems’ experienced team has developed 2nd and 3rd generation imaging systems for the life science market, thus applying their technical and market knowledge in creating innovative industry standard setting platforms.


Tram Tran
(925) 307-7127

Multicolor fluorescence imaging with Sapphire used to probe the structure of DNA recombinase

Fluorescence imaging Publication Spotlight

In a recent publication in Nature Communications, Caldwell et al used the multi-color fluorescence imaging capacity of the Azure Sapphire Biomolecular Imager to study the structure of DNA recombinase1.

The RecT family of recombinases contains over 1500 members. These enzymes bind to and catalyze the annealing of two complementary pieces of single-stranded DNA (ssDNA). No protein structures of RecT family members from bacteria or prophages have been solved. Some structures of RAD52, a potential human homolog of RecT, have been reported without DNA and with ssDNA, but not with two pieces of ssDNA which would represent an important intermediate of the annealing process.

Since the release of this interview, 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.

Solving the LiRecT Structure

Gel-shift assays imaged using fluorescent imaging on Azure Sapphire Biomolecular Imager
A portion of Figure 8a from Caldwell et al (2022). Structure of a RecT/Redβ family recombinase in complex with a duplex intermediate of DNA annealing. Gel-shift assays testing whether mutated forms of the LiRecT protein can bind to Cy3- and Cy5-labeled strands of ssDNA. For each mutant and the wild-type (WT) version of the protein, each strand was added individually (labeled in the figure as 3 for Cy3, 5 for Cy5) or sequentially (labeled 35). This portion of the figure shows that for some amino acids, double mutations interfered with DNA binding while single mutations did not. The gel was imaged on the Azure Sapphire Biomolecular Imager. Licensed under CC BY 4.0.

In this new work, Caldwell et al have solved the structure of LiRecT, a member of the RecT family from the prophage Listeria innocua, bound to a DNA duplex intermediate. The 3.4Å structure shows that multiple LiRecT subunits form a helical filament with an external groove that binds an extended and “un-wound” DNA duplex. The structure confirms that there is some structural similarity between the RecT family and the human protein RAD52. Though there is a great deal of difference between the structures, the authors identified a common core of structural similarity in the DNA-binding groove, supporting the hypothesis that the enzymes share a common underlying mechanism of protein-mediated DNA annealing.

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.

Can mutated proteins still bind to ssDNA?

To confirm the importance of specific amino acids that appeared to contact the DNA in their structure, the authors mutated 21 amino acid residues to see if changing them interfered with DNA binding. Gel-shift assays were used to see if the mutated proteins could still bind to ssDNA. RecT is known to bind weakly to individual strands of ssDNA and strongly to two complementary strands when they are added sequentially. Therefore, the authors used two complementary strands of ssDNA, one labeled with Cy3 and the other with Cy5, and conducted gel-shift assays in which the DNA strands were added individually or sequentially to the mutated enzymes. The results were imaged on the Sapphire, whose multi-color fluorescence capability allowed the researchers to detect the migration of each oligonucleotide on the same gel. The results generally supported the interactions suggested by the structure, though double mutations were often needed to completely disrupt DNA binding.

The work is an important contribution to understanding the mechanism of action of the RecT family and in what ways it resembles and differs from that of RAD52.

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.


  1. Caldwell BJ et al. Structure of a RecT/Redβ family recombinase in complex with a duplex intermediate of DNA annealing. Nat Commun. 2022;13(1):7855.

Imaging Coomassie-stained gels using NIR fluorescence and white light

Imaging SDS-PAGE

What is Coomassie?

Coomassie is a blue stain used in proteomics-related studies to detect proteins during electrophoresis, SDS-PAGE, and Bradford assaysIn Western blot analysis, Coomassie is used as a loading control and an anionic pre-antibody stain. Coomassie binds non-specifically to almost all proteins. It is also the most common method of in-gel protein detection, whose popularity can be attributed to its characteristics of being efficient, quick, and affordable. 

In this blog post, we will cover how to use white light and NIR fluorescent to image coomassie-stained gels.

How to image Coomassie stained gels using white light or using NIR fluorescence

To begin, soak the gels in the dye. To reduce background and make the bands easier to visualize, elude the extra stain with a solvent; this step is called destaining. Destaining can take as little as 10 minutes, to as long as overnight to produce bands with clear background.

Doing these steps allows for the visualization of proteins as blue bands on a clear background. Below we show a serial dilution of bovine serum albumin (BSA) run on an SDS-PAGE gel and stained with a colloidal Coomassie stain. The same gel was imaged 3 ways.

Coomassie-stained gel imaged on the Azure 300 Imager on a white light transilluminator
Figure 1. Coomassie-stained gel imaged on the Azure 300 Imager on a white light transilluminator
Coomassie-stained gel imaged on the Azure chemiSOLO
Figure 2. Coomassie-stained gel imaged on chemiSOLO using white light

The white light images were captured after placing the gel on a white light transilluminator table using white light on the Azure 300 Imager (Figure 1) and new chemiSOLO (Figure 2).

Coomassie-stained gel imaged in the 700 channel (NIR fluorescence) on the Azure Sapphire Biomolecular Imager
Figure 3. Coomassie-stained gel imaged in the 700 channel (NIR fluorescence) on the Sapphire Biomolecular Imager

The NIR fluorescence image was captured using the 700 channel (excitation wavelength 685nm) on the Sapphire Biomolecular Imager (Figure 3). On all images, the protein bands are easily visible down to the lowest amount loaded: 90 ng.

NIR fluorescence imaging of Coomassie-stained gels

About 15 years ago, it was reported that Coomassie blue-bound protein fluoresces in the near infrared3. Since that time, infrared fluorescent imaging of Coomassie stained gels has not become routine, perhaps limited by the availability of instruments able to carry out near-infrared fluorescence imaging3. Butt et al conducted a systematic study comparing the sensitivity and linear dynamic range of seven commercial Coomassie stains and seven published stain formulations when imaged using NIR fluorescence3.

The Coomassie stains were also compared to Sypro Ruby, which, though expensive, is a popular fluorescent stain due to its high sensitivity, high dynamic range, low interprotein variability, and mass spec compatibility. Butt et al found that some Coomassie stain formulations exceeded Sypro Ruby with respect to sensitivity and linear dynamic range3.

In examining 2D gels of mouse brain extract, Butt et al found more proteins were detected in gels stained with Sypro Ruby. The authors hypothesized the difference was due to attenuation of Sypro Ruby fluorescence in high-abundance spots, which may allow longer imaging to detect low abundance spots without signal saturation. This would suggest a tradeoff between sensitivity and quantitative accuracy when deciding between Sypro Ruby and Coomassie to stain a 2D gel3.

The sensitivity and linear dynamic range observed in this study suggests Coomassie may be an attractive, less expensive option for fluorescent staining of protein gels in some situations.

Scientist using pliers to place gel inside Azure chemiSOLO interface
The new chemiSOLO is used to image Coomassie-stained gels, chemiluminescent Western blots, and more.

White light imaging of Coomassie-stained gels

In a Coomassie-stained gel, the protein bands appear blue in a clear background may be examined by eye on the bench or on a light box. For record keeping, the gel may be imaged photographed with white light. The sensitivity of traditional Coomassie staining depends on the specific protein being assessed. Detection limit per protein band is about 30 to 100 ng of protein and can approach 10 ng2, 3. The sensitivity of traditional Coomassie staining is about 10 ng2.

In addition to R-250 (the original Coomassie Brilliant Blue ) and G-250 (the demethylated derivative of Coomassie Brilliant Blue ), colloidal Coomassie stains are available in which the dye is present in micelles1. Colloidal stains result in lower background than traditional Coomassie stains because the micelles are too large to enter the polyacrylamide gel matrix1. Colloidal Coomassie blue staining can provide higher sensitivity, as low as 1 ng2. depending on the formulation3,4, 5.

Why Coomassie staining is so popular for SDS-PAGE gels

Coomassie blue is one of the most commonly used dyes for staining proteins in SDS-PAGE gels. A variety of Coomassie stain formulations can be purchased commercially or mixed in the lab. The staining protocol typically involves incubating the gel in staining solution until bands are visible, followed by destaining to remove background.

The entire staining process with Coomassie can take from a few hours to overnight. Destaining reduces sensitivity and is essential with traditional Coomassie staining because high background is common. Coomassie dyes bind protein through electrostatic and hydrophobic interactions1.

Advantages of using Coomassie

Advantages of Coomassie staining include simplicity and affordability. It is also compatibile with downstream mass spectrometry analysis. 

Disadvantages of using Coomassie

Disadvantages include low sensitivity compared to other staining methods. Staining intensity can depend in part on amino acid composition of the proteins, which complicates the use of Coomassie for relative quantitation of different proteins1.

Looking for better ways to visualize your proteins? Azure Biosystems offers a range of imagers capable of imaging Coomassie-stained gels under white light (epi or trans-illumination). The new Azure chemiSOLO is able to easily and quickly image chemiluminescent Western blots without additional software downloads. It’s the first personal Western blot imager of its kind on the market! Get a quote for chemiSOLO by clicking here.

Frequently Asked Questions about coomassie

Coomassie blue stain binds proteins non-covalently so it is compatible with downstream analyses such as mass spectrometry which is frequently used to identify the protein in a spot or band from a gel. Multiple formulations of Coomassie stain are commercially available and a variety of staining techniques have been published. Read more about Visible Gel Imaging here.

When used in acidic conditions, Coomassie will bind to the hydrophobic, basic residues in proteins. Its color will change from a dull copper, red/brown hue to an intense shade of blue.

There are a number of issues that can arise during a Western blot transfer.  Many times it can be resolved by adjusting the transfer time, temperature or simply remaking the buffers. More troubleshooting tips are available in this blog post: Trouble-free Transfers.


  1. Smejkal GB. The Coomassie chronicles: past, present and future perspectives in polyacrylamide gel staining. Expert Rev Proteomics. 2004;1(4):381-387.
  2. Butt RH and Coorssen JR. Coomassie Blue as a near-infrared fluorescent stain: a systematic comparison with Sypro Ruby for in-gel protein detection. Mol Cell Proteomics. 2013;12(12):3834-3850.
  3. Butt RH and Coorssen JR. Coomassie Blue as a near-infrared fluorescent stain: a systematic comparison with Sypro Ruby for in-gel protein detection. Mol Cell Proteomics. 2013;12(12):3834-3850.
  4. Neuhoff V, Arold N, Taube D, Ehrhardt W. Improved staining of proteins in polyacrylamide gels including isoelectric focusing gels with clear background at nanogram sensitivity using Coomassie Brilliant Blue G-250 and R-250. Electrophoresis. 1988;9(6):255-262.
  5. Candiano G, Bruschi M, Musante L, et al. Blue silver: a very sensitive colloidal Coomassie G-250 staining for proteome analysis. Electrophoresis. 2004;25(9):1327-1333.