Antibodies have been indispensable tools in biology since Albert Coons and his colleagues at Harvard first conjugated fluorescein isothiocyanate, a fluorescent dye, to antibodies in 1941.1 Following that foundational work, Georges Köhler and César Milstein’s development of hybridoma-based monoclonal antibodies in 1975 marked the next major leap in antibody technology, forging a clear progression from polyclonal to monoclonal to single-domain antibodies.2 In the decades that followed, fluorescent antibodies have powered discoveries in cell signaling, neuroscience, immunology, and molecular imaging. However, as imaging techniques advance toward higher resolution and quantitative precision, the limitations of conventional antibodies grow increasingly apparent. These limitations include their large size, steric hindrance, and increased background from nonspecific binding when using secondary antibodies. To meet the demands of powerful new labeling and imaging techniques, researchers are turning to single-domain antibodies (SdAbs), also known as Nanobodies®. Many consider SdAbs to be the next generation of compact, high-performance antibody reagents.
What Are Single-Domain Antibodies?
SdAbs are the smallest functional fragments of an antibody that still retain full antigen-binding capability. They’re most commonly derived from certain types of antibodies found in cartilaginous fish and camelids, a group of animals including camels, llamas, and alpacas. In the early 1990s, researchers studying dromedary camels discovered that they produce a distinct class of heavy-chain-only antibodies, which lack the light chains present in conventional antibodies (Fig. 1).2 The single antigen-binding domain from these antibodies, the VHH, was found to function independently, marking the development of what we now call SdAbs.
Explored initially for therapeutic use due to their high tissue penetration and low immunogenicity, SdAbs are now used in basic research for immunofluorescence, super-resolution microscopy, protein purification, and live-cell imaging.2-4 Their simplicity, stability, and modularity make them ideal tools for a variety of imaging and other immunodetection applications.
Figure 1. Alpacas produce three IgG subclasses: IgG1, IgG2, and IgG3. Camelid IgG1 has both heavy and light chains and is most similar to the widely used IgG antibodies from species such as mouse or rabbit. Camelid IgG2 and IgG3 are heavy-chain-only antibodies (HCAbs) lacking light chains and the CH1 domain. MiniMab™ SdAbs are the recombinant form of the VHH variable domain of these HCAbs. Created in BioRender. Criado, M. (2025) https://BioRender.com/.
Small Size, Big Advantages: How SdAbs Outperform Conventional Antibodies
SdAbs stand apart from conventional antibodies not only in size but in performance. They consist of a single variable domain with a diameter of ~2.5 nm and a molecular weight of roughly 15 kDa, about one-tenth that of a typical immunoglobulin G (IgG) antibody, and are still readily conjugated or fused to fluorophores, enzymes, or affinity tags.3 Their compact structure allows them to reach epitopes in cavities or grooves that are otherwise inaccessible and enables deeper tissue penetration for more uniform labeling in dense or fixed samples.2-4 They are also highly soluble and are more stable under conditions that would denature most antibodies, such as high temperatures, detergents, and fixation treatments, making them compatible with a wide range of workflows.4-5
Nanobodies can be efficiently produced recombinantly in prokaryotic systems such as E. coli, bypassing the need for complex eukaryotic expression systems.11 Their small, simple structure allows easy expression, purification, and large-scale production, making them valuable for biosensors, diagnostics, and therapeutics.11 High-yield, cost-effective, and easily scalable production supports both clinical applications and research, broadening access for scientists with limited resources.11 Additionally, their recombinant nature also makes them highly engineerable. The result is faster, cleaner labeling with minimal background.4
Redefining Resolution: How SdAbs Can Improve Super-Resolution Imaging and Beyond
One potentially transformative application for SdAbs is their use in super-resolution microscopy. In widefield or confocal imaging, which have a resolution of ∼250 nm at best, the small uncertainty (~20 nm) introduced by labeling targets with primary and secondary antibodies is negligible. However, as imaging resolution increases with techniques such as STED, SIM, and dSTORM, the physical size of IgG antibodies, combined with the multiple dye labels on secondary antibodies, introduces significant localization error in the target position.6 This “linkage error” can add 20-30 nm between the fluorophore and the epitope when using secondary antibodies and 10-15 nm when using primary antibodies, reducing effective resolution. SdAbs largely eliminate linkage error, therefore, significantly improving localization accuracy and resolution.7
SdAbs are also enhancing signal quality across many assays. In immunofluorescence, they penetrate tissue more efficiently and produce cleaner labeling with reduced Fc receptor-mediated background.3-5 The Fc portion of conventional antibodies, usually IgG, binds Fc receptors on cells (macrophages, microglia, dendritic cells, neutrophils, some endothelial cells, etc.), producing unwanted staining that looks like real antigen signal. Because sdAbs lack an Fc region, they largely avoid that artifact. In Western blotting, SdAbs HRP or fluorescent conjugates provide sharper bands and improved signal-to-noise.12 Their small size also benefits flow cytometry, where reduced steric hindrance enables more accurate detection of densely expressed surface markers.4
Beyond their ability to improve imaging precision, SdAbs also offer significant practical advantages in assay performance. Their rapid binding kinetics, driven by a smaller molecular size and reduced diffusion barriers, allow them to reach equilibrium faster than conventional IgGs, enabling shorter incubation times. SdAbs are also exceptionally stable, retaining their structure and antigen-binding activity under elevated temperatures, low pH, and in the presence of detergents that can denature conventional antibodies.2 In addition, their high solubility and low aggregation propensity make them easier to handle and more consistent in demanding workflows such as multiplex labeling, harsh wash conditions, or long-term storage.2 Together, these features make SdAbs robust and efficient tools for diverse imaging and analytical applications.
Additionally, SdAbs can be genetically encoded to be expressed inside cells as “intrabodies,” which bind endogenous proteins in real time, enabling live-cell visualization of dynamic processes.8 Emerging computational design and AI-assisted SdAbs discovery methods are expected to accelerate their customization for intracellular targets.9-11
Figure 2. Key advantages of single-domain antibodies (SdAbs). Due to their small size and robust nature, SdAbs exhibit minimal epitope displacement for precise localization, faster binding kinetics, enhanced stability under harsh conditions, improved signal-to-noise ratios, and superior tissue penetration, making them powerful tools for advanced imaging and detection applications. Created in BioRender. Davis, J. (2025) https://BioRender.com/.
MiniMab™ SdAbs: Next-Generation Tools for Precision Imaging
At Biotium, we’ve developedMiniMab™ SdAbs reagents to bring these advantages to everyday research. MiniMab™ SdAbs are directly conjugated to our CF® Dyes to deliver exceptional brightness, stability, and target specificity.
Figure 3. Rat eye cryosection stained with CF®568 VGLUT1 rVHH (SdAb2412.VGLUT1) MiniMab™ (orange). Nuclei are stained with NucSpot® 680/700 (magenta). Scale bar: 50 μm.
Because MiniMab™ SdAbs are much smaller than conventional antibodies, they penetrate tissue samples more efficiently and minimize linkage error in super-resolution imaging. Each MiniMab™ conjugate is validated for immunofluorescence, and a subset is validated for Western blotting and flow cytometry, offering scientists a plug-and-play solution for clear, reproducible results.
MiniMab™ SdAbs are particularly advantageous for neuroscience, where fine structural detail and low background are essential for mapping synaptic and cellular architecture. By combining the precision of SdAbs design with the brightness and photostability of CF® Dyes, MiniMab™ reagents enable researchers to visualize targets with exceptional clarity, helping bridge the gap between standard immunofluorescence and true molecular-resolution imaging. MiniMab™ SdAbs are also available labeled with our top-performing CF® Dyes for STORM.
SdAbs: Redefining What’s Possible
As the demand for high-precision imaging and quantitative protein analysis grows, SdAbs are redefining what’s possible in the lab. Their small size, exceptional stability, and direct labeling capability make them indispensable for cutting-edge applications, from super-resolution microscopy to live-cell imaging.4-7
Whether you’re mapping intricate neuronal structures, analyzing protein complexes, or simplifying your immunofluorescence workflow, SdAbs offer a smarter, more efficient path forward. Biotium’s MiniMab™ SdAbs bring this power to your bench, delivering brighter signal, lower background, and higher resolution for your research.
References
Ortiz Hidalgo C. Immunohistochemistry in historical perspective: knowing the past to understand the present. Methods Mol Biol. 2022;2422:17–31. doi:10.1007/978-1-0716-1948-3_2.
Alexander E, Leong KW. Discovery of nanobodies: a comprehensive review of their applications and potential over the past five years. J Nanobiotechnol. 2024;22:661. doi:10.1186/s12951-024-02900-y.
Muyldermans S. Nanobodies: natural single-domain antibodies. Annu Rev Biochem. 2013;82:775–797. doi:10.1146/annurev-biochem-063011-092449.
Beghein E, Gettemans J. Nanobody Technology: A Versatile Toolkit for Microscopic Imaging, Protein–Protein Interaction Analysis, and Protein Function Exploration. Frontiers in Immunology. 2017;8. doi:https://doi.org/10.3389/fimmu.2017.00771.
Helma J, Cardoso MC, Muyldermans S, Leonhardt H. Nanobodies and recombinant binders in cell biology. J Cell Biol. 2015;209(5):633–644. doi:10.1083/jcb.201409074.
Carrington G, Tomlinson D, Peckham M. Exploiting nanobodies and Affimers for super-resolution imaging in light microscopy. Mol Biol Cell. 2019;30(22):2737–2740. doi:10.1091/mbc.E18-11-0694.
Ries J, Kaplan C, Platonova E, Eghlidi H, Ewers H. A simple, versatile method for GFP-based super-resolution microscopy via nanobodies. Nat Methods. 2012;9:582–584. doi:10.1038/nmeth.1991.
Traenkle B, Rothbauer U. Under the microscope: single-domain antibodies for live-cell imaging and super-resolution microscopy. Front Immunol. 2017 Aug 24;8:1030. doi:10.3389/fimmu.2017.01030.
Fridy PC, et al. A new generation of nanobody research tools using improved mass spectrometry-based discovery methods. J Biol Chem. 2024. doi:10.5281/zenodo.10805358.
Reddy S, et al. Advancements in nanobody generation: integrating conventional, in silico, and machine learning approaches. Biotechnol Bioeng. 2024. doi:10.1002/bit.28816.
Zhu H, Ding Y. Nanobodies: from discovery to AI-driven design. Biology (Basel). 2025;14(5):547. doi:10.3390/biology14050547.
Bruce V, McNaughton B. Evaluation of nanobody conjugates and protein fusions as bioanalytical reagents. Anal Chem. 2017;89(7):3819–3823. doi:10.1021/acs.analchem.7b00470.
Nanobody is a registered trademark of Ablynx N.V.
The future of science begins in the classroom, and products that facilitate hands-on lab experiences for students are key to making that future brighter. At a time when schools are seeking to prepare students for an increasingly technical and research-driven world, the right tools can make all the difference.
This article explores how Biotium’s GelRed® and other innovative, safer reagents are transforming molecular biology education in middle and high schools. Through real-world insights from a high school science educator and a recent middle school science fair success, we’ll examine how Biotium’s products are empowering students, teachers, and the next generation of scientists.
The Need for Safe, Effective Tools in School Labs
Educators working in middle and high school science classrooms often juggle limited budgets, outdated equipment, and serious safety considerations. Traditional DNA stains like ethidium bromide pose mutagenic risks and require UV transilluminators for visualization, equipment that is costly and potentially hazardous.
In the classroom, particularly at the secondary level, educators must strike a delicate balance: maintaining scientific rigor while ensuring student safety and engagement. Hands-on learning is powerful, but only when it’s feasible and safe to implement.
Non-mutagenic and environmentally safer than ethidium bromide.
Highly sensitive, allowing for clear visualization of DNA bands.
GelGreen® is compatible with blue light imaging systems, which are safer and more affordable than UV light equipment.
Educators value these products for their simple protocols, minimal preparation, and reliable performance. The accessibility of these reagents allows teachers to focus more on facilitating meaningful lab experiences and less on troubleshooting safety or technical issues.
In the Classroom: Insights from a High School Educator
Dr. Heather York, a biology teacher at Watkinson School, integrates Biotium’s products into her curriculum with great success. With a Ph.D. in zoology and a deep interest in field biology, she brings her enthusiasm for science into her classroom lab every day with engaging projects.
Dr. Heather York earned her Ph.D. in Ecology and Evolutionary Biology with a focus on the ecology of short-tailed fruit bats.
Dr. York incorporates GelRed® Prestain Plus 6X DNA Loading Dye into her course’s studies of the ecology of Wolbachia, in which the students learn core molecular biology techniques through hands-on electrophoresis labs. For many, it’s their first time using equipment like micropipettes and gel boxes. Drawing on her field biology expertise and resources provided by the Wolbachia Project at Penn State University, Dr. York designed the project to include the collection of arthropods in the field, which students then screen for the presence of Wolbachia DNA.
“Wolbachia is a bacterium capable of infecting a broad range of arthropods,” she explained, “and it's responsible for some very interesting evolution because it affects individuals' fertility. I'm very interested in doing fieldwork with students to capture and identify arthropods and then use PCR to see if Wolbachia sequences are present. In fact, I've even gotten into tardigrade biology because tardigrades are arthropod-like and can also host Wolbachia.”
The students first isolated DNA from the samples they collected and then performed polymerase chain reaction (PCR) to amplify segments of the extracted DNA, targeting two specific genes: mitochondrial CO1 from the arthropod and 16S rRNA from Wolbachia. They then determined the presence or absence of amplified DNA in their samples via visualization on an agarose gel using GelRed®. Arthropod and Wolbachia DNA, if present, were distinguished based on the size or base pair (bp) length of the DNA molecule.
A representative tardigrade and agarose gel from Dr. York’s classroom. (A) A tardigrade collected for examination. (B) An example agarose gel from the Wolbachia project targeting a 438 bp amplicon. Samples from wells 09–15, a negative PCR control (Neg), and a 100 bp DNA ladder (Lad; New England Biolabs) were run on an agarose gel. Each sample lane contained 4 µL of PCR product mixed with 2 µL of GelRed® Prestain Plus 6X DNA Loading Dye (1:1 GelRed and Sigma Gel Loading Solution), and 4 µL of the mixture was loaded. Electrophoresis was performed from bottom to top as shown.
This integration of lab and field work allows students to engage with the full scientific process, from specimen collection to DNA analysis, and the student response has been overwhelmingly positive. Dr. York emphasizes how these tools help students connect textbook concepts to real-world applications, transforming science from something they study to something they do. Even setbacks, like gels that don’t run properly, become learning opportunities in troubleshooting and persistence.
“I’m sort of self-taught on how to run PCR,” she says. “When it doesn’t work, I tell them, ‘let’s figure out where we can tweak it.’ They see that even teachers are learning, and that’s powerful.”
Science Fair Success: Biotium in Middle School Education
In 2023, Biotium was proud to support Ruhani Singh, Jiya Kohar, and Prisha Patel, a local middle school team that placed first at the California Science and Engineering Fair.
They used GelGreen® Nucleic Acid Gel Stain to investigate the relationship between gene SLC6A4, serotonin secretion, and general mood. SLC6A4 is a gene that codes for the serotonin transporter protein (SERT) in humans, and previous medical studies have implicated changes in SERT metabolism with many different conditions, including alcoholism, clinical depression, and generalized social phobia.¹ DNA gel electrophoresis and GelGreen® were used to confirm the SLC6A4 genotype of each volunteer in their study. They found that the SLC6A4 genotype did not correlate with mental health in teenagers.
“Our project just wasn’t about some science fair we wanted to compete in; we wanted to truly help make a positive change within our own schools and community through our research,” said Prisha Patel. “Throughout our study, I learned skills that I will continue using all throughout my career as well. Our first time in a lab, our first time analyzing DNA samples, and our first time conducting a full study with human samples was an experience that taught me so much and has developed something that could help so many students when dealing with their complicated emotions.”
Project awardees from left to right: Ruhani Singh, Jiya Kohar, Prisha Patel.
This victory showcases how accessible and safe reagents can empower young scientists to explore complex topics and laboratory techniques. It’s one thing to read about DNA extraction—it’s another to run your own gel electrophoresis and analyze your own results.
Early successes like these not only inspire confidence, they set the stage for lifelong engagement in science and research. For more information on their project, check out our press release.
The Bigger Picture: Cultivating the Next Generation of Scientists
Access to safe, hands-on lab tools increases student engagement and builds crucial science literacy. Students who gain early exposure to real lab techniques are better prepared for college-level coursework and future STEM careers.
Biotium is committed to supporting educators like Dr. York by offering safer, innovative, and reliable reagents that work in real classrooms. We understand that educators don’t just need high-performance products; they need ones that are accessible, affordable, and easy to integrate into their curriculum.
From responsive technical support to a growing library of resources and kits, Biotium aims to make molecular biology more approachable for teachers and students alike.
The Future is Bright!
From the buzz of a high school classroom hunting for tardigrades to a middle school team taking home science fair gold, Biotium’s products are making advanced science more accessible than ever.
Murphy, D. L., & Moya, P. R. (2011). Human serotonin transporter gene (SLC6A4) variants: their contributions to understanding pharmacogenomic and other functional G× G and G× E differences in health and disease. Current opinion in pharmacology, 11(1), 3-10.
The development of the polymerase chain reaction (PCR) in 1983 by Kary Mullis and coworkers revolutionized molecular biology, allowing scientists to amplify DNA sequences for various applications.1 Roughly a decade later, research by Higuchi et al. led to the discovery that PCR amplification can be monitored in real time using fluorogenic nucleic acid probes, allowing for quantitative real-time PCR (qPCR).2 This technique now underpins applications like measuring levels of gene expression, rapid forms of paternity testing, scanning for mutations in target genes, and the diagnosis of many infectious diseases.
Initially, ethidium bromide (EtBr) was commonly used as a DNA-binding fluorescent dye for monitoring qPCR. Its drawbacks, including being a potential human mutagen, led to the introduction of SYBR® Green I. SYBR® Green I has brighter fluorescent signal than EtBr, but researchers soon realized that it was also a mutagen. This, along with SYBR® Green I’s propensity for background fluorescence, its inhibitory properties, and its lack of stability following multiple freeze-thaw cycles and under PCR conditions inspired Biotium scientists to develop EvaGreen® and EvaGreen® Plus Dye for qPCR. These dyes have now been helping scientists optimize their PCR, qPCR, LAMP, HRM, and ddPCR (droplet digital PCR) data for over 15 years.
Figure 1. EvaGreen® Dye binds to dsDNA via a novel “release-on-demand” mechanism.
EvaGreen® Dye uses a novel concept of DNA binding via a “release-on-demand” mechanism, which means the dye is essentially non-fluorescent by itself but becomes highly fluorescent upon binding to dsDNA (Fig. 1). The dye is constructed of two monomeric dyes linked by a flexible spacer. In the absence of DNA, the dimeric dye assumes a looped conformation that is non-fluorescent in the absence of DNA. When DNA is available, the looped conformation shifts via an equilibrium to an active form that is capable of binding to DNA and emitting fluorescence. This unique property of EvaGreen® suppresses background fluorescence of unbound dye, allowing for significantly better signal-to-noise when compared to other qPCR dyes. In this article, we’ll explore a few techniques that grew from Mullis’ and Higuchi’s groundbreaking discoveries and consider some of EvaGreen® Dye’s key advantages in these applications.
Enhancing PCR and qPCR Performance
The unique properties of EvaGreen® Dye have made it particularly useful in qPCR. Compared to SYBR® Green I, it is generally less inhibitory toward PCR and less likely to cause non-specific amplification thanks to the dye’s stronger preferential binding to dsDNA over ssDNA12. Therefore, EvaGreen® Dye can be used at a much higher dye concentration than SYBR® Green I, resulting in more robust PCR signal (Fig. 2). EvaGreen® is also compatible with multiplex PCR, as the lack of dye migration from amplicon to amplicon enables the detection of multiple PCR products by melt curves. This is thanks to EvaGreen® Dye’s higher affinity for DNA as a dimer.
Figure 2. A comparison of the raw fluorescence signal from qPCR reactions performed with two EvaGreen®-based master mixes (Forget-Me-Not™ EvaGreen® and Fast EvaGreen®) and QuantiNova® SYBR® Green I. EvaGreen® Dye is less inhibitory than SYBR® Green I, allowing for a much brighter signal.
Unveiling the Full Potential of LAMP
Loop-mediated isothermal amplification’s (LAMP) simplicity and speed make the method particularly attractive for point-of-care diagnostics in resource-limited contexts. It also shines in field settings where it’s used for rapid detection of plant pathogens or infectious disease agents like malaria, tuberculosis, and SARS-CoV-2.10, 11 Both LAMP and PCR amplify DNA, but there are a few key differences:
Temperature: The strand-displacing properties of Bst DNA polymerase used in LAMP allows for rapid amplification of a specific sequence at one temperature (60-65° C). Standard PCR requires cycling between multiple temperatures to denature the DNA, anneal primers, and extend the new strand.
Generated product: PCR generates many copies of discrete, single-sized amplicons while LAMP creates long concatemers of the same repeated sequence.
Reaction duration: PCR thermocycling protocols often take at least 1.5 hours, while LAMP reactions are complete in 30-40 minutes.
Because of SYBR® Green I’s inhibitory effect on DNA polymerization at higher concentrations, it’s mainly used as an end point reporter in LAMP.4 As summarized by Fishbach et al., EvaGreen® is more suited for real-time monitoring of LAMP reactions as it does not impact DNA polymerization and, therefore, can be present throughout the entirety of the reaction.4, 7, 8
Increasing Precision in High-Resolution Melting (HRM®) Experiments
HRM® is a post-PCR method for analyzing the melt curves of amplified DNA fragments. The method requires a DNA binding dye like EvaGreen®, and an instrument with exceptional thermal sensitivity that can distinguish melt curves between fragments with small sequence variations. The analysis can be used for DNA mapping, mutation scanning, species identification, zygosity testing, DNA fingerprinting, and more.
Despite its popularity, SYBR® Green I has been found to be unreliable for HRM® due to problems caused by dye redistribution at higher concentrations. EvaGreen® was designed to be less inhibitory in the PCR reactions, permitting the use of saturating dye concentration for maximal signal and better high-resolution DNA melt analysis with less dye redistribution. 5,6
The only DNA dye suitable for multiplex ddPCR
Droplet digital PCR (ddPCR) is a digital PCR method that is based on water-oil emulsion droplet technology. The sample is fractionated into tens of thousands of droplets, which act as individual test tubes or wells in a plate in which the PCR reaction takes place. The technique allows researchers to multiplex targets from the same sample and requires smaller sample amounts than other commercially available digital PCR systems. This results in lower sample and reagent volume requirements and reduced overall cost while maintaining the sensitivity and precision of digital PCR. Digital PCR techniques, like ddPCR, also offer the ability for absolute quantification of molecules.
TaqMan® probes have traditionally been used for this technique, but their expense and the required complexity in assay design are significant drawbacks. EvaGreen® is less likely to inhibit PCR and promote mispriming than competitor DNA-binding dyes, and it shows superior performance when used in qPCR and for high resolution melt curve analysis. This makes it a promising tool for ddPCR. McDermott et al. confirmed its effectiveness in their 2013 paper in which they identified EvaGreen® as the only DNA-binding dye validated for ddPCR in their experimental system. In this study, they also found that EvaGreen® increased resolution and allowed for the quantification of multiple target species in a single well of droplets.9
A Breakthrough in Safety
DNA-binding dyes are inherently dangerous due to their potential to cause mutations. Ohta et al. found SYBR® Green I to be even more environmentally toxic than ethidium bromide, a well-known mutagen.13 No safety data currently exists on other common PCR and HRM dyes (e.g., SYTO®9, LCGreen®, BRYT Green®, and ResoLight). However, these dyes are known for quickly permeating cells and posing a potential risk of genotoxicity. This motivated Biotium scientists to create EvaGreen®, a dye that is impenetrable to latex gloves and cell membranes, with metabolites that have little or no interaction with DNA. Testing by Biotium and independent testing services confirmed that it is noncytotoxic and nonmutagenic at concentrations well above the working concentrations used in qPCR.
References
Saiki, R. K., Scharf, S., Faloona, F., Mullis, K. B., Horn, G. T., Erlich, H. A., & Arnheim, N. (1985). Enzymatic amplification of beta-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science (New York, N.Y.), 230(4732), 1350–1354. https://doi.org/10.1126/science.2999980
Higuchi, R., Fockler, C., Dollinger, G., & Watson, R. (1993). Kinetic PCR analysis: real-time monitoring of DNA amplification reactions. Bio/technology (Nature Publishing Company), 11(9), 1026–1030. https://doi.org/10.1038/nbt0993-1026
Ishino, S., & Ishino, Y. (2014). DNA polymerases as useful reagents for biotechnology - the history of developmental research in the field. Frontiers in microbiology, 5, 465. https://doi.org/10.3389/fmicb.2014.00465
Fischbach, J., Xander, N. C., Frohme, M., & Glökler, J. F. (2015). Shining a light on LAMP assays-a comparison of LAMP visualization methods including the novel use of berberine. BioTechniques, 58(4), 189–194. https://doi.org/10.2144/000114275
Wittwer, C. T., Reed, G. H., Gundry, C. N., Vandersteen, J. G., & Pryor, R. J. (2003). High-resolution genotyping by amplicon melting analysis using LCGreen. Clinical chemistry, 49(6 Pt 1), 853–860. https://doi.org/10.1373/49.6.853
Giglio, S., Monis, P. T., & Saint, C. P. (2003). Demonstration of preferential binding of SYBR Green I to specific DNA fragments in real-time multiplex PCR. Nucleic acids research, 31(22), e136. https://doi.org/10.1093/nar/gng135
Mair, G., Vilei, E.M., Wade, A. et al.Isothermal loop-mediated amplification (lamp) for diagnosis of contagious bovine pleuro-pneumonia. BMC Vet Res 9, 108 (2013). https://doi.org/10.1186/1746-6148-9-108
Tomlinson, J. A., Barker, I., & Boonham, N. (2007). Faster, simpler, more-specific methods for improved molecular detection of Phytophthora ramorum in the field. Applied and environmental microbiology, 73(12), 4040–4047. https://doi.org/10.1128/AEM.00161-07
McDermott, G. P., Do, D., Litterst, C. M., Maar, D., Hindson, C. M., Steenblock, E. R., Legler, T. C., Jouvenot, Y., Marrs, S. H., Bemis, A., Shah, P., Wong, J., Wang, S., Sally, D., Javier, L., Dinio, T., Han, C., Brackbill, T. P., Hodges, S. P., Ling, Y., … Lowe, A. J. (2013). Multiplexed target detection using DNA-binding dye chemistry in droplet digital PCR. Analytical chemistry, 85(23), 11619–11627. https://doi.org/10.1021/ac403061n
González-González, E., Trujillo-de Santiago, G., Lara-Mayorga, I. M., Martínez-Chapa, S. O., & Alvarez, M. M. (2020). Portable and accurate diagnostics for COVID-19: Combined use of the miniPCR thermocycler and a well-plate reader for SARS-CoV-2 virus detection. PloS one, 15(8), e0237418. https://doi.org/10.1371/journal.pone.0237418
El-Tholoth, M., Bau, H. H., & Song, J. (2020). A Single and Two-Stage, Closed-Tube, Molecular Test for the 2019 Novel Coronavirus (COVID-19) at Home, Clinic, and Points of Entry. ChemRxiv: the preprint server for chemistry, 10.26434/chemrxiv.11860137.v1. https://doi.org/10.26434/chemrxiv.11860137.v1
Mao, F., Leung, W. Y., & Xin, X. (2007). Characterization of EvaGreen and the implication of its physicochemical properties for qPCR applications. BMC biotechnology, 7, 76. https://doi.org/10.1186/1472-6750-7-76
Ohta, T., Tokishita, S., & Yamagata, H. (2001). Ethidium bromide and SYBR Green I enhance the genotoxicity of UV-irradiation and chemical mutagens in E. coli. Mutation research, 492(1-2), 91–97. https://doi.org/10.1016/s1383-5718(01)00155-3
SYBR and SYTO are registered trademarks of Thermo Fisher Scientific; LCGreen is a registered trademark of Biofire; BRYT Green is a registered trademark of Promega; HRM is a registered trademark of Idaho Technologies; QuantiNova is a registered trademark of Qiagen.
Antibodies are one of the most specific tools for the detection and capture of molecular targets. The right antibody can play a key role in revealing the presence, quantity, dynamics, and even binding interactions of a target protein. However, finding the right antibody from over 2 million commercial antibodies and 300+ suppliers can be an overwhelming and confusing task.1 We’ve compiled 7 essential tips to help jump-start your search and hone in on the perfect antibody for your experiment.
1. Choosing the right target of interest
Identifying the most effective target protein, or antigen, and understanding its complexity is key in choosing the right antibody. Homology with closely related proteins can result in cross-reactivity. Proteins may have multiple names, isoforms, splice variants, and post-translational modifications. If your research depends on the antibody recognizing a specific portion of your target, note the epitope the antibody was raised against and confirm it’s within the domain of interest. Fixation methods can alter an antigen as well, and permeabilization or antigen retrieval treatments may be required to make the target or epitope accessible to the antibody. However, both fixation and permeabilization can affect immune affinity reactions. Protein databases, like UniProt and GeneCards, are great resources for more information.
2. Primary or secondary antibodies?
With “direct” labeling, primary antibodies that bind to the antigen are directly detected. “Indirect” labeling is when labeled secondary antibodies are used to detect the primary antibody against your chosen antigen.
Created with BioRender.com
The use of directly-labeled primary antibodies requires fewer steps and reagents and is the preferred method for some applications that require multiplexing (i.e., flow cytometry), but the signal intensity is lower as only the fluorophores bound to the primary antibody are present. Depending on your method of detection and the degree of expression of the target, this may be acceptable. Some dyes, such as our CF® Dyes, can help minimize the issue of low signal-to-noise as they allow for a higher concentration of dye molecules per antibody and may produce lower background due to improved solubility and less non-specific binding. Directly-labeled primary antibodies also allow researchers to multiplex using a combination of antibodies from the same host. For more information on this, see Biotium’s Tech Tip on Combining Direct and Indirect IF Using Primary Antibodies from the Same Host.
Using secondary antibodies requires more steps and reagents, but increases sensitivity due to the signal amplification from multiple secondary antibodies binding to a single primary antibody. Secondary antibodies can also be used in Tyramide Signal Amplification (TSA) to further enhance the signal of low-abundance antigens and provide opportunity for multiplexing in microscopy. See this helpful Tech Tip to learn more about multi-color labeling using Tyramide Amplification Kits.
3. Species compatibility
Due to target variation between species, you should confirm that your antibody has been validated for your sample species. In most cases, you will want to select an antibody created in a different host species from your sample species, particularly if using secondary antibodies. Validated antibodies are usually available for widely researched model organisms. If you’re unable to find an antibody validated for your sample species, you will need to assess the antibody’s performance and specificity yourself. This is worthwhile as antibodies are often raised against relatively preserved domains and may recognize your target even if the sequence homology of the epitope is as low as 75%.2
The different types of immunoglobulins (IgG, IgM, IgY, etc.) may also impact cross-reactivity and multiplexing. For example, secondary antibodies that react with IgG (H+L) will react with epitopes on both heavy and light chains, so they will react with other isotypes of primary antibody or different subtypes of IgG. Secondary antibodies that specify a specific isotype for their reactivity (e.g., IgG2a) are cross-adsorbed against other isotypes for specific binding.3 If you seek signal strength and broadness of detection over specificity, consider polyclonal antibodies, which are a mixture of antibodies that recognize different epitopes on a given target. If your experiment requires a higher degree of specificity and consistency, you may want to consider monoclonal antibodies.
4. Formulation and degree of purification
Antibodies are available in various formulations and degrees of purification. Antibodies may be supplied as pure IgG in PBS but may also come with stabilizers such as BSA, gelatin, glycerol, or amino acids. Crude preparations of antibody in serum, ascites fluid, or supernatant may contain unwanted immunoglobulins or other components that may need to be removed to avoid skewing your results. This is especially important if you plan to label your own antibody with a fluorescent dye or other type of label, as additives like amino acids, glycerol, or Tris can affect labeling efficiency.
5. Antibody and application validation
Selecting an antibody that has been independently validated can help you get successful results. Suppliers often list validated applications online or in the product documents. Never assume that antibodies validated for one application, such as western blotting, will work well in another, such as flow cytometry, as the state of the antigen may change with the technique (e.g., the antigen will typically be denatured in western blots while the native form is usually present for microscopy or flow cytometry). Finding published examples of an antibody being used in an experiment similar to yours is one of the best predictors of success. PubMed and CiteAb are great resources for determining which antibodies other researchers are using. It is also worthwhile to look for antibodies validated by the seller. Biotium offers a selection of monoclonal Biotium Choice Antibodies that have been carefully curated and extensively validated in-house for flow cytometry.
6. Multiplexing
Multiplexing allows for simultaneous detection of multiple markers in a single sample. Careful consideration of species and immunoglobulin classes is essential to minimize cross-reactivity and background. Highly cross-adsorbed antibodies work best, and most researchers prefer to use directly-conjugated primary antibodies, as some experiments can require up to dozens of different antibodies. If using fluorescently labeled antibodies, remember to choose conjugates that don’t have overlapping emission spectra!
Rat intestine cryosection blocked with TrueBlack® IF Background Suppressor (Permeabilizing) and stained with CF®647 Mix-n-Stain labeled anti-Pan cytokeratin (magenta) and CF®488A Mix-n-Stain® labeled anti-Histone H1 (green), diluted in TrueBlack® IF Blocking Buffer (Permeabilizing).
7. Supplier-provided resources
Most experiments require troubleshooting and optimization. Choosing a supplier with knowledgeable technical support and helpful resources will save valuable time and reagents. Refer to these materials to determine how to best reconstitute, store, and aliquot your antibody, and take note of any special considerations listed. These documents typically provide suggestions for optimal concentrations for different applications as well. In addition to product information documents, Biotium offers an antibody selection tool and a variety of tech tips and general protocols for antibody-based detection. Our Technical Support Scientists are always happy to help with experimental troubleshooting once your antibodies have been put to use, as well.
There are reportedly more than a billion tissue samples archived in hospitals and tissue banks around the world, representing a wealth of rich clinical data.1 Most of these samples are formalin-fixed and paraffin-embedded (FFPE). This preservation technique, developed over 100 years ago, remains the gold standard for diagnostics in pathology. FFPE tissue samples are regularly used to diagnose and study disease processes in clinical settings and FFPE processing is common practice for tissue preservation in many non-clinical research laboratories.2 FFPE processing is advantageous for many reasons: it allows for the preservation of morphology, provides convenient long-term storage and sectioning, and reduces the risks associated with infectious agents. Lab equipment for preparing biopsies for pathologists to read by scientists. Selective focus. trairut noppakaew/Shutterstock.com
With the modern rise of next-generation sequencing (NGS) platforms, scientists are now looking to apply these powerful molecular technologies to the vast pool of existing FFPE samples. The following are a handful of key ways in which NGS technologies can be applied to FFPE tissue:
Analysis of Clinical Samples: FFPE tissue samples are routinely collected in clinical settings and are often associated with extensive clinical and pathological data. By accessing the genetic material of these samples, researchers can perform retrospective studies to identify genomic alterations, discover and validate biomarkers, and pinpoint potential therapeutic targets. This allows for a better understanding of diseases, prognosis, and treatment options.
Longitudinal Studies: FFPE samples are frequently collected over long periods of time, allowing for the analysis of samples from different time points within the same patient. This longitudinal analysis can provide insights into disease progression, treatment response, and the evolution of genomic alterations over time.
Historical Archives: FFPE samples often come from archival collections of patient specimens, some of which may date back several decades. Accessing the genetic material in these samples enables retrospective studies that can shed light on the genetic basis of diseases and their historical trends. This can be particularly valuable in studying diseases with long latency periods.
Unfortunately, the fixation process also leads to extracted nucleic acids with poor quality, degradation, fragmentation, and deamination of cytosine bases, complicating transcriptomic-level investigations.2 “Formalin tends to crosslink the tissue to such an extent that it is hard to get fragments of DNA that are longer than, maybe, a hundred or two hundred base pairs,” says David Rimm, a pathologist at Yale University, who was interviewed by Nature in 2007 on this topic.1 The race is now on to generate methods for analyzing the degraded and modified biomolecules obtained from FFPE samples for molecular analysis.
Opening the Genetic Time Capsule
Historically, it has been prohibitively difficult or impossible to isolate quality nucleic acids from FFPE tissue suitable for downstream analysis, but more recent advances in DNA and RNA purification have enabled researchers to gather crucial genetic and molecular information from these samples. Common extraction techniques require de-waxing, proteolytic digestion, and purification before genetic-molecular analysis (see Figure 1).3
Figure 1. Procedures used to extract nucleic acids from formalin-fixed paraffin-embedded tissues.
Paraffin removal is often done with xylene or other organic solvents, but there are alternate methods such as direct melting with microwaves, using mineral oil and incubation at 90°C, or using specialized extraction reagents.3 Most researchers agree that the de-waxing step does not have significant effects on downstream nucleic acid analysis, though the toxicity of the deparaffinization reagent is worth considering.3
The extraction of nucleic acids from FFPE tissues is the step that most impacts sample quality, because the reagents, incubation time, and incubation temperature involved in the extraction process have all been shown to significantly affect the quality of the DNA or RNA. Traditionally, researchers relied on high temperatures to remove crosslinks and adducts, but this is only partially effective and leads to additional fragmentation of labile nucleic acids such as RNA and denaturation of double-stranded DNA. Recently, new chemistry-based techniques for the reversal of crosslinking have been developed. The RNAstorm™ 2.0 FFPE RNA Extraction Kit and the DNAstorm™ 2.0 FFPE DNA Extraction Kit utilize proprietary CAT5™ catalytic technology to reverse crosslinking (Figure 2). Chemical de-crosslinking greatly accelerates the removal of formaldehyde damage and allows the use of milder conditions, resulting in markedly improved recovery of amplifiable nucleic acids.4
Figure 2. Workflow of DNAstorm™ 2.0 and RNAstorm™ 2.0 extraction from FFPE tissues.
Compared to other methods, these kits greatly enhance the recovery of amplifiable nucleic acids suitable for a variety of applications, including NGS, qPCR, microarray, or other gene expression analysis.
Decoding the Genomic Cache with NGS
Sanger sequencing, the first method for sequencing DNA, was first described in 1977 and involved sequencing a single base at a time. A little under 30 years later, modern NGS was developed and greatly reduced the time and cost of genome sequencing. This second technological leap was based on parallel sequencing of millions of relatively short DNA fragments and required reconstruction of the short reads when sequencing long stretches of DNA. Common sequencing platforms from manufacturers like Ion Torrent, Illumina, and GeneReader utilize this technology. While revolutionary, second-generation sequencing can be challenging, especially in the presence of structural variations and low-complexity regions. This was the impetus for third-generation sequencers (e.g., Pacific Biosciences and Oxford Nanopore Technology instruments), which can read lengths of more than 10 kb and are able to directly target unfragmented DNA molecules in real-time. Unfortunately, due to the fragmented and relatively poor quality of nucleic acids extracted from FFPE samples, long-read sequencing platforms are typically not compatible with data gleaned from FFPE tissue.5 Therefore, there remains the need for more effective methods for extracting genetic material extraction from FFPE tissue as well as downstream sequencing.
From Tissue Banks to Genomic Discoveries
Optimizing the use of FFPE tissue with NGS promises great rewards in the form of access to a vast wealth of clinical and non-clinical data which can be used to study cellular processes in tissue. While challenges remain in extracting high-quality nucleic acids from FFPE tissue, recent advances in extraction methods offer hope for successful NGS analysis. Unlocking the genomic cache stored in FFPE tissue has the potential to revolutionize our understanding of diseases and pave the way for personalized medicine and scientific breakthroughs.
References
Blow, N. “Tissue Issues.” Nature vol. 448, no. 7156, 1 Aug. 2007. doi: 10.1038/448959a.
Cancer is a leading cause of death worldwide, with a reported 1.7 million new cancer cases within the United States in 2019 alone. When cancer is suspected, a tissue biopsy is often the first step toward a diagnosis. This procedure, in which cells are collected for examination, remains the gold standard for cancer diagnosis. A tissue biopsy allows healthcare providers to determine if cancer is present, shows what type of cancer it is, and gives clues about the patient’s prognosis. Unfortunately, it’s often invasive, risky, costly, and painful. These factors make repeated biopsies difficult and an impractical method for tracking tumors over time.
In the search for complementary or alternative methods, researchers have found promise in minimally invasive liquid biopsies. In this approach, bodily fluids like blood or urine are collected, and their components, including circulating tumor cells (CTCs), extracellular vesicles (EVs), and cell-free DNA (cfDNA), are analyzed for biomarkers of disease. Exosomes, a subset of EVs, have been identified as a particularly promising component in liquid biopsies.1-4
Overview of exosomes as a new target for liquid biopsy. Figure from Yu et al. 2022.1 Reproduced under the Creative Commons license.
Why Exosomes?
These tiny EV’s were originally thought to be only vehicles for the disposal of cellular waste products, but exosomes are now being recognized for their important role in intercellular communication. Growing evidence suggests this is done through a variety of functional biomolecules, including nucleic acids, proteins, and lipids that exosomes transport. Now, researchers and clinicians are exploring whether the information carried by exosomes from their cells of origin might be useful for diagnosing and monitoring diseases. Tumor cell-derived exosomes, for example, have been shown to have immunomodulatory functions and play a role in signaling, direction, and invasion of metastatic cancer cells.1, 4-5
In addition to the informative cargo they transport, exosomes also display an array of other properties that make them well suited for disease diagnosis and monitoring. The following properties of exosomes make them attractive liquid biopsy candidates:
Exosomes exist in almost all body fluids, and their membrane-bound structure protects their contents from degradation.
They are secreted by living cells, including cancer cells, and contain information about these parent cells thought to be more representative than cfDNA, which is secreted only during cellular necrosis or apoptosis.
Identifying exosomes is clear and simple relative to other liquid biopsy components due to their distinctive protein components and their size and shape when imaged via electron microscopy.
Despite the promise exosomal analysis holds for cancer diagnostic applications, additional research is needed before it becomes an accepted clinical procedure. As Lynn Sorbara, Ph.D., of NCI’s Division of Cancer Prevention, put it: “I think the major stumbling block for moving these liquid biopsy tests forward is there is not enough clinical verification and validation to know and feel comfortable that what we’re detecting with them is clinically meaningful.”1, 3-4
Publication frequencies of studies investigating different contents of exosomes in liquid biopsies for disease diagnosis over 10 years based on PubMed searches in January 2020. The number of studies in this new and fast moving field continues to grow. Figure from Zhou et al. 2020.4 Reproduced under the Creative Commons license.
The Current State of Research on Exosomes for Liquid Biopsy
Researchers have been hard at work over the past decade to validate the use of exosomes for liquid biopsies and mitigate the challenges associated with this technique. One of the main hurdles is effective, high-purity isolation due to their small size and heterogeneity in biofluids. For example, cancerous exosomes represent only a small percentage of exosomes in body fluid samples, which increases the requirements for ultrasensitive and specific detection. To solve these challenges, some researchers are developing methods to purify sub-populations of exosomes using immuno-capture with tissue-specific or cancer-specific antibodies.1, 4
Both traditional and advanced technologies are used to image or separate exosomes from various body fluids and detect their cargoes. The current push to categorize tissue-specific exosome surface proteins and components has allowed for better labeling and separating of exosomes, as well as exosome subsets, with antibodies. Though effective, antibodies validated for EV detection via flow cytometry and western blot are limited, and many more must be validated for the growing number of exosome and exosome subset-specific markers. Fluorescent dye-based detection is another common, more traditional technique for detecting EVs and exosomes, though it often requires tedious troubleshooting. One reason for this is that lipophilic fluorescent dyes used to stain exosome membranes typically have high levels of aggregation. These aggregates can be mistaken for exosomes by flow cytometers. Specialized dyes for exosomes are now being developed to mitigate this issue. Researchers have also developed several effective fluorescence-based techniques such as immunosorbent assays, the single molecule array (SiMoa) platform, and aptamer-based detection methods in recent years that build on more traditional detection methods. Advanced techniques like density gradient ultracentrifugation, size exclusion chromatography (SEC), immune-capture with magnetic beads, and acoustic-based isolation techniques have also been developed and hold promise.1
Cancer-derived exosomes are enriched in differentially expressed proteins and nucleic acids, which may act as biomarkers for the early diagnosis, stage classification, and prognosis prediction of different cancers. Figure from Yu et al. 2022.1 Reproduced under the Creative Commons license.
The Future of Exosomes in Cancer Detection and Treatment
Early screening and accurate diagnosis are known to be the primary factors in reducing mortality and increasing the recovery rate for patients with tumors or precancerous lesions. Exosomes hold great potential as possible diagnostic and prognostic biomarkers and have been investigated in a variety of cancers. In addition, exosomes are involved in metastasis, and tumor immune suppression, suggesting that they might one day play a role in disease treatment as well. In fact, some clinical trials have already begun to explore the efficacy of using exosomes as therapeutics. 1, 3
References
Yu, D., Li, Y., Wang, M., Gu, J., Xu, W., Cai, H., Fang, X., & Zhang, X. Exosomes as a new frontier of cancer liquid biopsy. Molecular Cancer21 (2022). https://doi.org/10.1186/s12943-022-01509-9
Zhou, B., Xu, K., Zheng, X., Chen, T., Wang, J., Song, Y., Shao, Y., Zheng, S. Application of exosomes as liquid biopsy in clinical diagnosis. Signal Transduction and Targeted Therapy 5, 1–14 (2020). https://doi.org/10.1038/s41392-020-00258-9
Liu, Y., Shi, K., Chen, Y., Wu, X., Chen, Z., Cao, K., Tao, Y., Chen, X., Liao, J., Zhou, J. Exosomes and their role in cancer progression. Frontiers in Oncology11 (2021). https://doi.org/10.3389/fonc.2021.639159
The Rise of the Lab-on-a-Chip
With the advent of microchips, computers that once spanned whole rooms now fit into the palm of our hand. Cutting-edge science aims to scale down research experiments in a similar manner using microtechnology. Lab-on-a-chip (LOC) technology was born from this goal in 1979 and has evolved and grown in popularity significantly since then.1
LOC is a device that scales single or multiple lab processes down to mm-scale integrated circuits, or chips, to achieve automation and high-throughput screening.2 The development of these devices requires expertise in microfluidics, the study of the physical behavior of minute volumes of fluids. Manipulating reagents at this level allows scientists to exploit microscale effects like rapid heating and mixing, significantly reduce waste, and minimize exposure to dangerous chemicals.1 More recently, this technology has evolved to include micro-engineered single or multi-cell culture platforms. These devices mimic the in vivo environment and have become effective models of living cells and organs.1-2 Consequently, the concept of LOC has gained attention as a powerful tool for biological and biomedical research.3
A lung-on-a-chip microdevice. Image courtesy of Wyss Institute/Design Museum.
From Single Synapses to Multiple Organs
Three-dimensional (3D) cell culture technologies are still often used for drug development as these paradigms allow for more biologically relevant cell networks and high-throughput screening.4 However, there are many functional aspects of cells and organs that 3D culture models cannot replicate. These limitations include a lack of tissue-to-tissue interfaces, spatiotemporal gradients of chemicals, and mechanically active components. The application of LOC technology breaks through these barriers.5
Even tricky problems, such as mimicking physically separated biological structures, can now be tackled with LOC. Yamamoto et al. showed this in their use of microdevices to create precisely compartmentalized co-cultures of motor neurons and skeletal muscle fibers to form viable neuromuscular junctions. They showed that these cells are functionally identical to their in vivo counterparts in the presence of an excitatory neurotransmitter, inhibitory chemicals, and mechanical force. These cells can be labeled with fluorescent dyes and imaged on-chip with minimal adjustments to standard protocols, as shown by the use of lipophilic carbocyanine dyes, like CellBrite® Green, to stain motor neuron axons in this system.6
As scientists gained the ability to generate increasingly complex systems on chips, they began creating organoids, leading to the birth of the organ-on-a-chip (OOC) subfield.3 With these stem cell-derived, self-organizing miniature organs, researchers seek to replicate the critical structural and functional characteristics of their in vivo counterparts. Functional OOC could provide cost effective, high-throughput disease model systems as an alternative for drug testing in animals or human tissue.7
Bauer et al. demonstrated the potential for complex OOC by generating human pancreatic islets and liver spheroids on-chip. This two-organ-chip model, complete with a working feedback loop between the liver spheroids and the insulin-secreting islet microtissues, gave the researchers the ability to study pancreatic islet–liver cross-talk based on insulin and glucose regulation. The robustness and reproducibility was validated by conducting experiments in two independent laboratories, making it a promising type 2 diabetes model.8
An illustration of the two-organ-chip used by Bauer, et al. with media circuits, respective culture compartments, and micropump valves highlighted in red. The panel on the right shows the pancreatic islet microtissues and liver spheroids used in the experiment immunostained for insulin (pseudocolored red) and glucagon (CF®594, pseudocolored green). Credit: Bauer, et al. https://doi.org/10.1038/s41598-017-14815-w reproduced under the Creative Commons license.
The Future of LOC Technology
While LOC and OOC have laid the groundwork for many potential methods of revolutionizing medicine and scientific research, most current end goals still lie on the horizon. Multi-organ chips are currently at the cutting edge of this field. To fully reflect the complexity, functionality, and integrity of an organ, one must account for the array of physiological pathways and inter-tissue interactions that exist in vivo.1 With multi-organ chips, like the liver and pancreatic circuit demonstrated by Bauer et al., researchers seek to simulate physically interconnected multi-organ systems.8
Experts hope that multi-organ chips will eventually lead to the creation of patient-on-a-chip technology. This precise approach would entail creating a complex, individualized multi-organ chip model allowing clinicians to develop treatment plans tailored to each patient’s unique biology. It would also create an attractive alternative to the use of animal models. While many limiting factors still exist, including the expense of the components, the need for universal cell culture media, and the challenge of managing and interpreting data from such a complex system, the potential reward of revolutionizing our approach to personalized medicine and laboratory research keep scientists pushing forward.1
Azizipour, N., Avazpour, R., Rosenzweig, D. H., Sawan, M., & Ajji, A. (2020). Evolution of Biochip Technology: A Review from Lab-on-a-Chip to Organ-on-a-Chip. Micromachines, 11(6), 599. https://doi.org/10.3390/mi11060599
Langhans S. A. (2018). Three-Dimensional in VitroCell Culture Models in Drug Discovery and Drug Repositioning. Frontiers in Pharmacology, 9, 6. https://doi.org/10.3389/fphar.2018.00006
Huh, D., Hamilton, G. A., & Ingber, D. E. (2011). From 3D cell culture to organs-on-chips. Trends in Cell Biology, 21(12), 745–754. https://doi.org/10.1016/j.tcb.2011.09.005
Yamamoto, K., Yamaoka, N., Imaizumi, Y., Nagashima, T., Furutani, T., Ito, T., Okada, Y., Honda, H., & Shimizu, K. (2021). Development of a human neuromuscular tissue-on-a-chip model on a 24-well-plate-format compartmentalized microfluidic device [10.1039/D1LC00048A]. Lab on a Chip, 21(10), 1897-1907. https://doi.org/10.1039/D1LC00048A
Bauer, S., Wennberg Huldt, C., Kanebratt, K. P., Durieux, I., Gunne, D., Andersson, S., Ewart, L., Haynes, W. G., Maschmeyer, I., Winter, A., Ämmälä, C., Marx, U., & Andersson, T. B. (2018). Publisher Correction: Functional coupling of human pancreatic islets and liver spheroids on-a-chip: Towards a novel human ex vivo type 2 diabetes model. Scientific Reports, 8(1), 1672. https://doi.org/10.1038/s41598-017-14815-w
Go Long to See Deeper
Imaging in visible light hits a wall when it comes to imaging in tissue for two main reasons: visible light is absorbed and scattered by tissue and tissue autofluorescence occurs chiefly in the visible spectrum. Both factors limit the usefulness of visible light for looking deep in tissue. Moving into the far red and near-infrared (NIR) region of the spectrum allows light to penetrate deeper into tissue with less absorption, scattering, and autofluorescence, improving signal-to-noise and depth of imaging. NIR spans from 650-2000 nm and contains two spectral “windows” for biologic imaging where water absorption is minimal: NIR-I ranges from 650-900 and NIR-II ranges from 1000-1300, overlapping with short-wave IR (SWIR), which spans 1000-2000 nm. NIR and SWIR imaging are not without drawbacks: spatial resolution decreases with longer wavelengths of light and reduced but present scattering can still degrade image quality. Fortunately, advances in imaging technology including adaptive optics and deep learning can mitigate these limitations.1–5
Clinical Applications of NIR Imaging
Conventional medical scans like ultrasound, magnetic resonance imaging (MRI), computed tomography (CT) or positron emission tomography (PET) scans lack cellular resolution; thus, identifying tumor locations and margins in practice remains a large challenge with these technologies. On the other hand, optical imaging using NIR fluorescent dyes can visualize individual cells. While some NIR dyes preferentially stain tumors over untransformed tissue through mechanisms that aren’t clearly understood, targeted NIR probes—NIR dyes conjugated to an antibody or ligand—can be used to visualize and define tumor margins for surgical excision.3–7
CF®750 for in vivo small animal imaging. Tumors in mice were imaged using an IVIS® imaging system (Perkin Elmer) 24 hours (left), 48 hours (center), and 96 hours (right) after IV injection of CF®750 Avastin® conjugate. Image courtesy of Caliper Life Sciences.
Indocyanine green (ICG), an FDA-approved dye, is useful for imaging vasculature in several organs. ICG is used in practice for coronary surgeries, liver laparoscopy, visualizing infant cerebral blood flow, and is being explored for lymph node mapping in oncology.8,9
Indocyanine green labeling of coronary arteries in a rat heart. Credit: Alander, et al. https://doi.org/10.1155/2012/940585 reproduced under the Creative Commons license.
Dyes for NIR
A plethora of probes for NIR imaging have been developed, including organic dyes such as ICG and Biotium’s long wavelength CF® Dyes; nanoparticles such as quantum dots, nanodiamonds, and carbon nanotubes; fluorescent proteins; and bioluminescent protein-substrate complexes. Most NIR organic dyes emit in the NIR-I range, with few emitting in NIR-II and SWIR, which provide even better signal to noise in biological tissues. Most NIR-II emitters are nanoparticles that are poorly biocompatible, containing cytotoxic materials such as heavy metals. Small organic dyes such as near-IR CF® Dyes can penetrate into tissues easily and have a linear response to illumination; the NIR spectral region offers low tissue autofluorescence, producing high sensitivity.1,3,5,7,9
Imaging in NIR-I (left) compared to imaging in SWIR using ICG, showing markedly higher contrast in SWIR despite lower ICG emission. Credit: Carr, et al. https://doi.org/10.1073/pnas.1718917115 reproduced under the Creative Commons license.
Future Developments
While NIR imaging provides unmatched in vivo imaging capability, there are some remaining limitations: high-resolution imaging at depth still has some scattering, which blurs the image despite light being able to penetrate. This is improved by imaging in NIR-II and SWIR; however, most organic dyes emit in NIR-I. Advances in deep learning and generative adversarial networks (GANs) have made it possible to translate NIR-I images to NIR-II quality with sufficient model training.1
Deep learning models can approximate a NIR-IIb (1500-1700 nm) image, with its improved signal-to-noise, from a NIR-IIa image (900-1300 nm, left). “Generated” shows the AI-created image, while “expt” shows an actual NIR-IIb image for comparison. Scale bar 5 mm. Credit: Ma, et al. https://doi.org/10.1073/pnas.2021446118 reproduced under the Creative Commons license.
Another developing technology that can improve image quality is adaptive optics. Adaptive optics were originally developed for astronomy to counter distortions by the atmosphere, but the principle can be used in biologic imaging to counteract the image produced by scattering in the tissue, even permitting super-resolution imaging at depth.2
For clinical use, several new NIR dyes have been undergoing clinical trials for use in human surgery and photodynamic therapies, largely in oncology. Biocompatible materials will lead the way in NIR dye development, both in and out of the clinic. The advantages of NIR in basic biological research are also gaining traction as volumetric microscopic imaging techniques such as light sheet become more common. Imaging in NIR-II and SWIR is also expected to become more common, as the InGaAs (indium gallium arsenide) sensors required to detect long wavelength photons become more commonplace.3,4,7,10
Understanding microbes and their ecology is crucial for modern medicine and functioning society. Microbes cause disease, but are also vital components of a healthy gut and skin microbiome. Microbes fix nitrogen for plants, produce oxygen in marine environments, facilitate rusting and infrastructure breakdown, and come with us to space. By better understanding microbial diversity and their interactions with hosts and each other, we can reduce infections and food-borne illness, improve wastewater treatment and irrigation, and better human health (Emerson et al., 2017).
Culture viability—growing microbes in controlled media and counting colonies—has been the standard method for identifying and quantifying living microbes, but culture growth takes time. Worse still, certain organisms cannot be cultured with current technology. Cultures are ideal when looking for known, culturable species of interest, but these viable but non-culturable species limits their utility for ecological and complex microbiome studies. Culture-free methods eliminate these limitations but require molecular measures of viability.
Which are the dead ones?
With a few exceptions, the cell membrane breaks down and becomes permeable in dying and dead cells. Propidium iodide (PI)—a fluorescent dye commonly used in microscopy and flow cytometry—enters cells with compromised membranes and intercalates into the DNA, activating fluorescence for optical detection. PI, however, can show “false labeling” under certain conditions, such as with adherent biofilms or with extracellular debris, overestimating the population of dead cells (Rosenberg et al., 2019). PI performance is also species and growth phase-dependent: one study found that up to 10% of PI-labeled Gram-negative Sphingomonas species and 60% Gram-positive M. fredriksbergense were actually viable (Shi et al., 2007). It is unclear if PI false labeling involves cell entry or if fluorescence is extracellular; if the latter, then non-optical methods could circumvent problematic labeling.
PCR by itself cannot distinguish between DNA from living or dead cells. In 2006, Biotium invented and remains the primary supplier of propidium monoazide (PMA) (Madrid, 2018; Nocker et al., 2006). Like PI, PMA is excluded from living cells but labels DNA in dead or dying cells (Emerson et al., 2017; Nocker et al., 2006). With light activation, PMA bonds covalently to DNA via its azide functional group, rendering DNA less soluble and a poor template for amplification (Emerson et al., 2017; Nocker et al., 2006). Viability PCR (v-PCR) with PMA “blocks” extracellular and dead-cell DNA from amplification in many downstream PCR applications (Emerson et al., 2017).
Left: Standard PCR amplifies all sources of DNA in a sample, including those from living cells (blue), dead cells (red), or extracellular sources (green). All of these species are amplified. Right: Addition of PMA labels extracellular DNA and DNA from dead cells (red) while leaving DNA in living cells unlabeled (blue). PMA labeled DNA is not amplified, only unlabeled DNA is amplified. Credit: Emerson et al. https://doi.org/10.1186/s40168-017-0285-3 reproduced under the Creative Commons license.
A Clinical Research Tool
High throughput sequencing and microbial profiling with v-PCR can enhance our understanding of microbial communities and their evolution. Lung infections are a significant cause of complications and mortality in cystic fibrosis (CF) patients (Lyczak et al., 2002). PMA-treated v-PCR and 16S-rRNA gene sequencing of sputum samples from 30 CF patients identified microbes—including potential pathogens—that were missed without PMA treatment (Rogers et al., 2013).
Validating Household Cleaners
At home, our plumbing hosts biofilms that can breed pathogens. Optimizing benzalkonium chloride formulations to kill biofilms isolated from household kitchen drains was made possible with v-PCR (Forbes et al., 2017): simple aqueous solutions had little effect on the drain biofilms, but a complex formulation with additives typical for household cleaners reduced DNA recovery from PMA-treated biofilms, indicating microbial lethality. v-PCR allows us to counter threats to health by better measuring and understanding microbial communities in health and disease.
Surveilling Risks for Food-borne Illness
Waste runoff and contaminated irrigation are sources of foodborne illness worldwide. Beyond bacteria, v-PCR has been used in viability studies of viruses and eukaryotes to improve sanitary practices in agriculture and environmental management (Alonso et al., 2014; López‐Gálvez et al., 2018; Madrid, 2018; Prevost et al., 2016; Randazzo et al., 2016). v-PCR has detected pathogenic protozoan viability (Alonso et al., 2014) and can distinguish between infectious and non-infectious norovirus in water samples (Prevost et al., 2016; Randazzo et al., 2016). No country is immune to foodborne illness from many pathogens, and the ability to evaluate and improve sanitary measures at scale through viability testing remains as crucial as ever.
A Microbial Space Odyssey
As we reach for the stars and plan interplanetary missions, we need to understand the microbial environments that form in space to protect astronauts and future spacefarers. Clean rooms on Earth are often used to simulate space conditions; samples taken from the International Space Station (ISS) and cleanrooms at the Jet Propulsion Lab revealed substantial differences in their microbiomes (Checinska et al., 2015). Airborne and settled debris in the ISS had only 1.7% and 2.7% viable population, respectively, and the living constituents were enriched in Actinobacteria—human skin commensals. Being desiccation-resistant, actinobacteria were especially dominant in the ISS HEPA filtration units. Because of the vanishingly small proportion of surviving microbes, these key differences between space and Earthbound environments would not have been possible without PMA to block material contributions from non-living sources.
Future Directions
v-PCR is a powerful technique that has grown immensely since the late-2000s (Madrid, 2018); its versatility shows strength in viability and diversity studies with myriad applications. The strengths of PCR in identifying bacterial, protozoan, viral, and even archaeal constituents have been limited only by selectivity for live cells; PMA entered the scene as a more selective replacement for ethidium monoazide (Nocker et al., 2006). Like PI, PMA’s performance varies by species and may suffer when used in complex, multispecies environments (Wang et al., 2021). PMAxx™, Biotium’s next-generation v-PCR dye, has improved selectivity and dead cell permeability compared to PMA and will lead the way in coming v-PCR applications (López‐Gálvez et al., 2018; Madrid, 2018; Randazzo et al., 2016)!
Emerson, J. B., Adams, R. I., Román, C. M. B., Brooks, B., Coil, D. A., Dahlhausen, K., Ganz, H. H., Hartmann, E. M., Hsu, T., Justice, N. B., Paulino-Lima, I. G., Luongo, J. C., Lymperopoulou, D. S., Gomez-Silvan, C., Rothschild-Mancinelli, B., Balk, M., Huttenhower, C., Nocker, A., Vaishampayan, P., & Rothschild, L. J. (2017). Schrödinger’s microbes: Tools for distinguishing the living from the dead in microbial ecosystems. Microbiome, 5(1), 86. https://doi.org/10.1186/s40168-017-0285-3
Lyczak, J. B., Cannon, C. L., & Pier, G. B. (2002). Lung Infections Associated with Cystic Fibrosis.Clinical Microbiology Reviews, 15(2), 194–222. https://doi.org/10.1128/CMR.15.2.194-222.2002
Rogers, G. B., Cuthbertson, L., Hoffman, L. R., Wing, P. A., Pope, C., Hooftman, D. A. P., Lilley, A. K., Oliver, A., Carroll, M. P., Bruce, K. D., & van der Gast, C. J. (2013). Reducing bias in bacterial community analysis of lower respiratory infections.The ISME Journal, 7(4), 697–706. https://doi.org/10.1038/ismej.2012.145
As I pondered the title of this article, my thoughts immediately went back to a request by my PhD advisor to look at the effects on cell viability by an untested drug we suspect inhibits mitochondrial protein import. Cell viability you say? Where to begin…. The myriad of pathways, mechanisms, assays, and associated readouts to measure “viability” quickly made me realize just how comprehensive the term was. For those of you wondering how to get started with cell viability, let’s explore the many tools and metrics to help provide you with some direction.
Are there changes in metabolism or cellular activity?
Assays that measure cellular activity offer the most traditional readouts for viability (often directly called cell viability assays). These assays generally rely on determining relative levels of key metabolic co-factors or ATP. They are also commonly called cell proliferation assays, because they can be used to measure the relative number of live cells in different wells or samples.
Reductive potential
Reductive potential is closely associated with important cellular redox cofactors, particularly nicotinamide adenine dinucleotides (NAD+/NADH and NADP+/NADPH). These co-factors drive cellular metabolism, signaling, and gene regulation. As a result, the reductive potential of a cell sample is commonly used as a marker for overall cell viability. Assays that measure reductive potential will use a molecule that produces a measurable output when reduced, either fluorescent or colorimetric. These molecules include MTT and XTT which produce colorimetric compounds. For these molecules, reduction by NADH produces a purple compound called formazan. A similar type of assay includes resazurin (aka alamarBlue™) which reduces to resorufin and can be measured either through colorimetric absorbance or fluorescence. While MTT assay requires cells to be lysed before measurement, XTT and resazurin assays do not, allowing multiple measurements to be made over time. The WST-1 assay is also used to measure cellular activity by direct cleavage of mitochondrial dehydrogenases to form formazan.
Reduction of MTT by NADH to form formazan product. From Riss et al. (2004). Reproduced under the Creative Commons license.
ATP levels
Another metric for cellular activity is the level of ATP, the molecule that provides the energy that drives cellular reactions, because only metabolically active cells can synthesize ATP. The most widely used method for measuring ATP is through a luminescent enzymatic assay that involves luciferase. This type of assay begins with a cell lysis step to release intracellular ATP. The ATP within the lysate is combined with luciferase and its substrate D-Luciferin. The reaction then generates luminescence which can be measured by a luminometer. Biotium’s Steady-ATP™ HTS Viability Assay Kit is a glow-type luminescence assay that functions this way and can be used for highly sensitive detection of ATP.
Cell proliferation assays
Cell viability can also be assessed by monitoring the cell proliferation rate of the culture. CFSE and ViaFluor® stains are membrane-permeant dyes that are non-fluorescent until they enter viable cells where they are hydrolyzed by cytoplasmic esterases to release fluorescent amine-reactive dyes. The dyes then covalently react with amine groups on intracellular proteins and are retained in the cell. Each subsequent cell division reduces the fluorescent signal by half, allowing each division to be monitored by flow cytometry.
Calcein-AM assays
Calcein-AM based assays are cell viability assays that require both intracellular esterase activity and intact plasma membranes (see cytotoxicity assays below). Similar to ViaFluor® Dyes, calcein-AM is non-fluorescent until it is hydrolyzed by intracellular esterases to release the green fluorescent dye calcein. However, cells require an intact plasma membrane to retain calcein fluorescence, and therefore only live cells will generate a signal. This enables calcein-AM to be used as a true endpoint viability assay and for cell killing assays.
Are my cells undergoing apoptosis?
Rather than measuring broad cellular activity, assays that are specific for apoptotic activity offer a more detailed understanding of cell viability and the mode of cell death. This category includes a variety of assays that measure various hallmarks of apoptotic cells.
Caspase activity
Measuring the activity of specific apoptotic proteases is a popular method for monitoring apoptosis. This type of assay is generally end-point and typically relies on a fluorogenic protease substrate that fluoresces after being cleaved by a specific protease. Biotium’s NucView® Caspase-3 Substrates are a similar type of assay. However, unlike other fluorogenic protease substrates, NucView® Caspase-3 Substrates release a high-affinity DNA binding dye that is retained in the nucleus. This allows real-time monitoring of both caspase-3 activity and changes to nuclear morphology.
Principle of NucView® substrate technology. The substrate is non-fluorescent and does not bind DNA. After enzyme cleavage, the high-affinity DNA dye is released to bind DNA and becomes fluorescent.
Mitochondrial potential
Mitochondria aren’t simply the “Powerhouse of the Cell” we’ve come to know. They’re an extensive organelle network vital for essential cellular processes including metabolism, calcium signaling, apoptosis, and cell proliferation. Depolarization or loss of mitochondrial membrane potential is an early event in apoptosis. Consequently, assays monitoring mitochondrial potential are used as a key metric for assessing cell viability. These types of assays rely on cationic dyes that preferentially localize in the matrix of polarized mitochondria. Examples of cationic dyes include MitoView™ 633 as well as classic dyes like TMRM, TMRE, and JC-1. These types of dyes may also be used with NucView® for dual monitoring of caspase-3 activity and membrane potential in live cells.
Internucleosomal dsDNA breaks are another key indicator of apoptosis. Apoptosis-induced DNA breaks in fixed cells and tissues can be detected using the TUNEL (terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling) assay. This assay involves a TdT enzyme that catalyzes the addition of labeled dUTP to the 3’-ends of DNA double-strand breaks. The labeled cells can then be detected with microscopy or flow cytometry.
Are my cells dead? Cytotoxicity or Membrane Integrity Assays
An alternative way to determine the number of live cells in a population is to measure the number of dead cells. Because the loss of membrane integrity is a key indicator for cell death, cytotoxicity assays (aka live/dead assays) rely on membrane-impermeant dyes or substrates to be excluded outside of living cells. The widely known trypan blue dye that’s used for cell counting functions this way. Several of Biotium’s cell viability products also rely on membrane integrity to segment live and dead cell populations. This includes Live-or-Dye™ stains as well as PI, EthD-III, TOTO®-1, TO-PRO®-1, and other dead cell specific nucleic acid stains. PMA and PMAxx™ dyes also rely on membrane integrity to determine cell viability but are mainly used for viability PCR in microorganisms.
Principle of Live-or-Dye discrimination of live and dead cells.
Other questions to ask
Real-time or end-point?
Another important consideration is determining whether your study requires only a simple yes or no answer to cell viability, or requires consistent measurements in real-time. Cell viability assays that involve cell lysis (such as Steady-ATP™ or MTT assay) or require fixation before staining (such as TUNEL) are end-point and not suitable for real-time measurements. Other cell viability assays that are non-toxic and live cell-compatible such as NucView®, Annexin V, or dead cell nucleic acid stains are suitable for either end-point or real-time analysis.
To fix or not to fix?
You may need to evaluate methods depending on whether your samples are already fixed, or if you will need to fix cells after staining for further analysis, like immunofluorescence staining. Most viability or apoptosis assays require cells to be alive during the assay. Some assays like NucView® and Live-or-Dye™ can withstand fixation and permeabilization after staining is complete. Others like dead cell nucleic acid stains, MitoView™, or calcein-AM must be assayed in live cells and cannot be fixed. On the other hand, TUNEL staining is used to detect apoptosis in cells or tissue sections that are fixed before the assay is done.
Are you planning on doing high-throughput screening (HTS)?
Careful consideration of the assay workflow is needed when screening large libraries of mutants, compounds, or other conditions for affects on cell viability. End-point assays where measurements aren’t time sensitive are generally more suitable for high-throughput screening (HTS) because the samples can be done in multi-well plates. This includes our Caspase-3 DEVD-R110 Fluorometric HTS Assay Kit and reductive potential assays. However, automated microscopy screening platforms have been developed that can accommodate high-throughput imaging and detection of cellular phenotypes (Boutros et al., 2015). For example, NucView® Caspase substrates have been validated with Essen/Sartorius Incucyte® and Molecular Devices ImageXpress® imaging platforms for high-content monitoring of apoptosis (Aftab et al., 2014; Antczak et al., 2009). In addition, assays that require multiple wash or incubation steps such as TUNEL, calcein-AM, or Live-or-Dye™ staining may require more complex high-throughput screening workflows compared to homogeneous assays that can be directly added to the cell culture medium. Examples of single-step homogeneous assays include XTT, resazurin, NucView®, MitoView™, and dead cell DNA dyes.
Do you have the right equipment?
Before planning your cell viability experiment, you should ensure that you have the proper equipment to detect the relevant outputs. Assays where the signal fills the entire cell or is released into the medium can be used with standard microplate readers for fluorescence (calcein-AM, resazurin), or absorbance (MTT, XTT). Flash-type assays where sample measurement is time sensitive, a plate reader with injectors may be required. Other assays where the signal is localized to specific compartments within the cells require a specialized imaging microplate reader, fluorescence microscope, or flow cytometer. For time-lapse imaging a real-time imaging system is also required.
Is your assay validated for your model system?
There is considerable variance in cell physiology between different model organisms and experimental conditions. Consequently, consistent and reproducible results in your model system are critical to validating any conclusions on cell viability. When designing your cell viability experiment, it is important to validate the assay procedure in your model system as well as establish the appropriate positive and negative controls.
Conclusions
Understanding what data is needed to reach a noteworthy conclusion on cell viability is crucial to selecting the right type of assay. Hopefully, this article provides some insight on the mechanisms behind the various types of cell viability assays to help make that decision. Learn more about Biotium’s full selection of cell viability and apoptosis reagents, as well as stains and probes for a variety of cellular structures.
Boutros, M., Heigwer, F., & Laufer, C. (2015, December 3). Microscopy-Based High-Content Screening. Cell, Vol. 163, pp. 1314–1325. https://doi.org/10.1016/j.cell.2015.11.007
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