Methods for Protein and Antibody Bioconjugation to Gold

We discuss methods for protein and antibody bioconjugation to gold including theory, alternative approaches, and protocols.

Gold isn’t just used to make pretty jewelry. It is a highly versatile reagent used in a wide variety of applications. One important application of gold is for protein and antibody bioconjugation. Gold has been conjugated to a range of proteins such as antibodies, protein A, lectins, enzymes, toxins, and many others. This article discusses methods for protein and antibody bioconjugation to gold. These include…

Methods for protein and antibody bioconjugation to gold include passive non-covalent coupling, ionic coupling, coupling via thiol-maleimide chemistry, streptavidin-biotin interactions, and via amide bond formation.

We’ve also discussed how to label proteins with fluorescent probes instead of gold in our articles.

Protein and antibody bioconjugation to gold nanoparticles - TEM image
Gold nanoparticles can be coated with polymers that include functional groups. These functional groups can be used for antibody and protein bioconjugation. TEM image of gold nanoparticles (source).

Applications of Protein- and Antibody- Gold Bioconjugates

Since the early 90s, gold-protein bioconjugates have seen significant research into their applications. The technology and understanding of these bioconjugates have developed considerably in the last 30 years, opening up applications in many fields, particularly in biomedical science.

Applications of gold bioconjugates include gold staining for electron microscopy, biological assays, and biosensors. 

Gold Staining for Electron Microscopy

Gold is an excellent material for electron microscopy because of its high electron density which allows it to produce high contrast images (to learn more about gold in microscopy check out this article from Fourie et al.). Once a protein is bioconjugated to gold, the gold acts as a stain, giving much better imaging of the target protein. Electron microscopy using gold bioconjugates has been studied most extensively for the immune system in the field of immunohistochemistry. Interestingly, by staining with gold particles of different sizes, multiple objects can be labeled in a single sample. 

Gold Bioconjugate Bioassays

Gold bioconjugates are now used in a wide range of bioassays because of their excellent ability to bind to specific proteins and their widely adjustable optical and surface properties. Versatility allows gold bioconjugates to be modified to fit specific experimental conditions with high precision. Depending on the configuration of the assay, the type of gold conjugate, and the target protein, potentially thousands of proteins can be assayed in parallel and assessed for their various interactions. Gasparyan et al. describe the use of gold bioconjugates in immunoagglutination and DNA hybridization in their review here.

Related articles:

  • Immunoprecipitation can also be utilized for some bioassays as an alternative to gold conjugate based bioassays. However, it’s a much more involved and tedious technique.

Gold Biosensors

Extending from their incredible versatility in bioassays, the use of gold bioconjugates as biosensors has been explored in extensive detail. The flexible optical properties of gold nanoparticles make them excellent optical biosensors once bound to a target protein. Other gold-based biosensors use gold’s electrochemical or piezoelectric properties to create biosensors for specific proteins. Historically, even radioactive gold particles have been used in biosensing, although the use of such bioconjugates has seen a significant decline for obvious reasons.

Gold biosensors provide several advantages compared to traditional techniques such as ELISAs – they can provide constant time-series information, used without taking time points, and assays are much cheaper after the initial biosensors have been developed. Gold nanoparticle biosensors are described in detail in this review article.

Immuno-gold labelling, TEM

Antibody bioconjugation to gold allows scientists to explore biological phenomenon using TEM using gold’s optical properties. Here’s an example of immune-gold labeling. TEM image of antibodies (orange) marked with gold particles (black dots) (source).

How Does Gold Bioconjugate to a Protein?

The bioconjugation of gold to a protein occurs through highly complex mechanisms which often vary depending on the type of gold, the functionalization of the gold, the target protein, and the local chemistry. While this seems overwhelming, these complex mechanisms can be simplified into two general types.

Types of protein bioconjugation to gold can be classified into two general types: passive conjugation which involves non-covalent interactions and covalent conjugation.

Type 1. Passive Bioconjugation of a Protein to Gold

Passive conjugation is the traditional method of conjugating gold to a protein. The interaction occurs passively between the protein and the gold particle through van der Waals and ionic forces. Depending on the conditions, a protein can spontaneously conjugate with a gold particle. By varying the size of the particle, and its ratio to the protein present, the conjugate can potentially bind multiple proteins to a single gold particle or surface.

Passive conjugation is useful because it’s a quick and easy way to produce a functional protein conjugate. However, it isn’t perfect. Passive conjugates often lack long-term stability and require very specific conditions for each protein used or conjugation may not occur. Worse, as conditions change, passively conjugated proteins can detach from the gold if the conditions in the experiment change.

Type 2. Covalent Conjugation of a Protein to Gold

It’s important to have a sensitive and stable conjugate that doesn’t decouple when the experimental conditions change. By covalently reacting your protein with gold particles, a significantly more stable gold bioconjugate can be formed. This allows for more complex experiments to be performed and potentially harsher conditions to be used. 

Covalent conjugation is achieved by using functionalized gold particles. These groups cross-link the protein to the gold creating a strong bond. However, covalent conjugation is usually a more involved process requiring a higher investment of time and money. Additionally, it requires pre-functionalized gold particles that you can react with.

For other protein conjugation chemistry methods, take a look at our article.

Methods for Antibody Bioconjugation to Gold Nanoparticles

Gold nanoparticles are an important type of metallic gold that is commonly used for bioconjugation with antibodies in immune system studies. Gold nanoparticles are metallic gold structures that are roughly 1 to 100 nm in size. Their outer layer can be customized to provide the desired functionality. 

Methods for antibody bioconjugation to gold nanoparticles include passive conjugation in water and EDC/NHS covalent conjugation.

Method 1. Passive Antibody Conjugation to Gold Nanoparticles:

As mentioned above, passive conjugation is a straightforward and easy technique to conjugate proteins to gold. This applies to antibodies and gold nanoparticles too. Here’s a step-by-step example of how to conjugate antibodies to gold nanoparticles based on the nanoComposix BioReady gold nanospheres:

Step 1. Gather your Starting Materials including Gold Nanoparticles, Antibodies, Clean Sample Tubes, and DI water.

Collect the appropriate gold nanoparticles, your target antibody, and sample tubes to perform the reaction in. You’ll need access to DI water. The nanoparticles should be suspended in DI water and the antibodies should be free of additional proteins or salt additives. 

Step 2. Mix the Antibodies with the Gold Nanoparticles in DI Water. 

Aliquot your gold nanoparticles into a sample tube. Rapidly add your purified antibodies to the gold nanoparticle solution and cover the sample tube.

Step 3. Incubate the Sample while Conjugation Occurs.

The conjugation reaction is quick but not instantaneous. Allow your sample to incubate at room temperature for 30 minutes or so with gentle stirring/rotating.

Step 4. Centrifuge the Sample and Collect the Conjugate. 

Your sample needs to be centrifuged at 3500 RCF for 10 minutes. After, carefully remove the supernatant. Resuspend your conjugate in DI water.

Step 5. Store your sample at 4°C / 39.2°F

Your sample needs to be stored at low temperatures to ensure it lasts as long as possible before decoupling. However, do not freeze your conjugate as this can cause the sample to decompose. 

Method 2. Covalent Conjugation of an Antibody to Gold Nanoparticles Using an NHS Ester Reaction.

Biomedical and nanomaterial suppliers often provide ready-made kits for performing protein conjugations to gold. These typically use EDC and NHS reactive groups to active carboxyl groups on activated gold nanoparticles which then couple to the protein. Stratech offers a kit that used carboxyl-activated gold nanoparticles to couple with antibodies using EDC/NHS chemistry. Here’s a step-by-step example of that process:

Step 1. Gather your materials: Carboxyl-activated gold nanoparticles, the target antibody, the conjugation reagent (EDC/NHS), buffer solutions, and suitable clean glassware.

Again, the antibody to be conjugated must be free of contaminants such as salts or other proteins. Make sure your glassware is clean and has been rinsed with DI water. 

Step 2. Prepare the conjugation reagent. 

Prepare the conjugation reagent in a buffer solution to ensure its stability. It should be prepared fresh, right before conjugation.

Step 3. Mix your gold nanoparticles in the conjugation reagent. 

This activates the gold nanoparticles and prepares them for conjugation. The mixture needs to be gently stirred while incubating at room temperature for 30 minutes.

Step 4. Add a coupling buffer to the solution and centrifuge the mixture for 30 minutes. 

The kit comes with a buffer for the coupling step. Carefully add that to the mixture and centrifuge it for 30 minutes. 

Step 5. Add your antibody to the mixture and sonicate for 10s. 

Carefully add the desired amount of antibody to the conjugation reagent and gold nanoparticles. Sonicate in a water bath sonicator for 10s. 

Step 6. Incubate the sample again for 2 to 4 hours at room temperature. 

Gently stir the mixture while it incubates so that no particles can settle at the bottom.

Step 7. Add more buffer solution and then centrifuge the mixture for 30 mins. Remove most of the supernatant. 

To ensure the stability of your conjugate, add more buffer solution, and then centrifuge the sample again for 30 minutes. Carefully remove most, but not all, of the supernatant. 

Step 8. Add a washing buffer and store conjugate at 4°C / 39.2°F ready for use. 

Finally, add a washing buffer to the solution and store it at 4°C / 39.2°F. Don’t freeze your conjugate.

Protein Bioconjugation to Gold Nanoparticles and Surfaces

Gold bioconjugates aren’t just used in immune system studies. Beyond antibodies, gold conjugates can be added to various proteins such as protein A, lectins, enzymes, toxins, and many others.

Methods for protein bioconjugation to gold nanoparticles include covalent conjugation using NHS/EDC reactions, covalent conjugation using thiol reactions, and chemisorption. 

Methods for protein bioconjugation to gold surfaces include physical conjugation to functionalized gold surfaces, dative binding of gold conducting electrons to amino acid sulfur atoms on the protein, and ionic interactions in polar solvents.

For more information on biopolymer surface functionalization techniques, take a look at our article.

Method for Protein Bioconjugation to Gold Nanoparticles via Thiol Reactions

Many proteins contain amino acids with sulfur atoms in their structure. These sulfur atoms can be reacted with functionalized gold nanoparticles to create thiol crosslinkers. Abcam produces a gold conjugate kit which uses gold nanoparticles to react with thiol-group containing proteins:

Step 1. Add the Protein into the Diluent Reagent.

Your protein must be free of additives such as other proteins or stabilizers.

Step 2. Add a Buffer to the Mixture.

Add the buffer slowly into the mixture with gentle stirring.

Step 3. Pipette the Sample Directly onto the Gold Nanoparticles.

The mixture should be resuspended gently by withdrawing and re-dispensing the liquid twice using a pipette. 

Step 4. Incubate the Sample for 60 Minutes.

Incubating the sample for longer has no negative effect on the conjugate.

Step 5. Add the Quenching Reagent to the Sample.

Slowly add the quenching reagent to the mixture with gentle stirring. The conjugate can be used after 20 minutes. 

Step 6. Store the Sample at 4°C / 39.2°F.

Don’t freeze the sample as it may damage the conjugate.

Method for Protein Bioconjugation to Gold Surfaces by Ionic Interactions

Proteins can be bioconjugated to gold surfaces using ionic interactions. This relies on the attraction between the negatively charged gold surface and the positively charged protein. The interactions can be adjusted based on the conditions in the mixture. This requires experimentation to optimize the conditions for conjugation to the surface. Once bound to the gold surface, these conjugates can potentially be used as biosensors or assays. Here’s a step-by-step method to help optimize your conjugate production based on an article by Rayavarapu et al.:

Step 1. Create Solutions Containing the target Protein

Ensure the protein is free from additives like salts or other proteins. 

Step 2. Add the Protein to the Gold Surface. 

Add to the gold surface with gentle mixing.

Step 3. Incubate the Sample for 60 Minutes

The sample will passively conjugate by ionic interactions. 

Step 4. Centrifuge the Sample and Remove the Supernatant.

The sample should be centrifuged at 3500 RCF for 10 minutes.

Step 5. Assess the Sample.

Analyze your sample to assess how well the protein has conjugated to your gold surface.

Step 6. Adjust the Conditions and Repeat the Experiment

Adjust the conditions of your reaction. This could be the concentration of protein, the pH of the solution, the incubation time, the temperature, etc. 

Streptavidin Gold Conjugation

Streptavidin is a protein purified from Streptomyces avidinii. It has a high affinity for biotin (vitamin B7), which is one of the strongest non-covalent interactions in nature. The strong interaction between streptavidin and biotin can be used to attach biomolecules, requiring harsh conditions to break the binding. By creating a gold-streptavidin bioconjugate, researchers gain access to a powerful tool for electron microscopy and the detection of biotinylated compounds. 

There are many methods to bioconjugate streptavidin to gold. 

To bioconjugate streptavidin to gold,  use EDC/NHS reactions to create covalent bonds or passively conjugate them in pH-sensitive conditions.  

Commercially Available Gold Conjugation Kits

Gold conjugation kits can be purchased from many nanomaterials and biomedical science suppliers such as Abcam, Nanocomposix, and Sigma Aldrich.

Kit NameKit SupplierHow does it work?Price in USD
Ab154873AbcamUses covalent conjugation and is designed to survive even the most extreme conditions. Gold particles are 40nm, 20 OD575
High sensitivity gold conjugation kitNanocomposixComplete kit containing everything needed to optimize a lateral flow assay.895
NHS ester functionalized conjugation kitCytodiagnostics – Sigma AldrichGold conjugate kit with NHS ester functionalized gold nanoparticles of 40 nm. Can be used to produce a SARS-CoV-2 conjugate for COVID-19 detection. 275

Protein Labeling with Fluorescent Probes – Theory and Methods

We discuss types of fluorescent probes, how to select a fluorescent probe for protein labeling, protein labeling kits, and protocols for protein labeling using kits.

Protein labeling is an extremely useful process in many fields including biology, biotechnology, medicine, forensics, genetics, and more. In simple terms, protein labeling is the use of a ‘label’ to bind to a protein in one way or another so that it can be detected, monitored, analyzed, and even purified. Being able to label a target protein opens scientists up to a range of possibilities for interacting with a target protein and understanding the many complex processes that proteins perform in the body. While there are many different types of protein label and labeling techniques, this article focuses on protein labeling with fluorescent probes.

Protein labeling with fluorescent probes can be accomplished by linking cyanine dyes, rhodamine dyes, or fluorescein to cysteines, lysines, tyrosines, or the N-terminus of your target protein.

What Are Fluorescent Probes?

Fluorescent probes are molecules that absorb light at a specific wavelength and emit it at a specific wavelength that can be detected. Fluorescent probes are also known as fluorescent tags or fluorescent labels.

Take a look at this page from Nature about fluorescent probes. The absorbance and fluorescence of a fluorescent probe are dependent on a range of factors including its chemical structure, the solvent it is dissolved in, its binding target, and more. Fluorescent probes are mainly used in biological studies, but can also be used in other applications. This includes tracers, dyes and stains, indicators, and more.

In protein labeling, fluorescent probes are typically a reactive derivative of a fluorescent molecule known as a fluorophore. Each specific fluorophore is chosen so that it will selectively bind to the target protein in the desired location. In some uses of fluorescent probes, the fluorophore isn’t always chemically bound to the protein and instead binds by other mechanisms. For example, some fluorophores are adsorbed into the protein binding sites and can be used to learn more about the binding site structure and its affinities. Here’s a great chapter from Methods in Cell Biology that explores protein labeling with fluorescent probes in more detail.

Protein labeling with fluorescent probes such as fluorescein can help analyze brain slices in mice
Green Fluorescent Protein (GFP) fluorescing in a mouse brain slice (source)

What Types of Fluorescent Probes Are There for Protein Labeling?

Since there is so much diversity in proteins, it of course makes sense that there is a huge range of fluorescent probes available to choose from. Organic fluorescent dyes are the most common way of labeling proteins. They are excellent choices for protein labeling because they can be fine-tuned by changing their structure to make them more target-specific or to adjust their fluorescence.

Types of fluorescent probes for protein labeling include cyanines, rhodamines, fluorescein, biological proteins like GFP, and quantum dots. 

Cyanine-Based Fluorescent Probes for Protein Labeling

These synthetic dyes contain conjugated polymethine chains with quaternary nitrogens as part of the system. Their fluorescent properties are easy to adjust depending on the functional groups and length of the conjugated chain. They often yield brighter and more stable fluorescence than alternative organic dyes. Here’s a great overview of cyanine dyes from Science Direct.

Related articles:

Rhodamine-Based Fluorophores

This family of dyes is mainly used for dying paper and as inks, but they also make excellent protein labels. They are high-performance dyes that are excellent for labeling antibodies in particular.

Fluorescein Labels for Proteins

The grandfather of organic fluorescent dyes for protein labeling. Fluorescein is one of the most important and successful fluorescent dyes. It is even listed as one of the WHO’s essential medicines.  There are countless fluorescein derivatives in a huge range of applications so they remain the kings of protein labeling.

Biological Fluorophores

These fluorophores are formed from biological structures that can fluoresce. While they are often more expensive and time-consuming to use, they can be bound to proteins very effectively in specific cases and introduced into living cells, bacteria, or even entire organisms. Biological fluorophores have the advantage of being less likely to result in issues of toxicity by negatively affecting the proteins they are bound to. They can be made from other proteins, enzymes, antibodies, or other common biological structures that can be bound to a protein.

Quantum Dots 

These relatively new fluorophores are significantly brighter and more stable than organic fluorescent dyes. They are tiny (nanometer scale) crystals made from semiconductors. Their fluorescence is linked to their size and shape, and they are exceptionally stable (one study reported quantum dots fluorescing for 4 months in vivo!). However, they are still novel technology and need more research. The main challenge with quantum dots is their toxicity. Since they are made from heavy metals and have high stability, they are potentially very toxic depending on a range of parameters such as their size, shape, composition, and more. They are powerful tools for protein labeling as they can be coated in different ways to optimize their binding. 

Quantum dots are stable fluorescent probes for protein labeling
The fluorescence wavelength of quantum dots can vary depending on their size (source).

Tips to Select A Fluorescent Probe for Protein Labeling

Proteins are large, complicated biological structures with potentially thousands of relevant functional groups and binding sites, as well as a specific 3D structure. This means that choosing the right fluorescent probe for your target protein is essential. However, there isn’t always literature on every protein and fluorophore, so you’ll have to do some experimentation to make sure you get the right fluorescent probe for the job.

To select a fluorescent probe for protein labeling, choose a fluorescent probe that’s specific to your protein, in a detectable wavelength, stable in your experimental conditions, and doesn’t interfere with other fluorophores or components in your experiment.

1. Determine What Protein You Want to Label

This seems obvious but it’s the most important factor. You need to know exactly what you’re looking for, or there isn’t much point trying to label a protein. Are you targeting a specific family or protein? Or one particular protein? The more you know about your target, the easier it will be to choose a fluorescent probe. You’ll also need to analyze how to purify your recombinant protein with your fluorescent label.

2. Determine if Experimental Conditions Will Quench or Interfere With a Fluorophore

What are you trying to do once you’ve labeled your target protein? Monitoring intracellular protein processes in real-time might require a very stable fluorescent probe that has low toxicity. What if you’re trying to purify your target protein? That might open your options to include fluorescent probes that aren’t as long-lived but are brighter and will give you more precision.

3. Choose a Fluorophore That Binds Specifically to Your Target Protein

Your fluorophore needs to be specific in binding to your target protein. It’s no use to you if your label binds to various proteins that you aren’t interested in and gives you false readings. Worse yet, a poorly chosen fluorophore might bind to multiple proteins at once. We’ve discussed protein conjugation chemistry in detail in another article.

4. Find a Fluorophore That Can Be Detected Easily 

Great, you’ve added your fluorescent probe to your protein, but can you see it? Your fluorescent probe needs to be bright and easy to detect in your sample. If your sample matrix affects the fluorescent of your probe, you’re not going to get an accurate measurement. It’s especially important that it doesn’t fluoresce at the same wavelength as any component of your sample of you’re going to get a false reading.

5. Ensure That Your Fluorophore Is Stable in Your Experimental Conditions

Your fluorescent label of choice needs to be stable in your sample and when bound to your target protein. It will need to remain stable long enough for you to complete your experiment, but without it affecting the biological system you’re monitoring.

6. Minimize Interference Between Your Fluorophore and Other Fluorophores or Experiment Components

In many cases, your fluorescent probe needs to not interfere with the function of the protein, or the function of the biological system you’re monitoring the protein in. There’s no point labeling a protein if its function is completely impaired by the binding of your label. Worse yet, you won’t be able to monitor proteins in cells or other living organisms if your probe is so toxic that it kills them. When working with living samples, make sure to select a probe with suitable toxicity for the duration of your experiment.

Techniques to Analyze Proteins Labeled With Fluorescent Probes

Once you’ve labeled your protein with a fluorescent probe, there’s an amazing range of potential uses for your labeled protein. Proteins can be observed, quantified, and studied in great detail once you’ve attached a label.

To analyze your protein labeled with a fluorescent probe, you can use techniques such as fluorescent microscopy to quantify and monitor your target protein, flow cytometry to sort proteins, and even monitor living cells using fluorescent live-cell imaging with multiple labeled proteins. 

Analyzing Proteins Labeled With Fluorescent Probes Using Fluorescence Microscopy 

Labeled proteins can be identified in cells and cellular components with exceptional specificity. Many uses extend from this. For example, the levels of proteins expressed in certain tissue can be quantified to understand the effects of a specific gene. Another example is the use of multiple fluorescent probes to monitor a protein and the components it interacts with to provide more detail about the mechanisms occurring within a cell. In medical diagnostics, cancer-specific proteins can be labeled to learn more about specific cancers. We’ve written about immunofluorescence microscopy in detail and discussed how to use antibodies attached to fluorescent probes.

Flow Cytometry Can Sort Proteins Labeled With Fluorophores

Labeled proteins can be sorted and quantified in real-time as they pass through a light beam of detectors that measure their fluorescence. In medicine, this technique can be used to rapidly screen for medically relevant proteins. This technique is used in immunology (for example, antibodies), hematology (blood proteins and other markers), oncology (cancer-specific proteins as mentioned above), and even genetics (measuring gene expression by protein markers). Flow cytometry is a popular technique because it is relatively cheap and reliable. We’ve discussed the flow cytometry method and theory behind this technique here.

Live-Cell Imaging Can Be Used to Visualize Proteins Labeled With Fluorescent Probes in Biological Systems

Labeled proteins are extremely useful in monitoring intracellular processes in real-time using time-lapse fluorescence microscopy. It provides insights into the life of a cell and the active processes within. As mentioned above, the use of multiple fluorescence probes can provide intricate details to the internal structure of a cell. However, a careful balance needs to be found between collecting enough data without producing phototoxic effects from overusing the fluorescent probes.

Related articles:

  • Fluorescent western blotting is a simple and effective technique for analyzing labeled proteins

Where Can I Buy Fluorescent Probes?

Most major chemical suppliers sell specific fluorophores. Organic fluorescent dyes are the most commonly available types but now more unusual types like quantum dots are becoming available.  Many suppliers produce protein labeling kits that contain everything needed to complete the process in a short time. Some offer the ability to label up to 10 mg of protein.

Fluorescent probes can be bought from most life science suppliers such as Anaspec, ThermoFisher, and LiCor.

Here are some examples of protein labeling kits found on the market:

AnaTag 5 microscale protein labeling kits. These labeling kits use fluorescein isothiocyanate (FITC) as a fluorophore. The kit is suitable for biological applications and can label 3 x 200 ug of protein.

Alexa Fluor protein labeling kits from ThermoFisher. These kits offer a very straightforward way to covalently label 1 – 10 mg of protein with a fluorescent dye. They offer a range of dyes with a broad range of excitation and emission wavelengths.

LI-COR IR Dye protein labeling kits.  These kits are designed for labeling antibodies for use in flow cytometry where fluorophore-conjugated antibodies are required.

Step-by-step Example of Labeling a Protein With a Fluorescent Probe

The most straightforward way to label a protein with a fluorescent probe is to use a protein labeling kit like those mentioned above. Most of these processes use a similar process to label the proteins. Below you’ll find the general steps that protein labeling kits like this one. They are very easy to use and require a small time investment. 

Materials Needed for Protein Labeling

·        Protein labeling kit

·        A suitable amount of purified protein

·        30 minutes of hands-on time

·        2 hours until the protein is ready

General Steps to Label With Fluorescent Probes

Step 1. Add Your Protein Into a Vial With the Fluorophore and a Magnetic Stir Bar 

Most kits come with a premeasured quantity of dye and vial for you to perform this step.

Step 2. Let the Reaction Occur With Gentle Stirring 

The kit will tell you how long you need to stir the reaction. Be careful not to stir too aggressively as some proteins are sensitive to agitation.

Step 3. Purify the Protein Using the Size Exclusion Column Provided 

These are usually gravity-fed size exclusion columns that purify out your protein.

Step 4. Collect Your Purified Labeled Protein

Collect your protein from the column. Now you can perform your experiments on the protein.

As you can see, the process is fairly simple and requires minimal effort as all of the equipment you need to label the protein is provided in the kit.

Protein Conjugation Chemistry – Theory and Examples

We discuss protein conjugation chemistry techniques such as lysine, cystein, or site-specific conjugation using bifunctional linkers.

Before we do a deep dive into protein conjugation chemistry, let’s first describe what a conjugated protein is.

A conjugated protein is a protein to which another chemical group or molecule has been attached. Typically, covalent bonding interactions are used to conjugate molecules to proteins.

Proteins that contain only amino acids are generally referred to as free proteins

Conjugated proteins can be attached to carbohydrates, lipids, organic complexes and may even have stabilized metal ions. 

Conjugated proteins contain free proteins, as defined earlier, attached to molecules, called prosthetic groups.  When carbohydrates are bound to a protein, the protein is called a mucoprotein or glycoproteins (considered to be the most abundant and largest conjugated protein). Some categories of conjugated proteins also take the form of proteolipids or lipoprotein. Proteolipids have the tendency to behave typically like lipids and are quite soluble in organic solvents. Lipoproteins (lipids attached to a protein molecule) behave typically like a protein. The lipid inside a lipoprotein imparts a lower density to the protein; which means a lipoprotein has a relatively low physical weight.

You can also have Metalloproteins and Chromoproteins, which are other derivatives of conjugated proteins. A metalloprotein has a metal group specifically bound to an amino acid moiety. A chromoprotein is a colored protein composed of a protein and a chromophore (functionality that gives them color).

A typical metalloprotein called hemoprotein is represented below. Note the way the iron ion is bound.

A metalloprotein that has conjugated iron


Hemoglobin is a conjugated metalloprotein that binds iron. Image from ScienceDirect.

Related articles:

Bifunctional Reagents for Protein Conjugation Chemistry

A bifunctional reagent features reactive or some very active end groups or molecules that make bioconjugation (crosslinking) possible. These bifunctional reagents can actively bind to compatible functional groups through their active end groups. 

Research has shown four main functional groups make effective target moieties on proteins and prosthetic molecules. These functional groups include;

  • Primary amines: A primary amine has an amino functional group (- NH2). We can have the aliphatic amine (R-NH2) and the Aromatic amine (Ar-NH2). Aromatic amines are more reactive and easier to conjugate to proteins than aliphatic amines. 
  • Carbonyls: Identified by the carbonyl functional group (R-C=O).
  • Thiols (sulfhydryls): This is a functional group that has sulfur bonded to a hydrogen atom. They are usually denoted as R-SH.
  • Carboxyls: Denoted R–COOH. 

A closer look at all four bifunctional groups shows that they are also a part of amino acids, so they are easy to find on proteins. 

protein conjugation chemistry using cystine

Some common homo and hetero bifunctional reagents for protein conjugation. Image from SpringerLink.

Homobifunctional Reagents for Protein Chemistry vs. Heterobifunctional Crosslinkers

Bifunctional reagents can be homobifunctional or heterobifunctional. 

Homobifunctional reagents have the same reactive group on either side and can be utilized for one-step protein conjugation chemistry. Heterobifunctional reagents have different reactive groups on each side and are typically utilized in multistep conjugation reactions with proteins. 

Researchers have also utilized homobifunctional crosslinkers with N-hydroxylsuccinimide (NHS) esters to identify previously unknown interactions between proteins. 

How to Use Bifunctional Reagents For Protein Conjugation

If you’re wondering how to utilize a bifunctional reagent for protein conjugation, in general you need to:

To use a bifunctional reagent for protein conjugation, first determine the target moiety on your protein and your prosthetic molecule. Then find a bifunctional reagent that will bind both your free protein and your prosthetic molecule. Finally, utilize the appropriate protein conjugation conditions to attach the protein to your linker and then conjugate your protein, with the crosslinker, to the prosthetic molecule.

Here are more details on the above general steps. 

Step 1. Determine A Target Moiety On Your Protein First

Amino acids on proteins consist of carboxyl groups and amino groups. Since proteins are linear polymers consisting of amino acids, one end of the protein will have an unconjugated carboxyl and the other end will have an unconjugated amine group. The carboxyl end is called the C-terminus and the amine end is called the N-terminus. Both of these ends can be conjugated to. Protein labeling with fluorescent probes is a common reason to utilize the N- and C- termini.

You can quantify the N-terminus along with any free amine groups in a protein by reacting the protein with fluorodinitrobenzene (FDNB) or dansyl chloride. The fluorodinitrobenzene (FDNB) or dansyl chloride will link with any free amine inside the protein. If you notice that there are a lot of amines, then consider using bifunctional linkers with at least one NHS group to react to the amines on the protein. See below for lysine conjugation chemistry.

Other viable target moieties on your protein could include thiols from cysteine residues or the C-terminus carboxyl group. See the image below for amino acids that are easy to react with. 

lysine, cysteine, aspartic acid, and glutamic acid are excellent for making protein conjugates

Lysine, Aspartic Acid, Cysteine, and Glutamic Acid are good target moieties for protein conjugation reactions due to their amine and carboxyl side chains. Image from NEB.    

Step 2. Determine a Target Chemical Moiety on Your Prosthetic Molecule

Conjugated proteins are proteins linked with chemical groups on other molecules, called prosthetic molecules. Prosthetic molecules can be carbohydrates, lipids or even ions. 

You need to determine which chemical moiety on your prosthetic molecule you will conjugate to the protein of interest using the bifunctional reagent. 

The best moieties on your prosthetic molecule to bind to are amines, carboxyl groups, or sulfhydryls as we stated previously. 

If you can install an azide, tetrazine, or alkyne on your prosthetic molecule, these groups can make your protein conjugation chemistry really easy. 

If none of these groups are available on your prosthetic molecule, consider converting a hydroxyl to an aldehyde to make it more reactive.

Typically nanoparticles that are commercially purchased can include functionalized polymer surfaces that you can conjugate to proteins. We discuss protein and antibody bioconjugation to gold in our article.

click chemistry is commonly utilized for conjugating proteins

Azide-alkyne ‘click’ chemistry is a common method utilized for protein conjugation chemistry. It’s considered ‘bioorthogonal’ because azides and alkynes are uncommon in proteins. Image from Royal Society of Chemistry.

Step 3. Find a bifunctional reagent that can react with both chemical moieties

Bifunctional reagents are important in the protein conjugation process as we’ve discussed previously. 

The bifunctional reagent that you use to conjugate your protein to the prosthetic molecule must include chemical groups that can react with your protein (from step 1) and chemical groups that can react with your prosthetic molecule (from step 2).

If the two groups from Step 1 and Step 2 above are different, you need to use a heterobifunctional reagent. Western blotting can be utilized to prove that your conjugation reactions yielded higher molecular weight results.

Using heterobifunctional reagents for protein conjugation chemistry

To use a heterobifunctional reagent, first react it with the protein. This reduces steric hindrance compared to the opposite case where you first react with the prosthetic group because your linker bifunctional reagent will be able to reach deeper parts of your protein without the prosthetic group interacting with the protein. 

After reacting with the protein, you can react to the other functional group on the bifunctional reagent to react with your prosthetic molecule. 

Note that two steps are involved for heterobifunctional reagents in order to avoid polymerization or side linkage of functional groups. 

Using homobifunctional reagents for protein conjugation

Homobifunctional reagents can be utilized in one pot reactions containing both the protein and the prosthetic group or separately in two different conjugation reactions. 

Site-specific protein conjugation

For site specific protein conjugation, researchers typically include unnatural amino acids inside their protein during expression. Cysteine groups could be used for site specific conjugation if only one disulfide bond is present in the protein and hence only those sulfhydryl groups are accessible to your bifunctional reagent.

Bioorthogonal reactions for protein conjugation

Azids, ketones, and aldehyde can be coupled to a protein via bioorthogonal chemistry. Typically, bifunctional reagents utilized to conjugate to these won’t react with amines or carboxyls on the protein. Ketones and aldehydes can be reacted with aminooxy or hydrazide compounds to yield a very stable oxime or hydrazone linkage between the protein and the biomolecule. 

Lysine Conjugation Chemistry

Lysines are the most common targets for protein conjugation chemistry reactions. The amine on the lysine is essentially a nucleophile and reacts to amine-reactive chemicals such as N-hydroxysuccinimide (NHS) ester (these react via lysine acylation).

For successful Amine-NHS conjugation reactions, the pH of the reaction solution must be below 10.5, which is the pka of the ammonium group in lysine. This helps deprotonate the lysine groups. This means that the reaction should be carried out at ph around 8.5 to 9.0. 

Lysine may also react with isothiocyanates and isocyanates and benzoyl fluorides.

Advantages of lysine-based conjugation

  • Simple
  • Commonly utilized for antibody conjugation
  • Highly reactive, if they are accessible
  • Versatile because it can react several cytotoxic agents (especially onto antibodies)

Bioconjugation strategies for lysine residues on proteins. Image from Nature

Maleimide Protein Conjugation Reaction Chemistry

You can react cysteines in proteins with maleimide groups as long as your buffer is in a pH between 6.0 and 7.0. A common approach is to reduce disulfide bonds in proteins first with DTT or TCEP and then to react with a maleimide-containing linker.

At a pH between 6.0 to 7.0, the maleimide group can react with sulfhydryl groups resulting in the production of a stable thioether linkage and the reaction is irreversible. At pH above 8.0 (alkaline), say 8.5, the reaction seems to favor primary amine and the rate of hydrolysis of the maleimide group to maleimide acid (not reactive) will increase.  

You must eliminate compounds that contain thiol groups, like dithiothreitol (DTT) and β-mercaptoethanol (BME, aka 2-mercaptoethanol), from the reaction because they tend to compete with thiol groups on proteins. For instance, if you utilized dithiothreitol (DTT) to reduce disulfides in a protein and open up the sulfhydryl groups, you would need to thoroughly remove the DTT using a desalting column before you start the maleimide reaction. Instead of DTT, consider using the disulfide-reducing agent TCEP which lacks thiols. 

You can quench free maleimide left over after reacting your protein by adding free thiols. You can also check your conjugation reactions using our HPLC Step By Step guide.

Advantages of Maleimide Protein Conjugation

  • Thiols are only present in proteins on Cysteines
  • For many proteins, one or two disulfide bonds exist, which mean that you can reduce them and conjugate in a site-selective manner using thiol-maleimide chemistry
  • Cysteines can be readily introduced into proteins without affecting function, using site directed mutagenesis

Here are some strategies for using thiol-maleimide protein conjugation chemistry to create cleavable and permanent antibody drug conjugates. Image from RSC.

Protein conjugation protocol example

There are several protein conjugation kits that are available. Here’s an example from Vector labs. However, you can also utilize the chemical reactions above to create protein conjugates. 

The steps below show a typical example of conjugating to lysines on a protein, based on information from GBiosciences


  • A PEG linker containing NHS such as any of these
  • DMSO or DMF 
  • Amine free buffer at pH 7-8. Phosphate buffer works well
  • 0.5-1.0 NaOH
  • Protein containing Amines
  • Desalting column


  1. Dissolve 1-2mg the NHS ester reagent in 0.25 ml of DMSO or DMF
  2. Then add 2ml of phosphate buffer to the same reaction vial
  3. Add 100µl 0.5-1.0N NaOH to 1ml of the reaction from step 2.  Vortex for 30 seconds
  4. Check that the pH is between 8.5 and 9
  5. Add your protein in pH 8.5-9 phosphate buffer to this vial
  6. Use a desalting column to remove any unreacted linker molecules. Here’s a step by step method for protein purification of recombinant proteins.
  7. Utilize the other end of the PEG linker to conjugate a prosthetic molecule

Biopolymer surface functionalization: Simple Step-By-Step guide

Biopolymer Surface Functionalization of Biomaterials

Introduction to Surface Functionalization of Biopolymers

Immobilizing (or covalently attaching) proteins, lipids, carbohydrates, and other polymers on biopolymer surfaces is incredibly important for a number of reasons. Want your surface to be hydrophobic or hydrophilic? Want to attach interesting fluorescent molecules on a sensor surface? There are a ton of possibilities! One of the most common uses for biopolymer surface functionalization is Surface Plasmon Resonance. Here, a protein is covalently attached to a gold surface and several different ligands are flowed past the protein-surface. Researchers can then study the binding and unbinding of ligands to proteins on the gold surface and determine on/off rates etc.

For more information on methods for protein bioconjugation to gold, take a look at our article.

Take a look at the image below:

SPR Functionalized Biopolymers

Another reason for immobilization of materials on a biopolymer surface might be to make it more hydrophilic. For a long time we have known that functionalizing long hydrophilic polymers on a surface can help prevent clotting and protein binding. This is one of the key methods for improving the blood compatibility of biomaterials. Without proteins to bind the surface and subsequent activation of platelets, biomaterials can be used inside the body for longer periods of time and they can even be implanted!

Curious about how you can tell if your protein attached to the surface? Label your protein with a fluorescent probe with our method!

PEG Functionalized Biomaterial Surface

A Super-Simple EDC/NHS method for Surface Functionalization of Proteins on a Biopolymer

A common method for modifying the surface of a carboxyl-containing polymer with protein, is to attach the N-Terminus of the protein onto the surface. Here is a simple representation of the chemistry:

EDC NHS Biopolymer Surface Functionalization

Materials for EDC/NHS Surface Immobilization of Proteins

  • Coupling Buffer: We need to make sure that your protein is neutrally charged using an appropriate buffer (and your knowledge of the isoelectric point, pI, of the protein). Make a buffer with 100 mM Formic acid (pH 3-4.5) , acetic acid (pH 4.0 – 5.5), or maleic acid (pH 5..5 – 7.0) in water. Use NaOH for pH equilibriation.
  • EDC [1-Ethyl-3-(3-dimethylamoniminopropyl) carboodiimide] at 0.4 M in water. Store at -20 C in small aliquots.
  • NHS [N-Hydroxysuccinimide] dissolved in water at 0.1 M. Store at -20 C in small aliquots.
  • Ethanolamine Hydrochloride dissolved in water at 1 M concentration, pH 8.5. Store at -20 C in small aliquots.
  • Your Protein of Interest at 50 ug/ml in an appropriate buffer.

Step-by-Step Biopolymer Functionalization Methodology

  1. Wash the biopolymer surface with coupling buffer
  2. Thaw EDC, NHS, and Ethanolamine aliquots.
  3. Incubate the protein of interest with EDC and NHS at a 1:1 EDC:NHS ratio. Also, incubate the biopolymer surface with the EDC/NHS solutions. Leave at room temperature for 10 minutes.
  4. Wash the biopolymer solution with coupling solution 3 times.
  5. Add the protein + EDC/NHS solution onto the surface and incubate for 15 minutes.
  6. Wash the surface with coupling buffer
  7. Add ethanolamine solution onto the surface and incubate for 10 minutes
  8. Wash the surface with your protein storage buffer to re-equilibriate.

You can also utilize protein conjugation chemistry to impart unique tags onto your proteins that make them easier to functionalize onto surfaces.

Notes on this Surface Functionalization Methodology

  • If your biopolymer doesn’t have carboxyl groups, this methodology will not work. Choose an appropriate coupling technique based on the surface you’re trying to functionalize.
  • EDC is hygroscopic and breaks down quickly in water. Keep the solid EDC under dry gas and if you have any EDC solutions, make sure to use them quickly or freeze them!
  • Don’t reuse thawed EDC aliquots.
  • You can change the incubation lengths to improve coupling efficiency between the protein and the surface.

Other Surface Functionalization methods on SciGine

Great Homebrew Video of Surface Modification

Literature References for Biopolymer Surface Functionalization

Tips For Making Research Efficient

Insights from Priyamvada Jayaprakash. Connect at Her LinkedIn Profile. She’d love to discuss her PhD experience with you and get insights on opportunities related to tumor immunotherapy.

Challenges I Faced During my Ph.D

I love Biology

Biology has always fascinated me. Understanding how cells and their environment team up to create a functioning organism mesmerizes me. So, without a second thought, I decided to pursue my Bachelors in Biotechnology when I was a senior in high school. During my coursework, I gained an introduction to diverse areas spanning genetics, biochemistry, cell biology and immunology. I learned to appreciate not only the complexities of organisms at a molecular level but also the fact that some of these molecular underpinnings can be explored and manipulated for the betterment of people. Towards the end of my 4 years, I realized that my passion for science wasn’t quelled by my undergraduate education and I pursued my scientific journey by accepting a PhD position in the University of Southern California (USC), Los Angeles. My thesis was on understanding the importance of secreted heat shock proteins (Hsps), Hsp90α and Hsp90β, in wound healing and breast cancer metastasis.

Note: If you’re writing research papers, I highly recommend Grammarly – it’s a free grammar check plugin for Chrome. Try it out here

Like any other PhD student, I spent the initial few months (and a lot of time in the future) reading and understanding “literature”. This meant a thorough knowledge and understanding of what findings have been accomplished in my research area- what ideas have worked and more importantly, not worked. Fellow researchers would definitely know “Pubmed”, an extremely useful database from NCBI (National Center for Biotechnology Information), that lets you search for papers of a specific area- all in one location. I cannot emphasize enough how much literature is important in the life of a PhD. It is especially important for a a naïve undergraduate like me from a foreign land, with no previous experience in a research lab.

Question. What are the downfalls of “literature search” in general vs. PubMed specifically? Or are they the same?
Priya: Searching in general gives a lot of redundant and irrelevant information such as random articles/blog posts etc, most of which are not helpful. But I feel Pubmed search is more streamlined to identify publications of interest. To the best of my knowledge, finding papers is either through Pubmed or a general Google search.

The Research Process is Inefficient

Hence, it came as a shock to me when I realized Pubmed is probably the only database that lets you search for papers in one go. More appalling was the lack of accessibility to all journals and papers one might need. Most journals charge an “article processing fee” for making research work “open access” and unfortunately, many labs are not financially equipped to pay such fees. As a result, research is not freely available to everyone in a timely manner. This is further exacerbated by the fact that the research environment is for the most part competitive, rather than collaborative. A number of my friends and myself have been instructed to be cautious while sharing interesting findings that were still unpublished. Conferences are a great resource of getting to know what’s out there, but most labs are skeptical about sharing their new findings in the fear of getting “scooped”. However, an ideal research environment is one where everyone focuses on asking the right scientific questions, identifies relevant ways to answer the questions, establishes productive collaborations and works toward the betterment of the society. We should create a forum where researchers are able to share their findings with an open mind and get ideas from fellow colleagues- our scientific progress can only be improved by engaging more scientific minds.

Question. This supports ResearchGate. Do you think it adequately solves your challenges? What’s missing?
Priya: Honestly, I need to use ResearchGate more to comment on it. But from what I know, more scientists need to be part of it and be made aware of it as well.

Funding Issues have changed Science

In the recent past, there has also been a rapid decline in funding. NIH funding is being allocated to projects that have a high probability of being a success. Hence “risky” projects have taken a back seat, which means innovation, the most important quality of a research, is at stake. I did my PhD in a lab that was hit by the funding decline. The lab worked at the junction of basic and translational research. We performed in vitro experiments and identified secreted Hsp90α as an important pro-motility factor. We then performed in vivo experiments in mice and pigs to prove the importance of the protein in wound healing and breast cancer metastasis. Compared to labs that worked on drug screening, my PhD lab, similar to most labs, focused on identifying something that could definitely benefit people, in the long run. Hence, I had to restrict my ideas to experiments that we could afford. The funding decline also meant that scientists were not open to trying out new things. Scientists feared any change and “played safe” with their ideas. The goal has now become “publish or perish”.

Question. What does this physically mean? Is your research stifled?
Priya: The funding decline did not affect access to literature. My experience is mainly from inability to buy the right reagents/chemicals required for research. Recombinant proteins and antibodies are expensive and sometimes, you prefer buying these only from specific companies due to the quality. Thus, in case of a funding decline, you either do not buy the reagents you need or you settle for one of less quality. Thus research does get stifled.

Question. What does it mean to “play safe” and not really expand scientific discovery?
Priya: I can comment on this from the perspective of a researcher working in cancer biology. Cancer is a complex and heterogenous disease and 1) multiple targets are required to treat the disease 2) multiple approaches to identify the targets are required. Thanks to modern technology, there are high throughput screening methods to simultaneously identify multiple targets specifically expressed by cancer cells. However, these methods are expensive to execute and many labs are unable cannot afford them. Thus, many target proteins are understudied, which means the number of “druggable” targets in cancer patients are being understudied.

From the point of view of a lab with low funding, identifying a single protein that is specifically expressed by cancer cells can fetch them a paper and therefore more funding. This highlights the “publish or perish” scenario, but does nothing to advance scientific contributions to cancer research. In other words, “playing safe” affects scientific progress globally.

Research Lacks the Right Tools

As a graduate student, there were a lot of things I learned, apart from the science. I spent a lot of time identifying the right reagents for my experiments. Any student who has done western blotting will know that many times antibodies either a) do not work, b) have a lot of background noise or c) have non-specific binding. Not all antibodies work for all purposes. Hence, antibodies from a specific company may not all be amazing. Even though I use literature as a starting point, it requires a lot of trial and error, which means a lot of time and effort on the my part. But, not many resources have helped with this. I used VWR as an online resource to purchase reagents. However, VWR is merely a database that lists different companies offering a specific product. It provides no information on how good the product is.

Hmm, this gets me thinking…I’m happy spending time on Amazon getting a case for my iPhone. Amazon gives me a table comparing different vendors, with specific features of each vendor, the associated product, the price and reviews of each. Why can’t there be an Amazon for science? It would make a huge difference to my professional life. I would highly appreciate a resource that does this and provide reference papers that have used the product. Similar to the star ratings we find on Amazon or eBay, it would be very helpful and time saving for researchers if there were a good rating system for life science reagents too.

Identifying the right protocol is a challenging task too. We need to conserve time and resources…it would be a great help if some online platform provided references to other researches that have already standardized the protocol, along with information on the reagents used.

Question. How do you currently search for protocols and products?
Priya: Currently, I use a general Google search to look for protocols and VWR for most of my purchases. The problem is mainly that I get a lot of redundant and irrelevant information. In addition, it is difficult to verify if the protocols have been standardized and come from a reputed lab or institution. Also, there’s no proof if the protocol has worked for them or not. Hence, if I have a visual evidence (aka reference papers), it is more convincing.

Conclusions and More Ideas

I consider myself very fortunate to have obtained admission into a prestigious graduate program in the United States. I came to the country with no prior experience. When I look back, I think that certain tools may greatly benefit many young students. One tool I can think of is a visual pathway explorer. Biological processes are dynamic in nature and I have always wanted resources that can teach me a specific biological process more visually. Currently, I use YouTube for this purpose, but it does not have videos for everything. I think it would be amazing to have an interactive tool where a pathway/process is taught illustratively and questions can be posted on the website, which will be addressed by experts. Imagine how many ideas can be sprung out of this simple idea!

Science is fascinating, but with challenges of failure and lack of accessibility to resources, it can get pretty frustrating too. Thus by providing the right accessibility to constructive collaborations, the right protocols, and the right reagents, the life of scientists can be more productive.

Question. If you wanted to make research more efficient. What kind of company would you make? What products would it have? Assuming you have an unlimited budget and/or time.
Priya: I would create a company that acts as a liaison between labs and service/reagent companies. For example, for my research, I would need diverse products ranging from a specific custom-made cell line, lentiviral knockdown systems, CRISPR/Cas9 reagents, specific transgenic mouse models or diverse services ranging from microarray analyses, RNA seq analysis etc. Diverse types of companies provide these reagents and services. For example, IDT is the company I use to generate my oligo constructs for cloning while I use Jackson labs for mouse models and Affymetrix for microarray services.

My company would be like a liaison that obtains the needs of specific lab (s) and get in touch with these diverse companies. Every lab can create a profile on the company website and use it as a single portal to add these diverse orders. My company would communicate with the different companies and provide the reagents & services to the labs. This will be the service wing of the company. In addition, my company will also provide analyses software for the different high throughput techniques at a subsidized rate for customers. If feasible, certain bioinformatic analyses might also be performed by my company for a subsidized rate.

In addition, my company will also have a scientific wing where I would have a board of expert scientists, postdoctoral researchers and other experts in different fields, for eg, neuroscience, immunology, proteomics etc. If a lab is confused about how to address a specific goal, they can post the questions via their profile on my company website and they will be addressed by the respective experts.

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

SciPrice and SciGine are Solving Challenges in Research

Many of the challenges that Priya has outlined are now being tackled by SciPrice and Scigine.

SciPrice is a marketplace for antibodies which focuses on providing all the information possible for each product. It’s an Amazon for Science.

Scigine provides protocols for Biology from reputed journals along with the materials that were used. In combination with SciPrice, both services help make research more efficient.

Thank you for reading!

Ligate Sticky Ends via DNA Ligation

DNA Ligation of Sticky Ends

Ligation of Sticky Ends, Summary of DNA ligation

We have already discussed a high level view of gene cloning in our Molecular Cloning Guide blog post. However, in that blog post we didn’t delve very deep into how we can perform each of the individual steps. Today’s blog post is about ligation. Ligation is the process by which two pieces of DNA can be glued together to form one piece. So, to begin, let’s assume you’ve already decided on a gene product that you want to clone. You’ve also designed primers and completed PCR on the open reading frame in your donor DNA (this could be genomic or non genomic DNA). Your next steps are to digest the PCR product with restriction enzymes and generate sticky ends. You’ll also want to digest your “shuttle” plasmid to generate complimentary sticky ends which will allow your “insert” DNA to click into position into your vector. It’s like a puzzle piece!

Note: If you’re writing research papers, I highly recommend Grammarly – it’s a free grammar check plugin for Chrome. Try it out here

Note: It might be useful to look at our RNA Extraction & Isolation guide if you’re planning on making cDNA related to your gene

The above summary is demonstrated here:
Ligate Sticky ends using Ligase

Sticky Ends Insert into a Shuttle Vector

Only some Restriction Enzymes Create Sticky Ends

As you can figure out, generating sticky ends and complimentary ends is extremely important to the above process. However, several different restriction enzymes are available and each of them has different locations where they cut. Also, the type of cuts that they introduce may be “sticky” or “blunt”. Depending on the cloning strategy you are using, you may mix and match different enzymes to achieve different end goals. Ligation of “sticky ends” is much more efficient than ligation of “blunt” ends. Typically 10-100 times more T4 Ligase is required for blunt ends.

Here’s an image with various restriction enzymes and the kinds of ends they produce. Depending on the type of ends, your DNA ligation will proceed very differently!

Restriction Enzymes for DNA Ligation

Ligate DNA via DNA Ligase

Once the restriction enzyme digestion is complete, you can proceed to the ligation step. But, before you digest anything, make sure you’ve planned everything properly! You need to make sure that the insert will be ligated in the proper direction in the shuttle vector. Only once you’ve vetted your overall strategy, should you proceed to ligation and transformation, etc.

There are several kinds of ligase enzymes but the enzyme produced by T4 bacteriophage-infected E. Coli is the most common one. This ligase is called T4 ligase. Whereas normal E. Coli produce DNA ligase that uses NADH as a cofactor, T4 infected E.Coli produce a ligase that uses ATP as a cofactor. This enzyme will find the 3′ Hydroxyl and 5′ Phosphate within your sticky ends and it will form a phosphodiester linkage. If this is confusing, check out the Polymerase chain reaction (PCR) guide for images on what DNA looks like. This is shown here:

Ligation Protocol for T4 Ligase

Phosphodiester Bond Formation during Ligation

Protocol for Ligation of Transgene Insert into Shuttle Vector

Ligation enables fragments of DNA to be combined, such as the cut ends of transgene inserts and plasmids during cloning. This protocol describes the directional cloning of a XbaI/SalI-digested transgene into a shuttle vector, pAdtrackCMV, via cohesive end ligation.

Materials for DNA Ligation

XbaI/SalI digested, gel-purified insert (approx. 1 kb) and pAdTrack-CMV shuttle vector (approx. 9.3 kb; Plasmid #16405, Addgene)
Quick Ligation Kit (contains DNA ligase and 2X Reaction Buffer; #M2200S, New England Biolabs)
Agarose plate containing ethidium bromide
DNA standards

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

Ligation Methodology

  1. Estimate the DNA concentration of purified insert and vector preparations by applying 1 µl to an agarose gel plate (+ethidium bromide) alongside a range of DNA standards and visualizing under UV light.
  2. Prepare the ligation mix as follows:

    XbaI/SalI digested pAdtrackCMV 50 ng

    XbaI/SalI digested insert 17 ng

    Add water up to 10 µl total volume.
  3. Add 10 µl of 2X Reaction Buffer and mix.
  4. Add 1 µl of DNA ligase and mix.
  5. Microcentrifuge briefly to settle liquid to the bottom of the tube and incubate at 25°C for 5 min.
  6. Place on ice* and transform into desired bacterial strain.

Tips and Tricks for DNA Ligation

  • This reaction setup is using a digested insert to vector DNA molar ratio of 3:1. Inserts of different sizes will require a different amount to be added. Important ligation control reactions to include are (1) digested vector only and (2) digested insert only.
  • Ligation reactions can be stored at -20°C for future use

Applications of Ligation on SciGine

Construction of PB42 Vectors Via Ligation
Plasmid Construction via PCR and Ligation
Plasmid Ligation and Transformation in Yeast
Construct with Human p275UTR
Different DNA ligation methods discussed

Video Tutorial About Sticky Ends and Ligation


He et al Proc Natl Acad Sci U S A. 1998 Mar 3. 95(5):2509-14.
Sticky Ends Explained Well
DNA Ligation Theory
Gaastra et al. Ligation of DNA via T4 Ligase
Tsuge et al. One Step Assembly of DNA fragments

Protein Purification of Recombinant Proteins

Protein Purification of Recombinant Proteins

Protein Purification Summary

In our previous blog posts we have explored Gene cloning with Plasmid Vectors in Bacteria, Transient transfection into Mammalian Cells with Calcium Phosphate, and how we can use newly introduced proteins to control biology. Proteins made this way are considered recombinant because they aren’t natively produced in the organism that you got them from.  We really like recombinant technology because it allows us to scale up protein production and generate therapeutic and/or interesting fusion proteins that we can use. If you want some human protein, would you rather grow humans and isolate the protein for scale up (~30 years per doubling)? Or use bacteria instead (~20 minutes per doubling)? Note: this was a joke. Don’t grow humans for protein production 🙂

In this blog post, we are going to explore how “recombinant” proteins can be purified after cells have expressed the gene products that you cloned into them. The strategies explored here can be applied to all sorts of proteins so let’s begin!

Note: You can easily troubleshoot your protein purification procedure by labeling your protein with a fluorescent probe. We’ve given you some more information in our article.

Protein Purification of therapeutic recombinant proteins

Strategies for Protein Purification

Let’s say you have some bacteria that you’ve produced a protein inside. Your first step is to lyse those bacteria and neutralize any proteases that are now in your lysate. Proteases will wreak havoc on all the proteins in solution…so this step is important. Next, we have to think about the recombinant protein that we created in order to purify it. Note that conjugated proteins can utilize their unique tags for purification.

Several different purification methods can be used based on your properties:

    • Protein Charge: If your protein has a overall charge because of excess arginine or aspartamine residues, perhaps it can be purified by running it through an ion exchange column. For negatively charged proteins, use anion exchange chromatography, and for positively charged proteins use cation exchange chromatography. The steps here are simple…Dissolve your protein in a buffer and incubate it with the resin. Wash the resin with some low salt buffer. And then elute the bound protein with some high salt buffer (which breaks the ionic interactions with the resin).

Using Ion Exchange columns for protein purification

    • Protein Size: Dialysis and Size Exclusion chromatography can help you isolate proteins based on their size. In the case of dialysis, you incubate your protein in a dialysis bag and stir it while replacing the buffer outside. Your protein and larger proteins are retained in the bag while smaller proteins are filtered out through diffusion. Size exclusion chromatography (SEC) works similarly to separate out larger molecules from smaller ones. Take a look at our HPLC Step by Step guide to understand chromatography in general.

Recombinant proteins separated by size via dialysis

    • Protein Affinity: If you are lucky enough to buy resin with antibodies vs. your protein, you can simply pass your protein through the resin and it will selectively bind your protein. Then wash it a little bit with buffer so no other proteins are bound and finally elute it by disrupting the antibody-protein interaction.

Affinity based separation of recombinant proteins

SciPrice, biology product search engine

    • Protein Substrate: If your protein is an enzyme with a binding pocket, you can also immobilize your substrate on a column and use that for purifying your protein. Simply pass your protein through the column multiple times so it binds the substrate while other non-functional proteins are easily washed away.

Substrate affinity column for Protein purification

A typical protein purification strategy will involve using several of these techniques in combination. No single technique is 100% efficient, so each time you purify with one of these methods, your protein will get more and more pure. Use a western blot to analyze how clean your protein is. You can also use a silver stain to determine purity. I’ll discuss this technique in the future.

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

Purification of Recombinant Proteins with His Tags

Above, we have already discussed the purification of recombinant proteins via their charge and using their binding pocket. Another strategy that’s very popular is to introduce at least 6 Histidine residues into the N- or C-Terminus of a protein via cloning. Then, when it’s time for purification we can run the protein through a divalent nickel column. Histidine residues, at a high pH (~7.6), can chelate Nickel and hence will be bound on the nickel column. The column can be washed with a low concentration of Imidazole (~20 mM) and then eluted with 150 mM+ of Imidazole.

Cloned His Tags easily chelate nickel for separating recombinant proteins

Step by Step Guide to Purification of a His-Tagged Fusion Protein


Neurospora culture
Lysis Buffer (50 mM Tris-HCl, pH 7.5, 0.1 mM EDTA, 1 mM B-mercaptoethanol)
Protease inhibitor cocktail
Phosphate buffered saline (pH 7.0)
Wash buffer (50 mM Phosphate Buffer pH 7.0, 300 mM NaCl, 1 mM Imidizole, pH 7.0 final)
Elution buffer (50 mM Phosphate Buffer pH 7.0, 300 mM NaCl, 150 mM Imidizole, pH 7.0 final)
Collection tubes for washes and elutions


  • Grow culture and lyse in lysis buffer at 4 C for 45 min
  • Homogenize lysate and centrifuge at 12000 g for 20 min
  • Discard the pellet
  • Dialyze the supernatant against PBS (pH 7.0) for 1 hour at 4 C. Replace the buffer outside the dialysis bag and continue to dialyze for 1 hour more.
  • Prepare the Nickel-Agarose column according to the manufacturers instructions.
  • Add in your protein dialysate from the previous steps on top of the column.
  • Allow the material to diffuse to the bottom and load the filtrate on the column once again
  • Wash the column with wash buffer (use 10x the volume of the beads in the column)
  • Elute the column with elution buffer (use 1-3x the volume of the beads in the column)
  • Collect the eluant in 1 ml fractions and assay each fraction for protein
  • Assaying the protein can be performed via a western blot or other protein assay

Tips and Tricks for Purifying Recombinant Proteins with His Tags

  • EDTA is used in lysis buffer to prevent protease activity
  • Use a dialysis membrane of the appropriate size to retain your protein’s molecular weight + 1000 Da at least. This way you can be sure that you aren’t losing a lot of your protein along with all the filtrate.
  • The size of the column that you use should be determined according to the instructions
  • Protein assays for determining activity are a broad category. For many enzymes  there are assays where the enzyme will be used to cleave a substrate and generate a fluorescent signal.

Protein Purification Protocols on Scigine

Powerpoint related to various Purification Processes


Guide to Protein Purification and Assays from NIH
Protein Purification Powerpoint Presentation
Applications of Protein Purification from Cornell
Manju Kapoor’s Guide to Protein Purification
Nickel-Agarose Purification Guide

Gene Cloning using Plasmids: Molecular Cloning Intro

Gene Cloning using Plasmids via Molecular Cloning techniques

Gene Cloning with Plasmids: Summary

We all know that DNA is the basic building block of biology. So, how can we make use of DNA to change cell biology? Well, today’s blog post will focus on “gene cloning” — making plasmids (circular DNA strands) so that we can introduce them into bacteria using our previous bacterial transformation method. With a plasmid inside the bacteria, you can a) use bacteria to make copies of the plasmid, b) make new proteins with the transformed bacteria and c) do the same inside mammalian cells using the Calcium Phosphate transient transfection method that we developed earlier. With molecular cloning techniques, we can control biology and make cells do some really cool stuff! Note: this is an overview post and does not have a step-by-step protocol associated with it. I’ll tease apart the different steps in future blog posts.

Note: If you’re writing research papers, I highly recommend Grammarly – it’s a free grammar check plugin for Chrome. Try it out here

Molecular Cloning of Plasmids: Primer Design

“Cloning” refers to the process of making a copy of a gene so that we can modify it and see what happens. Remember, if you modify genes, your cells start producing new proteins; these proteins could be therapeutic and/or give your cells some new skills. To start, you’ll probably want to review the PCR protocol & guide to remember how PCR works. Now, let’s say we have a gene that we want to clone already available. The next most important part of PCR based gene cloning is the primer…so to design a primer, we need the following:

  • Hybridization sequence: A series of bases that compliment the bases right before your “target gene” or gene of interest.
  • Leader sequence: A few extra bases for our restriction enzymes to make efficient cuts that don’t overlap with our gene of interest.
  • Restriction sites: Places that we will cut so that we can make the plasmid circular.

Take a look at this image to understand the above plasmid design:
Making primers for Gene Cloning PCR

Molecular cloning primer design

Gene cloning product

Be Careful Designing Plasmid Primers for Gene Cloning

Based on the above image, you can tell that if an enzyme’s restriction site is inside your gene of interest, you cannot use that restriction enzyme because you’ll cut your gene. Also, you’ll be putting this gene into a new plasmid. Make sure that the restriction enzyme you use is compatible with the “multiple cloning site” within this new plasmid. If you end up inserting this gene in random locations, the probability that this plasmid will be incorporated into the bacteria or expanded will be significantly decreased.

Look at the image below to understand these tips:
Gene cloning failure - wrong restriction enzyme

Primer mismatch -- Gene cloning error

Gene cloning with PCR

With the primer already designed, we are ready to clone our gene. The rest of the steps in the gene cloning process are:

  • PCR everything
  • Use restriction enzymes to digest the PCR product
  • Use Gel Electrophoresis to purify the insert and the “vector” (recipient plasmid)
  • Ligate the plasmid
  • Transform bacterial cells
  • Isolate our plasmid for future use
  • Analyze the PCR products

Since we already know how to do PCR from our previous blog post, let’s focus on the other stuff. The first step listed is to digest the PCR product. For this, we will use restriction enzymes and incubate them with the PCR products. If everything was designed properly, we would know exactly where the restriction enzymes will cut the DNA in both the “vector” and the “insert”. Next, we will run these restriction digests on a gel and pick out the bands corresponding to our vector and insert (which we already know the size of). Any other “junk” PCR products will be removed in this step. The vector and insert DNA will then be “ligated” to form our new plasmid. To confirm our gene is in this plasmid, we will transform some bacteria with it on a petri dish. Try to make dilutions of your bacteria so that you can grow colonies of bacteria and pick out colonies later on. With the colonies that you pick out, you’ll want to isolate their DNA and digest it to see if your vector and insert are inside. We’ve already isolated the vector and insert in the past, so it’s simple to find out if our insert is inside the bacteria. Finally, as another confirmation, we will sequence the DNA from the bacteria and confirm that everything exists. We will write more about each of these steps in the future, but we wanted you to see them together, as an overview, in this blog post.

Take a look at the steps below:

Preparing plasmid vectors for Gene Cloning

Electrophoresis and Ligation of Genes using Restriction Digests

Transformation of Bacteria and Isolation of Final Gene Clones

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

Tips and Tricks with this methodology


: Make sure you choose the melting temperature to match the part of the primer that binds the “open reading frame” (your gene of interest). If you choose the wrong melting temperature, you might get the wrong PCR products because either a) your Tm was too low and you didn’t split the ORF or b) your temp was too high and you got lots of non specific binding.

DNA Digestion

: Make sure DNA digestion occurs for a long time, preferable overnight, to make sure all your vector and insert products were cut and maximize your ligation in the next step. You may need to use alkaline phosphatase in this step. I’ll speak more about that in the future.

Gel electrophoresis

: During gel electrophoresis make sure that you run the correct controls and *know* what wells relate to each of the digested products. Also, make sure you skip lanes to make cutting the wells easier. After this, you’ll need to quantify your DNA so you have enough for the ligation step. You can use a UV spectrometer for this step.


: Ligation also requires you to have several controls. For example, you need a ligation reaction without any insert. This will tell you how much background self-ligation your recipient plasmid has. You also need a ligation with some of the other bands you see during your gel electrophoresis. This will tell you how much contaminant DNA there was in your ligation.

Methods related to Gene Cloning on SciGine

Video about Gene Cloning with Plasmids

Notes from our audience

  • “TA cloning is another approach if cloning doesn’t work in systematic way” –Swapnil Oke on Linked In
  • “I think for completeness I think it would be valuable to also mention a few other plasmid features that are important. I didn’t see mention of ribosomal binding sites (RBS) or origin of replication, etc” – Michael Kim on Linked In. — I plan on write about more details regarding plasmid design and purification in the future. For now, please don’t use the above blog post as a comprehensive guide…more like an overview 🙂


Molecular Cloning book about PCR based cloning
Addgene Plasmid Reference, a comprehensive guide
Chaokun et al., Fast Cloning

Bacterial Transformation Protocol with Competent Cells

Bacterial Transformation with Competent Cells

Bacterial Transformation using Competent Cells: Summary

Since we have already learned Calcium Phosphate Transfection with mammalian cells, let’s now focus on bacterial transformation of DNA with competent cells. In general, bacterial cells take up naked DNA molecules or plasmids via a process called transformation. Usually, this happens at a slow rate, but when bacterial cells die in close proximity to others, or when they are stressed, the transformation process occurs at a much higher rate. However, not all bacterial cells can be transformed, so biologists use ‘Competent Cells’ which are more inclined to take up DNA. The end goal of transformation is to get bacteria that have your genes of interest so that they will replicate your genes along with their own. If the bacteria contain your genes of interest, you can use them to mass produce proteins, or just store them for extended periods of time because bacteria are so hardy. A good way to test whether your genes of interest were transformed is to include antibiotic resistance in your plasmid. This way, you can be fairly certain that if your bacteria are resistant to antibiotics, they are also carrying genes of interest to you.

Note: If you’re writing research papers, I highly recommend Grammarly – it’s a free grammar check plugin for Chrome. Try it out here

Take a look at how natural transformation works:
Transformation Protocol with DNA

Transformation Biology in Bacteria

For bacteria, survival is key and transformation is one of their survival mechanisms. As biologists, we can make use of this survival mechanism for our benefit as well. To do this, we first incubate our competent bacteria with our plasmid and calcium chloride. Bacterial membranes are permeable to chloride ions, but not to calcium. So, as chloride ions enter the cell, the bacteria tend to swell (because they also intake water with chloride ions). Then we heat the bacteria in a process called ‘heat shock’ such that they turn on their survival genes. This causes the bacteria to uptake the surrounding plasmids. With the right design, this plasmid will then be recognized by bacterial DNA polymerases (remember our PCR Guide ?) and it will be expressed/replicated along with the bacteria’s normal DNA.

Take a look below to understand how biologists transform cells:

What is transformation

Transformation Biology

Selecting Transformed Bacteria with Antibiotic resistance in a plasmid

Selecting for Transformed Bacteria with the Lac Z Operon

Once your target plasmid is inside the bacteria, you still need to separate transformed cells from those that are not transformed. Another key challenge is that the transformation process may lead to some DNA being recombined so that your gene of interest is no longer functional. How do you select for cells that only contain functional target DNA that hasn’t been recombined? The trick is to use both antibiotic resistance and a Lac Z operon. By cloning your plasmid along with a Lac Z operon, you give your cells the ability to make a galactosidase protein. If cells have the galactosidase and you feed them X-Gal, they turn blue; cells without this operon are white. So, you first transform all your cells. Then you feed them IPTG to activate the Lac Z operon and cause cells to produce the galactosidase. Then you add in X-Gal and just pick out the bacteria that have functional Lac Z because the useful cells will be a bright blue!

Check out the figure below:

Transformation in Bacteria with LacZ

Transformation leads to Competent cells with LacZ operon

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

Bacterial Transformation Protocol

Transformation describes the uptake and incorporation of plasmid DNA into bacteria. Antibiotic resistance genes carried on plasmids allow selection of transformants. This protocol describes the transformation of DH5α E. coli with pAdtrackCMV (a vector carrying kanamycin resistance).

Materials for Bacterial Transformation

Ligation mix (20 µl) – insert ligated into pAdTrack-CMV shuttle vector (Plasmid #16405, Addgene)
DH5α competent cells (includes pUC19 DNA control; #18265017, ThermoFisher Scientific)
LB broth (#10855-021, ThermoFisher Scientific)
LB Agar selective plates (prepare from #22700025, ThermoFisher Scientific) with 50 µg/ml kanamycin (#15160054, ThermoFisher Scientific)

Step by Step Transformation Protocol

  1. Thaw competent cells on ice. Aliquot 50 µl into cooled Eppendorf tubes for each transformation reaction.*
  2. Add 5 µl of ligation mix to each tube.*
  3. Incubate on ice for 30 min.
  4. Heat-shock the cells for 20 sec in a 42°C waterbath.
  5. Place on ice for 2 min.
  6. Add 950 µl of warm LB broth per tube.
  7. Allow cells to recover at 37°C for 1 hour with gentle shaking.
  8. Spread 200 µl and 20 µl of each transformation mix onto warm selective plates.*
  9. Incubate plates overnight at 37°C.

Notes on this methodology

  • We will talk about “Ligation” in another future blog post
  • Step 1. Unused cells can be refrozen and stored at -80°C for future use.
  • Step 2. As a transformation control, add 1 µl of pUC19 plasmid to one aliquot of cells (pUC19 confers resistance to ampicillin so will need to be seeded onto different selective plates).
  • Step 8. Transformation mix can be stored at 4°C and plated the next day if required.

Bacterial Transformation Video Tutorial

Applications of DNA Transformation on Scigine


Excellent Book about Bacterial Transformation
Guide to Common terms in Transformation – Oklahoma University
Compilation of History of Transformation and related protocols
He et al Proc Natl Acad Sci U S A. 1998 Mar 3. 95(5):2509-14.

Guide: Measure Cell Proliferation with Thymidine and BrdU

Cell Proliferation, BrdU, Thymidine, EdU

Thymidine and BrdU, Cell Proliferation Assay Summary

How do you know if your cultured cells are growing? Does your new cancer drug affect cell proliferation? What’s the effect of VEGF on endothelial cells? As you can tell, knowing how to perform cell proliferation assays is an absolutely essential skill for anyone in biology, biochemistry, or pharmaceutics. Radioactive Thymidine cell proliferation assays have been used since for over 40 years to detect whether cells are growing. The principle is simple: cells will incorporate Thymidine into their DNA as they proliferate. However, dealing with radioactivity is painful and annoying, so new fluorescence-based, non-radioactive, BrdU and EdU cell proliferation assays have become the new mainstay technique. These molecules are both thymidine analogs and hence work using the same principle as radioactive thymidine. In today’s guide, we will learn Step-by-Step, the theory behind these assays and how to apply them in the lab. Combining our techniques of MTT Cell Viability assays and Flow Cytometry or FACS, we are really building up a great list of skills to analyze biological phenomena!

Note: If you’re writing research papers, I highly recommend Grammarly – it’s a free grammar check plugin for Chrome. Try it out here

Principle of Cell proliferation assays with nucleotide analogs

3H-Thymidine is a radioactive version of the Thymine DNA base (thymine + the sugar backbone = thymidine). When cells are incubated with thymidine, they use the radiolabeled thymidine to synthesize DNA and incorporate it into their DNA backbone. So, thymidine is an excellent measure of DNA synthesis in cells that have undergone the S-Phase of cell replication. Similarly, BrdU is a Thymidine analog that lacks the radioactivity from tritium and it is used identically to Thymidine. Just incubate cells in the presence of BrdU. However, unlike radioactive thymidine, BrdU is detected with Anti-BrdU antibodies.

A quick summary picture is shown below.
Cell Proliferation with Tritium Thymidine
BrdU Cell Proliferation no radioactivity, Thymidine has it.

Using Thymidine vs. BrdU. Cell Proliferation Assay Tips and Tricks

Taking things with a grain of salt: Note that DNA replication can happen even when cells are not proliferating. For example, if you have damaged DNA (ie. DNA repair is taking place). So, Thymidine and BrdU assays are really DNA replication assays and not perfect cell proliferation assays. But, for the most part, they are the gold-standard when looking for cell proliferation.

Thick tissue sections? Choose your cell proliferation assay wisely: The 3H-Thymidine assay uses radioactivity. And the beta particles that are generated by this method cannot penetrate very deep into tissue. So, if you’re labeling tissue sections, make sure they are extremely thin! In these cases, BrdU is a great option because it penetrates deep into tissue and can be detected even from 50 um thick slices. This is illustrated below in the picture.

Not enough signal? Add more: Since cells are substituting the radioactive thymidine into their DNA. Adding more thymidine means you’ll get more incorporation. And, more incorporation means you’ll get more signal! So, if your signal is low, just add more of the nucleotide. BrdU, however, doesn’t behave this way. There is a limit at which adding BrdU doesn’t increase your signal.

Want to preserve your tissue? Use 3H-Thymidine: BrdU immunohistochemistry requires you to digest and disrupt tissue for visualization because the antibodies that detect BrdU need to access all the tissue. Detecting radioactivity just needs you to use a scintillation counter.

Thymidine Cell Proliferation Assay vs BrdU Assay

Other Methods: EdU Cell Proliferation Assay

BrdU immunohistochemistry has the disadvantage that you need antibodies to detect it. Because of this, you need to disrupt the tissue you are staining. EdU is a new version of BrdU which has an azide functional group. This can then easily be detected with a “click” fluorophore. This is shown below:

EdU Fluorescence Cell Proliferation Assay
EdU vs. BrdU Cell Proliferation Assay

Step by Step Guide to BrdU Cell Proliferation assay in vivo

Here is how you can label proliferating cells in a mouse
Materials for BrdU Assay
BrdU labeling reagent (Invitrogen, #000103)
BrdU staining kit (Invitrogen, #933943)
Opaque dark container for storing stained tissue
16% Paraformaldehyde in Water (#47608, Sigma Aldrich)
4% Paraformaldehyde
1% Paraformaldehyde with 0.5 M EDTA, pH 8 [Demineralization solution]
Histo-clear (#50-329-51 Fisher Scientific)
Ethanol 100%
Ethanol 95%
Ethanol 70%
30% Hydrogen Peroxide (H2O2) in Water
Petri dish with wet paper towels and paperclips for humidifying tissue (see image)
Humidifying chamber for Cell Proliferation Assay

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

Step-by-Step Protocol for Cell Proliferation Assay
You’ll want to refer to our Immunofluorescence microscopy guide and our Immunoprecipitation / CoIP Guide to understand this section better. Especially the bit about biotinylated antibodies and their detection.

  1. Give mice 100 ul of the BrdU labeling agent per 10 g of mouse weight. I.P
  2. Wait 2-4 hours for labeling the skeletal tissue
  3. Asphyxiate mice using CO2
  4. Dissect mouse body parts and rinse with 1x PBS
  5. Fix mouse tissue in 4% paraformaldehyde solution for 48 h at 4oC
  6. Demineralize tissue with demineralization solution for 3 weeks. Change solution 1x per day, everyday.
  7. Rinse sample tissues 3x with PBS for 60 minutes each wash.
  8. Embed the tissues in paraffin and mount on slides. Make sure sections are less than 20 um thick.
  9. Air dry the frozen sections on the bench for 1 h.
  10. Remove paraffin by dipping the slides in Histo-Clear and leaving them for 5 minutes. Repeat this once again.
  11. Rehydrate sections by dipping in 100% Ethanol for 2 min, then 95% for 2 minutes, then 70% for 2 minutes
  12. Wash sections 3x with 1x PBS for 2 min at a time.
  13. Quench endogenous peroxidase activity by submerging sections in 10% H2O2 in methanol for 10 min
  14. Rinse 3x with PBS for 2 min each.
  15. Make trypsin solution according to BrdU Staining kit and cover tissue sections.
  16. Rinse with PBS 3x for 2 min each.
  17. Add DNA denaturing solution and cover tissue for 30 min.
  18. Rinse off excess with PBS wash 3x for 2 min each. Then remove the PBS by blotting with tissue paper around the edges of the tissue.
  19. Add the blocking solution and submerge tissue.
  20. Add the biotinylated mouse anti-BrdU antibody and incubate for 60 min in the humid chamber petri dish.
  21. Rinse 3x with PBS
  22. Add the streptavidin-peroxidase solution. Incubate for 30 min.
  23. Rinse 3x with PBS
  24. Freshly make the peroxidase staining solution from the BrdU staining kit. Incubate sections for 5 min in this.
  25. Add hematoxylin staining solution for 1 min. Not more!.
  26. Quickly rinse with PBS 2 times for 30 secs.
  27. Dehydrate slides by incubating in 70% ethanol for 1 min. Then incubate in 95% ethanol for 1 min. Then incubate in 100% ethanol.
  28. Add Histomount media and put a coverslip on top.
  29. Use a normal brightfield microscope to image the cells. Make sure to check multiple vieweing areas to get a representative sample of your tissue.

Notes on this BrdU Cell Proliferation Assay Methodology

  • Demineralization finishes when the tissue is pliable and can be bent without breaking.
  • Embedding tissues in Paraffin will be discussed later in another post.
  • After adding peroxidase staining solution, the tissue will become visibly browner
  • Staining with hematoxylin will make sections too blue and harder to analyze. Don’t leave in there for more than 1 minute!
  • Normal cells that are dividing will have brown nuclei. Non dividing cells will have blue nuclei. % Proliferation = dividing cells / (dividing cells+non dividing cells).

Checking Cell Proliferation in Zebrafish: Video

Applications of BrdU, EdU, and Thymidine Assays on SciGine

BrdU Immunofluorescence Cell Proliferation Assay
In Vivo BrdU Assay for Cell Proliferation of Chondrocytes
Thymocyte proliferation measured using [H]Thymidine
EdU Cell Proliferation Assay with Flow Cytometry
Lamprey Cell Proliferation using EdU Assay


Calculation of Cell Proliferation using Thymidine
Duque et al.: How Proliferation assays affect cell behavior
Mead et al.: How to use BrdU with Skeletal tissue sections