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.

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

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

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    • 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.

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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

Calcium Phosphate Transient Transfection Protocol & Guide

Calcium phosphate Transient Transfection Protocol and Guide

Transfection with Calcium Phosphate: General Summary

Molecular biology tools allow us to understand and manipulate DNA/RNA so that we can change how cells behave. In this next series of posts, let’s learn how to manipulate cells and make them do our bidding. Among the list of methods to learn, the first tool to understand is transfection – the process by which we introduce new DNA into a cell so that we can change what proteins it creates. Specifically, we will focus on Calcium Phosphate transient transfection because it is a common and powerful technique. We can then combine transfection with some of our protein-manipulation techniques to change cell behavior and confirm that our changes actually had an effect: Immunoprecipitation (IP) and Western Blotting. Note that other techniques for transfection including electroporation, DEAE:Dextran based transfection, and lipid mediated transfection will be discussed in the future.

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Transient vs. Stable Transfection

When you introduce DNA into a cell, it is possible for the cell to keep the DNA temporarily or permanently. Temporarily, a cell might keep your DNA as a packaged plasmid and express it until it divides. Permanent transfection takes place when the new DNA is integrated into the genome of the cell and it passes the DNA down through cell division into its progeny. It’s difficult to determine when genes will be integrated into the genome (stable transfection) and when they will be kept temporarily (transient transition). There is a lot of luck involved. However, it is possible to only keep cells that have your DNA by selection. Take a look at the image below:

Calcium phosphate Mammalian Transfection

DNA Transfection guide

Transient Transfection vs Stable transfection

Calcium Phosphate Transient Transfection

To introduce DNA into eukaryotic cells such as mammalian cells, we need to neutralize the charge on the DNA. This is because cell surfaces are negatively charged and DNA that is unshielded will be repelled from the cell surface. Even if some DNA enters a cell, the nuclear envelope will also reject the DNA due to its charge. (For a picture of the DNA polymer look at our PCR protocol) So, the classical technique for neutralization of DNA’s charge is to use Calcium Phosphate. The steps for transfection with Calcium Phophate are very straight forward:

  • Generate DNA strand (circular DNA is much easier to introduce)
  • Mix calcium phosphate with DNA and generate nanoparticulate precipitates
  • Incubate with cells
  • Select cells expressing the DNA of interest

Cells will tend to phagocytose the calcium phosphate nanoparticles and, with luck, they will allow the nanoparticles to enter the nucleus. Calcium phosphate transfection works well because of the stability provided by divalent calcium ions. Other methods such as lipofectamine and polyethylene imine based transfection also work similarly by neutralizing DNA’s charge. But lipids offer the additional benefit of making the DNA complex more hydrophobic and hence make it easier for it to pass through the phospholipid bilayer.

The general technique is shown below:

Calcium phosphate Nanoparticles and Aggregates

Phagocytosis of Calcium Phosphate leads to Transfection

Selection Media Confirms Stable Transfection with Calcium Phosphate Protocol

Tips and Tricks when optimizing your Calcium Phosphate Transient Transfection Protocol

Calcium Phosphate based transfection is a standard and well known technique. Calcium divalent ions bind the DNA polymers and neutralize their negatively charged phosphate backbones. However, optimization is necessary to maximize the DNA that is phagocytosed into your cell of choice. The variables that affect this technique’s efficacy are:

  • The pH of the solution: Even differences of 0.1 units will have drastic effects on the efficacy of your transfection protocol.
  • Amount of DNA in the precipitate:Some cell types require a lot of DNA in the precipitate such as primary human foreskin fibroblasts. Others will tend to die instead of uptaking DNA, if you add too much DNA.
  • Incubation time with the precipitate:HeLa and 3T3 cells are relatively easy to transfect within 16 hours. These cells can tolerate DNA nanoparticles for extended periods of time. However, other cell types may need shorter incubation times and may tend to apoptose if exposed too long.
  • Additional glycerol or DMSO shock: It may be useful to “shock” cells with a 10% Glycerol solution or a 10-20% DMSO solution for a short time (~3 minutes). Carefully optimize this time for your particular cell type.
  • Rate of Formation of DNA nanoparticles: High efficiency transfection techniques have been discovered whereby buffers like BBS allow DNA nanoparticles to form slowly and precipitate onto cells. When this happens, cells tend to phagocytose more of the adducts and tend towards higher viability/less toxicity.

To make sure that your DNA is being incorporated into cells, you should include a reporter plasmid such as one with neomycin resistance (neo). You will need to optimize the ratio of neo reporter DNA vs. the DNA you want to include.

Calcium Phosphate Transient Transfection Protocol

Materials for Calcium Phosphate Transfection
HeLa cells
Complete DMEM
DNA (10 – 50 ug per transfection)
2.5 M CaCl2 (#C3306 Sigma Aldrich)
2x Hepes Buffered Saline (0.28 M NaCl, 0.05 M HEPES [#H3375 Sigma Aldrich], 1.5 mM Na2HPO4, pH 7.05 exactly)
Culture Dish

Materials for BBS Calcium Phosphate Transfection
HeLa cells
Complete DMEM
TE buffer, pH 7.4 (10 mM Tris-Cl, 1 mM EDTA)
2.5 M CaCl2
2x BES-Buffer (BBS) (50 mM BES [#B9879 Sigma Aldrich], 280 mM NaCl, 1.5 mM Na2HPO4 pH 6.95 exactly)
Selection Medium

Transfection Protocol Steps

  1. Split cells such that there is space between cells.
  2. Clean DNA by adding in 100% ethanol for precipitation
  3. Dry DNA after aspirating supernatant from ethanol. Use air to make sure it is completely dry.
  4. Resuspend pellet in 450 ul of water with 50 ul of 2.5 M CaCl2 buffer
  5. Put 500 ul of 2x Hepes Buffered Saline in a 15 ml conical falcon tube
  6. Add the DNA/CaCl2 solution dropwise to this tube while agitating with a stir bar or other mechanism.
  7. Allow the precipitate to sit at room temperature for 20 minutes
  8. Spread the precipitate over the cells along with their medium . Shake gently to make sure the precipitate is even.
  9. Incubate in cell culture incubator at 37 oC with 5% CO2 for up to 16 hours
  10. Remove medium, wash twice with PBS, and feed cells with complete medium.
  11. Plate cells in selective medium.

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

BBS High Efficiency Transfection Protocol Steps

  1. Seed cells in a dish so that they can double atleast twice so they can be stably transfected
  2. Next day, dilute DNA in TE Buffer at 1 ug/ul
  3. Make a 0.25 M CaCl2 stock
  4. Mix 20-30 ug of DNA with 500 ul of 0.25 M CaCl2 stock.
  5. Add 500 ul of BBS to this mixture and vortex. Incubate at room temp for 20 min.
  6. Add this mixture to the cell culture dish dropwise and mix by gently shaking the plate.
  7. Incubate cells overnight for 24 h at 3% CO2 at 35 oC.
  8. Wash cells 2x with PBS and then incubate them in complete medium for 2 doublings.
  9. Split cells and incubate in selection media.

Notes on this transfection methodology

  • Cell density has to be low but not too low. The ideal cell density allows you to reach confluence at the end of the transfection period without making the media acidic.
  • Also, space between cells increases transfection efficiency because DNA phagocytosis is proportional to exposed surface area of cells.
  • For some cells, incubate with 10% glycerol or 10-20% DMSO for 3 minutes, and wash twice with PBS, prior to adding the DNA nanoparticles. This may increase your transfection efficiency. However, for the BBS method, this step is not necessary because it will not affect cell transfection efficiency.
  • Supercoiled DNA and plasmid DNA works best with these procedures.
  • Depending on the plasmids that you introduce into cells, your selective medium will vary.
  • pH is EXTREMELY CRITICAL for transfection procedures. At the end of the transfection, pH of your medium should be alkaline at 7.6, but prior to the procedure, make sure all your buffers are clean and at the right pH.
  • All buffers above may be frozen and stored as aliquots. But it is important to make sure that your pH is correct prior to using freshly thawed buffers.

Applications of Transfection on SciGine

Murine L cells transfected with Calcium Phosphate and BBS Buffer
HeLa cell transfection with Lipopolyamine
Calcium Phosphate with HEPG2 and HEK293
MDAMB436 cells with Calcium Phosphate Transient Transfection
Transient Transfection Protocol for HEK293T cells

Transient and Stable Transfection Video Tutorial


Calcium Phosphate Transfection by Kingston et al.
High Efficiency Transfection by Chen et al.
Transfection Review by Kim et al.

Immunofluorescence Microscopy Protocol and Method Guide

Immunofluorescence Microscopy Protocol and Method Guide

Immunofluorescence Microscopy Overview & Theory

Immunofluorescence Microscopy (IF) is a classical technique to observe the localization of molecules in cell/tissue sections. While most researchers try to look for proteins, it is also possible to look for DNA, RNA, and carbohydrates in sections of tissue. With the addition of this technique to your tool belt along with the immunoprecipitation scientific method and the western blot scientific method, you will now have a variety of different ways of manipulating and analyzing biomolecules from tissues, cells, and cell lysates. The principle is fairly straight forward: incubate your sample with an antibody generated towards your target molecule and then detect the antibody using fluorescence. In an immunofluorescence microscopy experiment, this takes the form of putting your cells on a microscope slide, probing with antibodies, and then using specialized fluorescence microscopes with red/blue/green filters along with specific laser emitters to visualize the antibodies. As you would expect, you could either incubate your samples with a antibody-fluorophore conjugate and visualize it (direct immunofluorescence), or you could first put in an antibody that recognizes your target and then put in a secondary antibody-fluorophore conjugate that recognizes the first antibody (indirect immunofluorescence).

Related articles:

Here’s an image which describes the above theory:
Immunofluorescence Microscopy Detect Proteins in Cells
Direct and Indirect Immunofluorescence
Microscopy for localization of DNA and RNA binding proteins

Advantages and Disadvantages of Direct vs. Indirect Immunofluorescence Microscopy

The most common method of performing an IF experiment is to use the indirect immunofluorescence technique. But both methods have their merits, and depending on your application, you may be limited to one method over another.

Direct Immunofluorescence microscopy:
Fewer steps: You don’t need to use a secondary antibody, so you have fewer wash and other steps.
Fewer Complications: Fewer steps also means less troubleshooting.
More expensive: Primary antibodies that are specific and also conjugated to fluorophores are hard to find and expensive.
Lower overall signal: Each primary antibody only has one fluorophore, so you have lower overall signal.

Indirect Immunofluorescence microscopy:
Higher signal: A single primary antibody may bind to multiple secondary antibodies (which each have a fluorophore)–so you get higher overall signal because of this amplification effect.
More versatile: Multiple types of secondary antibodies may be used to detect a single primary antibody. This allows you to probe the same sample multiple times, get higher signal, use a single secondary to detect multiple primaries, and various other strategies.
Cheaper: Secondary antibodies raised against a certain species of primary are very cheap and widely available.
More steps: With a secondary antibody incubation step, there is a possibility for more complications and troubleshooting.

Dealing with the weaknesses of Fluorophores in Immunofluorescence Microscopy

[Slightly technical] Fluorescence is a phenomenon whereby an electron receives some light energy, gets temporarily promoted to a higher energy orbital, and then relaxes back down to its baseline energy state. As the electron relaxes, it releases the light at a slightly lower energy that the initial incident light which hit it. [End Technical Section]

How Fluorescence works:

Immunofluorescence Microscopy Theory of Fluorescence and Electron Energy

This leads to some challenges when performing IF experiments: photobleaching, autofluorescence, and non-specific fluorescence.

Photobleaching during Immunofluorescence Microscopy and how to deal with it:
Use less energy: Some fluorophores will decompose after receiving a lot of energy from lasers. This is typically seen in microscopy when your sample slowly become dimmer and dimmer after imaging a section too long. To overcome this challenge you should use a lower energy intensity when looking around for the right locations in your sample, and then switch to a higher energy intensity when taking an image of your sample. Typically, good microscopy systems will take care of this automatically.

Antifade agents: these molecules scavenge singlet oxygen radicals that are caused by high energy lasers and can be used to maintain high fluorescence signals during microscopy. It is theorized that singlet oxygen species are the main culprit that cause localized damage to fluorophores.

High yield fluorophores: With fewer high-yield fluorophores you can get more signal. The “quantum yield” is an important number when considering which fluorophore you should use alongside your antibodies. If you can find quantum-dot conjugated antibodies, you’re going to be amazed with your microscope images! 🙂 I’ll write more about them in the future.

Cellular Autofluorescence:
There are a few molecules within cells that cause auto-fluorescence. Auto-fluorescence can lead to false positive data in flow cytometry experiments and rarely even in immunofluorescence microscopy. Be wary of the following molecules:

  • FAD and/or FMN: Flavinoids such as these have an Excitation @ 450 nm and Emission @ 515 nm
  • NADH: An Adenine dinucleotide with Excitation @ 340 nm and Emission @ 560 nm

FAD and NADH structures lead to auto-fluorescence:
Immunofluorescence Photobleaching and Autofluorescence

To avoid autofluorescence, use fluorophores with excitations and emissions far away from the above wavelengths so that the inherent fluorescence in your sample doesn’t affect your measured signal.

Non-specific Fluorescence:
Based on the excitation and emission of different fluorophores it is possible for you to get fluorescence signals even though you haven’t probed for them. Be wary of the different fluorophores that you use in your sample and the bandwidths with which you detect them. You may need to modify your microscope’s settings and filters to get really sharp and beautiful IF images.

To avoid non-specific fluorescence, consult a fluorophore chart as mentioned in the guide for the Flow Cytometry (FACS) scientific method.

Why do we have non-specific fluorescence? Take a look at this image:

Immunofluorescence Microscopy Non-specific Fluorescence
Overlapping fluorescence spectra cause problems with Microscopy
Use appropriate and compatible fluorophores

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

Immunofluorescence Microscopy Step-By-Step Guide

Example Immunofluorescence of Pancreatic Sections

This protocol describes the detection of insulin in paraffin wax-embedded sections of pancreatic tissue.

Normal donkey serum (#D9663 Sigma)
Guinea pig anti-insulin antibody (used at 1:3200; #ab7842 Abcam)
Biotinylated donkey anti-guinea pig antibody (use at 10 µg/ml; ##706-066-148)
Streptavidin conjugated Cy3 (use at 1:100; #016-160-084 Jackson Laboratories)
Proteinase K buffer (50 mM Tris pH 8.3, 3 mM CaCl2, 50% glycerol)
Proteinase K (20 mg/ml; #AM2548 ThermoFisher Scientific)
DAPI (#D1306 ThermoFisher Scientific)
SlowFade mounting media (#S36937 ThermoFisher Scientific)

Experimental procedure:

  1. Cut 8 µm paraffin sections at the microtome and mount onto glass slides.
  2. Heat slides at 60°C for at least 5 min to melt wax.
  3. Dewax slides in xylene for 2x 5min*, 100% ethanol for 2x 5min and immerse in running deionised water until clear.
  4. Circle sections using a wax stick to easily observe staining area in subsequent steps.
  5. Prepare Proteinase K by adding 1 µl to 1 ml warm (37°C) Proteinase K buffer. Perform antigen retrieval by adding 50 µl of diluted Proteinase K per section in a humidity chamber for 20 min at 37°C*. Wash slides 3x 5 min in PBS with stirring.
  6. Block sections with 50 µl of 10% normal donkey serum in PBS at room temperature for 30 min in humidity chamber.
  7. Tip off blocking solution and add 50 µl anti-insulin primary antibody diluted in 10% normal donkey serum in PBS. Incubate overnight at room temperature in humidity chamber.
  8. Wash slides 3x 5 min in PBS with stirring. Add 50 µl anti-guinea pig biotinylated secondary antibody diluted in 10% normal donkey serum in PBS. Incubate 2 h at room temperature in humidity chamber.
  9. Wash slides 3x 5 min in PBS with stirring. Add 50 µl streptavidin-Cy3 diluted in 10% normal donkey serum in PBS. Incubate 1 h at room temperature in humidity chamber in the dark.*
  10. Wash slides 3x 5 min in PBS with stirring. Add 50 µl DAPI (3 µM) diluted in 10% normal donkey serum in PBS to stain cell nuclei. Incubate 30 min at room temperature in humidity chamber.
  11. Tip off excess solution from slide and mount in one drop of mounting medium with coverslip.
  12. Examine at the fluorescence microscope*.

Procedural notes

  • Step 3. Perform in fume hood
  • Step 5. Perform incubations in humidity chamber to reduce evaporation of antibody/proteinase solutions
  • Step 9. Perform incubations with fluorescently labelled reagents in the dark
  • Step 12. Slides can be stored at 4°C until use
  • Consider buying “pre-adsorbed” secondary antibodies. These antibodies have been incubated with common cross-reacting species and they didn’t bind. Therefore, they are super specific to your species of choice and should provide very little background signal.

SciGine Immunoprecipitation Microscopy Protocols and Methods

Immunofluorescence of Arp3
Immunofluorescence Microscopy for MUC of Methanol Fixed Tissue
Use of Cy3 conjugated antibody for Immunofluorescence in Cos7 cells
Cells permeabilized with TritonX-100 for use in Immunofluorescence Microscopy
Analysis of Epithelial Polarization using Indirect-IF


Discussion of Immunofluorescence by Robinson et al.
Robertson et al. IF guide on Biomed Central
NADH and FAD autofluorescence

Immunofluorescence Theory Video by the HHMI

Immunoprecipitation (IP) Scientific Method Guide

Immunoprecipitation Method Guide on SciGine

Immunoprecipitation Overview

Immunoprecipitation is a method for extracting protein from a solution. Typically, this solution is a cell lysate which you want to analyze. Very frequently you’ll hear your colleagues say, «I’m going to do an IP-pull down on my protein» — when you hear this, you’ll know they’re talking about immunoprecipitation. This technique is a must-have for any biochemist who works with proteins because it’s so versatile. Once you remove a protein from solution, you can analyze it to see what it binds. You can also check out its molecular weight and structure. And, beyond proteins, IP can be applied to RNA and DNA pull downs as well.

In theory, this method is very simple. First, lyse your cells using some sort of lysis buffer. Next, add in an antibody that will bind your protein of interest and form a protein-antibody complex. Then drop in some resin which can bind the antibody. Typically, this resin is either agarose-based or superparamagnetic and is covered with Protein A and/or Protein G. These proteins are specifically designed to bind the heavy chains of antibodies so they can easily pull out your protein-antibody complexes. Agarose beads offer higher capacity per bead but magnetic beads are MUCH easier to separate because you can use a magnet to keep them in place. Finally, spin everything down and remove the supernatant. With the remaining bead-antibody-protein conjugates, you can either denature everything and run a SDS-PAGE western blot, or you can try to analyze your protein’s function with an activity assay, or run it on HPLC or LC-MS, etc. There are so many downstream applications of immunoprecipitation!

Take a look at the attached drawing to understand this method:
Immunoprecipitation Scientific Method
Immunoprecipitation Pulldown Assay
Immunoprecipitation IP Downstream applications

Related articles:

Different Types of Immunoprecipitation (IP) / Pull Down Assays

While the previous description shows the most straight forward and common version of immunoprecipitation, there are many variants of this method. It’s truly versatile and powerful, so let’s take a look:

  • Pre-immobilized Antibody Immunoprecipitation: Instead of adding in antibody for your target and then immobilizing the antibody onto agarose beads, you can pre-immobilize it and then add it into your protein mixture.
  • Co-IP, CoIP, or Co-Immunoprecipitation: In a Co-IP, you pull down multiple proteins along with your protein of interest as a complex. It’s very likely that this protein complex that is physiologically relevant to the signaling cascade that they trigger. As an example, you could pull out clathrin-binding proteins and you’d have a clue as to how these proteins enter/exit cells. Cool right? With this simple technique, you can now map out which proteins interact with each other to cause a certain phenotype!
  • ChIP or Chromatin Immunoprecipitation: In this method, the DNA that is bound to your target protein is what you are after. The idea is to first let your protein bind DNA. Then you crosslink it using formaldehyde and lyse the cells. Next, you fragment the DNA so your protein isn’t bound to ALL the cellular DNA…it’s only bound to the DNA that it interacts with. Then do a regular IP based on this blog post. With this precipitated protein, you can then detach the protein from the DNA using heat and perform PCR to look at what DNA segment is being bound.
  • RNA Immunoprecipitation or RIP: Jut like a ChIP, you can also pull out proteins that bind to RNA inside cells. The strategy is the same as ChIP but you need to use RT-PCR to analyze the RNA that you get. We’ll discuss this technique in the future.
  • Biotin/Streptavidin Immunoprecipitation: Some times, Protein A and/or G, based pull-downs don’t work because your antibody doesn’t bind to them very well (due to protein-to-protein variability). In these cases, you need an alternate strategy. If you can find biotinylated antibodies for your target protein, then you can use streptavidin beads to pull them out! Biotin-Streptavidin interactions are among the strongest molecular interactions known to science.
  • Covalent Capture Immunoprecipitation: If you cannot use Protein A and/or G, and you cannot find the biotinylated antibodies for your target protein, then you have to go back to chemistry. The strategy is relatively simple, but most biologists hate chemistry 🙂 All you need to do is to oxidize your antibody to introduce aldehydes in the backbone. This can be done using periodate oxidation on vicinal diols in the heavy-chains of the antibody. Then using amine-containing resins, you can capture the antibodies via schiff base formation and borohydride reduction. You can also flip this strategy on it’s head and use amine-containing antibodies with aldehyde containing resin.

Some of these strategies are shown here:
Preimmobilized antibody or Coip used in IP Pulldowns
Co immunoprecipitation and Chromatin immunoprecipitation
Steps of Chromatin Immunoprecipitation
Finding DNA binding regions of proteins using ChIP

Immunoprecipitation Scientific Method Step-By-Step

Immunoprecipitation of the Lck protein from T cell lysates has been used to analyse the phosphorylation patterns of this important T cell activation protein.

IP Materials:

Lysis buffer (20 mM Tris pH 7.5, 150 mM NaCl, 1% Nonidet P-40, 1 mM phenylmethane sulfonyl fluoride (PMSF))
Protein G-Sepharose beads (Protein G Sepharose 4 Fast Flow #17-0618-01, GE Healthcare Life Sciences)
Protease inhibitors (HaltTM Protease Inhibitor Cocktail 100X, #78430, ThermoFisher Scientific)
Purified mouse anti-human p56-Lck antibody (#551048, BD Biosciences)

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

IP Experimental procedure:

Perform all steps on ice to avoid protein degradation.

  1. Pellet cells (107) and resuspend in 800 µl ice-cold lysis buffer containing protease inhibitors. Lyse cells at 4°C for 15 min with mixing.*
  2. Pellet cell debris for 15 min at maximum speed on microcentrifuge at 4°C.
  3. Prepare Protein G-Sepharose beads following manufacturers’ instructions, by washing 1 ml of bead slurry three times in lysis buffer. Resuspend the final pellet in an equal volume of lysis buffer and store at 4°C until use.*
  4. Preclear lysate by adding 50 µl of washed Protein G-Sepharose beads at 4°C with mixing for 1 h. Pellet beads for 20 sec at maximum speed on microcentrifuge and retain lysate in a fresh tube.
  5. Add 1-2 µg of specific antibody (eg. anti-Lck) to lysate for 15 min at room temperature. Add 50 µl of washed Protein G-Sepharose beads and incubate overnight at 4°C with mixing.
  6. Spin for 20 sec at maximum speed on microcentrifuge to pellet immunoprecipitates. Wash complexes three times in 500 µl lysis buffer.*

IP Procedural notes:

  • Step 1. Prepare fresh lysis buffer on day of procedure. If protein is intended to be assayed for phosphorylation levels, also include phosphatase inhibitors.
  • Step 3. Protein G has a high affinity for mouse Ig; for other precipitating antibodies Protein A can be used.
  • Step 6. If samples are to be analysed by SDS-PAGE, add reducing sample buffer directly to washed immunoprecipitates prior to electrophoresis.
  • Protein A and/or G are not great way to immobilize antibodies if doing an IP with serum. Serum contains lots of other Protein A and/or G types which will compete with your IP resin.

SciGine Immunoprecipitation Applications

CDK2 Immunoprecipitation
Phosphoserine Pull-down from G418 clones
Immunoprecipitation and SDS-PAGE of FLAG protein
SRC Kinase Immunoprecipitation and SDS-PAGE
Immunoprecipitation of Rheumatoid Arthritis related RV202


Immunoprecipitation guide by Kaboord et. al
IP Protocol by Carey et. al
ChIP by the Haber Lab

Western Blot Theory and Method Guide

Western Blot Method Guide and Step by Step Procedure

Overview of Western Blot Method

A western blot enables sensitive detection of specific proteins from a solution containing multiple proteins. This is an essential biology technique and one of the cheapest methods that can be utilized to analyze proteins. To perform a western blot first separate proteins based on their mass and charge via gel electrophoresis, and then follow up by detecting the protein of choice with a specific antibody. Typically, researchers will use western blots to separate proteins from cell media or from cell lysates. For example, if you wanted to find out how much actin your cells are expressing, a western blot can easily compare actin amounts between different cell types. It’s also likely that you will be using western blots when producing proteins in mammalian and insect cells.

In a typical western blot procedure, cells will first be lysed and the amount of protein will be determined using a spectrophotometer. Then a gel will be made and the total protein from the cell lysate will be loaded into wells in the gel. After applying an electrical field, the proteins in the gel will begin to migrate down and separate into distinct bands based on the size and charge of the protein. After the smallest proteins reach the bottom of the gel, the electrophoresis will be stopped and all proteins on the gel will be transferred onto blotting paper so that they can easily be handled. Finally, antibodies that recognize the proteins of interest will be added and detected via chemiluminescence.

Related articles:

Here is a step by step illustration of how to perform a western blot:

Western Blot Scientific Method Guide

Western Blot Step By Step

SDS-PAGE Western Blot Step-by-Step Protocol

Western blotting can be used to examine the upregulation of RCAN1, a signaling molecule in neuronal cell types.

Materials for Western Blots:

  • Rabbit anti-RCAN1 antibody (#SAB2101967, Sigma-Aldrich)
  • SDS-PAGE gel (Criterion TGX precast Stain-free Any kD gel, #5678124, Bio-Rad)
  • TBS (20 mM TrisCl pH 7.6, 150 mM NaCl)
  • Running buffer (25 mM Tris, 192 mM glycine, 0.1% SDS)
  • Transfer buffer (25 mM Tris, 192 mM glycine, 20% methanol, 0.05% SDS)
  • 4X SDS-PAGE loading buffer (Laemmli’s sample buffer #1610747 Bio-Rad; add fresh dithiothreitol to 10 mg/ml on the day of experiment)
  • Transfer membrane (0.2 um polyvinylidene fluoride membrane, #03010040001 Roche)
  • Secondary antibody (donkey anti-rabbit horseradish peroxidase-conjugate; #711-035-152 Jackson ImmunoResearch)
  • Blocking buffer (5% skim milk powder in TBS with 0.1% Tween20)
  • Immun-Star WesternC Chemiluminescence reaction solutions (#170-5070 Bio-Rad)
  • Dual color ladder (#1610374 Bio-Rad)
  • Blotting paper (ProteanXL, #1703966 Bio-Rad)

Western Blot Experimental procedure:

  1. Unwrap precast gel and rinse wells three times with running buffer. Assemble gel in tank and fill with running buffer.*
  2. In an Eppendorf tube add protein sample (30 µg) to 10 µl 4X SDS-PAGE loading buffer and add water to a final volume of 40 µl.
  3. Heat samples to 95°C for 2 min and spin briefly to ensure contents are at the bottom of the tube
  4. Load gel with samples and include ladder in one lane.
  5. Run gel at 200V for 30 min.
  6. While gel is running, soak two pieces of blotting paper (cut to the same size as the gel) in transfer buffer (approx. 30 min). Activate transfer membrane (also cut to size) by dipping in methanol, then soak in transfer buffer for approx. 10 min.
  7. Remove gel from tank and place in transfer buffer.
  8. Assemble transfer “sandwich” by placing down soaked blotting paper, transfer membrane, gel and blotting paper onto open transfer cassette (Turbo Blot transfer unit; Bio-Rad). Use a glass rod to roll across the “sandwich” to remove any air bubbles.
  9. Close cassette and run in machine (standard minigel program for 30 min).
  10. Remove transfer membrane from cassette, taking care to snip one corner to ensure orientation.*
  11. Incubate transfer membrane in blocking buffer for 1 h at 4°C with rocking.
  12. Pour off blocking buffer and add diluted anti-RCAN1 antibody 1:200 in 5 ml TBS with 2.5% skim milk power and 0.05% Tween20. Incubate overnight at 4C with rocking.
  13. Wash membrane three times in TBS with 0.2% Tween20 at 4°C with rocking, for 10 min each time.
  14. Incubate with diluted secondary antibody 1:2500 in 5 ml TBS with 0.2% Tween20 at 4°C with rocking for 1 h.
  15. Wash membrane three times in TBS with 0.2% Tween20 at 4°C with rocking, for 10 min each time. Rinse membrane briefly in water.
  16. Mix 1 ml each of ECL reagents in a foil-wrapped tube and add to membrane for 5 min prior to imaging on ChemiDoc MP imager (Bio-Rad).

Procedural notes for this Western Blot Method:

  • This precast gel contains 18 lanes with a loading capacity of 10-40 ug protein in up to 30 ul per well
  • Small needle-point markings can be added to membrane in-line with color markers which reduce in intensity following subsequent incubation and washing steps.
  • To make sure you know which step you are on, cut the bottom right side of your gel after running the gel electrophoresis.
  • In this method, the protein is denatured prior to running on the gel. This is called SDS-PAGE. By denaturing, you ensure that the size and charge are all that matter, as opposed to native gel electrophoresis where the conformation of the protein also matters.
  • Blots can be regenerated (the antibodies that were used for probing can be removed) by using stripping buffer. However, blots can only be stripped a few times before they have too much background noise to be easily analyzed.

Applications of Western Blots on Scigine (Search Engine for Scientific Methods):


NIH Western Blot Reference
Western Blotting by Kurien et al

Good Video References Related to Western Blots:

HPLC: Biochemical Analysis. A Step-By-Step Method Guide

HPLC Analysis Step by Step

HPLC Method Overview

HPLC, or high performance liquid chromatography is an amazing analytical technique for chemical compounds including biopolymers, small molecules, and polymers. In this method, a sample is first dissolved to make a solution. This solution is then injected into a “column” that contains resin that will interact with the sample. This will slow down the movement of the sample through the “column” and as the sample comes out the other side of the column, it is detected. This allows you to know both the time at which the sample comes out and the intensity of the sample that was detected. Here’s an overview of this technique:

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

HPLC Bioanalytical Method Guide

So, while there is continuous flow of some buffer through the column, we also inject our sample and observe as different molecules within the sample come out at different “retention times”. The detector on the end of the column can be any kind of detector but the most common types are refractive index (RI), ultraviolet (DAD), and fluorescence (FLD). Each of these will detect different properties of the molecules that come out of the column and display a chromatogram.

HPLC Chromatogram Guide

Types of Chromatography

Different column resin compositions determine the kind of chromatography that you are running and what molecules you can separate.

  • Normal Phase: The column is filled with silica particles which are polar and the buffer running through the system is non-polar. Once you inject your sample, polar particles will stick to the silica more and have a longer retention time than non-polar molecules.
  • Reverse Phase: The column is filled with hydrophobic particles (actually they are silica particles with long hydrocarbons on the surface). The buffer that is running through the system is polar (such as acetonitrile/water or methanol/water mixtures). This means that hydrophobic molecules will stick to the resin more and be retained longer.

Complete Step by Step HPLC guide


HPLC autosampler vialsI only use autosamplers since manual injection is tedious 🙂
Centrifugal filters with 0.2 um poresTo clean up samples
Eppendorf vialsFor centrifuging
HPLC machine


In a typical HPLC procedure you can decide the following variables:

VariableWhat it does
Flow rateWith fast flow peaks come out sooner but there’s they’re harder to resolve and tend to blend together. For more resolution, run slower.
PressureAffected by flow rate and solvent
Solvent BuffersDetermines signal intensity, how quickly the peaks come out, signal fidelity
Column TypeDetermines the type of interaction with the sample
Detection ParametersIf using UV or FLD, you need to set the right excitation/emission wavelengths

Since HPLC is a very machine-variable technique, I can only provide general guidelines.

For sample preparation:

  1. Dissolve your biopolymers or small molecules in a suitable solvent such as methanol
  2. Centrifuge at 10,000 rcf in an eppendorf vial and keep the supernatant to remove any large particular matter
  3. With a centrifugal filter, add 500 ul of your sample solution onto the top
  4. Centrifuge at 10,000 rcf and collect the filtrate (the solution that successfully passes through the filter)
  5. Load this sample into an HPLC vial

For setting up the HPLC machine:

  1. Make sure you have all your buffers set up
  2. Open the purge valve and purge the system for 5 minutes.
  3. Add your samples into the autosampler tray
  4. Stop the purge
  5. Close the purge valve
  6. Run the system at a normal flow rate (1 ml/min) with your buffer to equilibrate the column for 10 minutes
  7. Make sure that your pressure is stable (ie, less than 2-3 bar of fluctuation)
  8. Set up your sequence and your method
  9. Run a standard before your actual samples or as part of the same sequence

Example buffer system to determine Fluoresceinamine levels in samples:
Sample: Add 10 ug of fluoresceinamine into 1 ml of Acetonitrile.
Buffer: Pure acetonitrile buffer on a C-18 column; this is “reverse phase”.
Flow rate: 1 ml/min.
Column: 4.6 mm x 30 cm size.
Detection: Detect via a fluorescence detector set to Excitation @ 485 nm and Emission @ 535 nm.

Side-note: Use Grammarly to check your research papers – it’s a free grammar check plugin for Chrome. Try it out here

Notes on HPLC methodology

  • To clean the system and equilibriate it, you need to run enough solvent. However, this amount varies column-to- column. A typical 4.6 mm x 30 cm column should be clean when you follow the procedure above.
  • Isocratic means that the solvent concentration stays constant throughout the run.
  • It is useful to run standards before your samples as well as with your samples. Standards make it easy to identify which peak pertains to your molecule of interest.
  • Always use HPLC grade solvents. This is especially true for solvents like THF which are frequently sold with inhibitors that also complicate your ability to detect your molecule of interest.

Applications of HPLC on SciGine

HPLC is such a versatile technique. Take a look at these methods on SciGine which assay different types of chemicals in various samples.


ChemGuide Summary of Technique
Method Guide from Waters
Overview on Wiki

ELISA: A Step By Step Method Guide

Step by Step ELISA Guide on SciGine

ELISA: A Step By Step Method Guide

ELISA Biological Method Overview

ELISA is the common acronym for Enzyme-Linked-Immunosorbent Assay. It’s a quick plate based technique for detecting an antigen from a solution. This antigen could be a peptide, protein, antibody, or small molecule. In general, for an ELISA, an antigen is first immobilized on a surface (Step 1 below). Next, an antibody specific to the antigen is flowed over the surface (Step 2). This antibody, is also attached to a chemiluminescence-related enzyme. Treatment with the chemiluminescent substrate facilitates detection of the antibody and the antigen (Step 3). Take a look at these pictures to get an overview of the strategy:

Conjugated proteins may require different antibodies in ELISAs. However you can easily track the ELISA process by labeling your protein with a fluorescent probe.

ELISA Steps - SciGine Biological Methods

Types of ELISAs

There are a few different types of ELISA assays but they all follow the basic strategy outlined above.  Essentially, one can choose how to immobilize the antigen on the surface and how the antigen is  detected via the antibody.

  1. Direct Assay: In this method, the antigen is immobilized to the surface and detected directly via an  antibody that’s bound to a chemiluminescent enzyme. (Same as above)
  2. Indirect Assay: In this method, the detecting antibody doesn’t have the chemiluminescent  enzyme. So, another antibody must bind to the first antibody to facilitate detection.
  3. Sandwich Assay: The most common type of ELISA. In this assay, a “capture” antibody is first  immobilized to the substrate. Then antigen is flowed over it so that it gets immobilized to the  surface along with the capture antibody. Finally the detection antibody is flowed over the  substrate and it binds the antigen. This detection antibody may be directly conjugated to the  chemiluminescent enzyme (just like a direct assay) or another antibody may be needed (just like  the indirect assay).

Types of ELISA Assays - SciGine

Related articles:

A Complete Sandwich ELISA protocol

Materials for ELISA

96 well polystyrene plate

Plate shaker


Coating buffer

0.2 M sodium carbonate/bicarbonate buffer, pH 9.4

Wash buffer

0.1 M phosphate, 0.15 M sodium chloride, pH 7.2 with 0.05% Tween 20

Blocking buffer

2% w/v Bovine Serum Albumin in Wash Buffer

Diluent buffer

2% w/v BSA in Wash buffer or a more appropriate buffer such as cell culture media

Stop buffer

2 M Sulfuric Acid

Capture Antibody Solution

15 ug/ml antibody in coating buffer

Detection Antibody Solution

10 ug/ml in (20% Diluent buffer/80% Wash Buffer)

Enzyme Conjugated Antibody Solution

200 ng/ml in (20% Diluent buffer/80% Wash Buffer)

HRP Substrate

TMB (3,3′,5,5′-tetramethylbenzidine). 1 mg/ml. Usually commercially available as a solution.

Step-By-Step Method for ELISA

  1. Prepare a standard curve with your antigen in Diluent Buffer spanning a wide range of concentrations from 0 pg/ml to 3 times your maximum expected antigen concentration (3000 pg/ml approximately)
  2. Dilute the capture antibody to 15 ug/ml and have enough for 100 ul/well
  3. Add the capture antibody to the polystyrene plate, cover, and incubate at room temp. for 2 hours
  4. Remove the solution from each well and add in wash buffer (200 ul per well). Shake for 5 minutes. Repeat 3-5 times.
  5. Add 200 ul of blocking buffer per well, cover and incubate at room temperature for 1 hour (or overnight at 4 oC).
  6. Prepare the samples and standards such that you have 100 ul per well
  7. Remove the wash buffer and add in your sample + standard antigens into different wells. Cover and incubate at room temperature for 1 hour
  8. Repeat Step 4  to wash the plate
  9. Add 100 ul of the Detection Antibody per well. Incubate at room temperature for 1 hour.
  10. Repeat Step 4 to wash the plate
  11. Add 100 ul of the Enzyme conjugated Antibody to each well and incubate for 1 hour at r.t.
  12. Repeat Step 4 to wash the plate (2 times). We need to make sure the plate is very clean and any non-specific binding is minimized.
  13. Add 100 ul of the HRP substrate solution (1 mg/ml TMB)
  14. Incubate until blue (usually about 10 minutes at room temperature)
  15. Add 100 ul of Stop Buffer. This should make the solution yellow.
  16. Measure using a plate reader at 450 nm absorbance.

Notes on this ELISA method

    • Note 1. Your standard curve needs to span beyond your antigen concentration because you need to determine the exact amount of your antigen within the linear range of the standard curve. If necessary, dilute your antigen solution down to a point where it is within your standard range.
    • Note 2. Concentration of antibodies used will need to be optimized. It is highly likely that you will need to dilute each of the antibodies down rather than increase their concentration because these are at the upper ranges of the necessary concentration.