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. Take a look at the image below:
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!
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:
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.
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!
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. 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).
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.
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.
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.
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.
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.
Step by Step Guide to Purification of a His-Tagged Fusion Protein
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.
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.
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 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:
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
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)
Materials for BBS Calcium Phosphate Transfection
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)
Transfection Protocol Steps
Split cells such that there is space between cells.
Clean DNA by adding in 100% ethanol for precipitation
Dry DNA after aspirating supernatant from ethanol. Use air to make sure it is completely dry.
Resuspend pellet in 450 ul of water with 50 ul of 2.5 M CaCl2 buffer
Put 500 ul of 2x Hepes Buffered Saline in a 15 ml conical falcon tube
Add the DNA/CaCl2 solution dropwise to this tube while agitating with a stir bar or other mechanism.
Allow the precipitate to sit at room temperature for 20 minutes
Spread the precipitate over the cells along with their medium . Shake gently to make sure the precipitate is even.
Incubate in cell culture incubator at 37 oC with 5% CO2 for up to 16 hours
Remove medium, wash twice with PBS, and feed cells with complete medium.
Plate cells in selective medium.
BBS High Efficiency Transfection Protocol Steps
Seed cells in a dish so that they can double atleast twice so they can be stably transfected
Next day, dilute DNA in TE Buffer at 1 ug/ul
Make a 0.25 M CaCl2 stock
Mix 20-30 ug of DNA with 500 ul of 0.25 M CaCl2 stock.
Add 500 ul of BBS to this mixture and vortex. Incubate at room temp for 20 min.
Add this mixture to the cell culture dish dropwise and mix by gently shaking the plate.
Incubate cells overnight for 24 h at 3% CO2 at 35 oC.
Wash cells 2x with PBS and then incubate them in complete medium for 2 doublings.
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.
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).
Here’s an image which describes the above theory:
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: Advantages 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. Disadvantages 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: Advantages 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. Disadvantages 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:
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.
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:
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.
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.
Why do we have non-specific fluorescence? Take a look at this image:
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)
Cut 8 µm paraffin sections at the microtome and mount onto glass slides.
Heat slides at 60°C for at least 5 min to melt wax.
Dewax slides in xylene for 2x 5min*, 100% ethanol for 2x 5min and immerse in running deionised water until clear.
Circle sections using a wax stick to easily observe staining area in subsequent steps.
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.
Block sections with 50 µl of 10% normal donkey serum in PBS at room temperature for 30 min in humidity chamber.
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.
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.
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.*
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.
Tip off excess solution from slide and mount in one drop of mounting medium with coverslip.
Examine at the fluorescence microscope*.
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
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:
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.
Immunoprecipitation of the Lck protein from T cell lysates has been used to analyse the phosphorylation patterns of this important T cell activation protein.
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)
IP Experimental procedure:
Perform all steps on ice to avoid protein degradation.
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.*
Pellet cell debris for 15 min at maximum speed on microcentrifuge at 4°C.
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.*
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.
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.
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.
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.
Here is a step by step illustration of how to perform a western blot:
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.
Unwrap precast gel and rinse wells three times with running buffer. Assemble gel in tank and fill with running buffer.*
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.
Heat samples to 95°C for 2 min and spin briefly to ensure contents are at the bottom of the tube
Load gel with samples and include ladder in one lane.
Run gel at 200V for 30 min.
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.
Remove gel from tank and place in transfer buffer.
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.
Close cassette and run in machine (standard minigel program for 30 min).
Remove transfer membrane from cassette, taking care to snip one corner to ensure orientation.*
Incubate transfer membrane in blocking buffer for 1 h at 4°C with rocking.
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.
Wash membrane three times in TBS with 0.2% Tween20 at 4°C with rocking, for 10 min each time.
Incubate with diluted secondary antibody 1:2500 in 5 ml TBS with 0.2% Tween20 at 4°C with rocking for 1 h.
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.
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.
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:
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.
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 vials
I only use autosamplers since manual injection is tedious 🙂
Centrifugal filters with 0.2 um pores
To clean up samples
In a typical HPLC procedure you can decide the following variables:
What it does
With fast flow peaks come out sooner but there’s they’re harder to resolve and tend to blend together. For more resolution, run slower.
Affected by flow rate and solvent
Determines signal intensity, how quickly the peaks come out, signal fidelity
Determines the type of interaction with the sample
If 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:
Dissolve your biopolymers or small molecules in a suitable solvent such as methanol
Centrifuge at 10,000 rcf in an eppendorf vial and keep the supernatant to remove any large particular matter
With a centrifugal filter, add 500 ul of your sample solution onto the top
Centrifuge at 10,000 rcf and collect the filtrate (the solution that successfully passes through the filter)
Load this sample into an HPLC vial
For setting up the HPLC machine:
Make sure you have all your buffers set up
Open the purge valve and purge the system for 5 minutes.
Add your samples into the autosampler tray
Stop the purge
Close the purge valve
Run the system at a normal flow rate (1 ml/min) with your buffer to equilibrate the column for 10 minutes
Make sure that your pressure is stable (ie, less than 2-3 bar of fluctuation)
Set up your sequence and your method
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.
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.
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:
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.
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)
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.
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).
A Complete Sandwich ELISA protocol
Materials for ELISA
96 well polystyrene plate
0.2 M sodium carbonate/bicarbonate buffer, pH 9.4
0.1 M phosphate, 0.15 M sodium chloride, pH 7.2 with 0.05% Tween 20
2% w/v Bovine Serum Albumin in Wash Buffer
2% w/v BSA in Wash buffer or a more appropriate buffer such as cell culture media
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)
TMB (3,3′,5,5′-tetramethylbenzidine). 1 mg/ml. Usually commercially available as a solution.
Step-By-Step Method for ELISA
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)
Dilute the capture antibody to 15 ug/ml and have enough for 100 ul/well
Add the capture antibody to the polystyrene plate, cover, and incubate at room temp. for 2 hours
Remove the solution from each well and add in wash buffer (200 ul per well). Shake for 5 minutes. Repeat 3-5 times.
Add 200 ul of blocking buffer per well, cover and incubate at room temperature for 1 hour (or overnight at 4 oC).
Prepare the samples and standards such that you have 100 ul per well
Remove the wash buffer and add in your sample + standard antigens into different wells. Cover and incubate at room temperature for 1 hour
Repeat Step 4 to wash the plate
Add 100 ul of the Detection Antibody per well. Incubate at room temperature for 1 hour.
Repeat Step 4 to wash the plate
Add 100 ul of the Enzyme conjugated Antibody to each well and incubate for 1 hour at r.t.
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.
Add 100 ul of the HRP substrate solution (1 mg/ml TMB)
Incubate until blue (usually about 10 minutes at room temperature)
Add 100 ul of Stop Buffer. This should make the solution yellow.
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.