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.
We have already discussed a high level view of gene cloning in our Molecular Cloning Guide blog post. However, in that blog post we didn’t delve very deep into how we can perform each of the individual steps. Today’s blog post is about ligation. Ligation is the process by which two pieces of DNA can be glued together to form one piece. So, to begin, let’s assume you’ve already decided on a gene product that you want to clone. You’ve also designed primers and completed PCR on the open reading frame in your donor DNA (this could be genomic or non genomic DNA). Your next steps are to digest the PCR product with restriction enzymes and generate sticky ends. You’ll also want to digest your “shuttle” plasmid to generate complimentary sticky ends which will allow your “insert” DNA to click into position into your vector. It’s like a puzzle piece!
As you can figure out, generating sticky ends and complimentary ends is extremely important to the above process. However, several different restriction enzymes are available and each of them has different locations where they cut. Also, the type of cuts that they introduce may be “sticky” or “blunt”. Depending on the cloning strategy you are using, you may mix and match different enzymes to achieve different end goals. Ligation of “sticky ends” is much more efficient than ligation of “blunt” ends. Typically 10-100 times more T4 Ligase is required for blunt ends.
Here’s an image with various restriction enzymes and the kinds of ends they produce. Depending on the type of ends, your DNA ligation will proceed very differently!
Ligate DNA via DNA Ligase
Once the restriction enzyme digestion is complete, you can proceed to the ligation step. But, before you digest anything, make sure you’ve planned everything properly! You need to make sure that the insert will be ligated in the proper direction in the shuttle vector. Only once you’ve vetted your overall strategy, should you proceed to ligation and transformation, etc.
There are several kinds of ligase enzymes but the enzyme produced by T4 bacteriophage-infected E. Coli is the most common one. This ligase is called T4 ligase. Whereas normal E. Coli produce DNA ligase that uses NADH as a cofactor, T4 infected E.Coli produce a ligase that uses ATP as a cofactor. This enzyme will find the 3′ Hydroxyl and 5′ Phosphate within your sticky ends and it will form a phosphodiester linkage. If this is confusing, check out the Polymerase chain reaction (PCR) guide for images on what DNA looks like. This is shown here:
Protocol for Ligation of Transgene Insert into Shuttle Vector
Ligation enables fragments of DNA to be combined, such as the cut ends of transgene inserts and plasmids during cloning. This protocol describes the directional cloning of a XbaI/SalI-digested transgene into a shuttle vector, pAdtrackCMV, via cohesive end ligation.
Materials for DNA Ligation
XbaI/SalI digested, gel-purified insert (approx. 1 kb) and pAdTrack-CMV shuttle vector (approx. 9.3 kb; Plasmid #16405, Addgene)
Quick Ligation Kit (contains DNA ligase and 2X Reaction Buffer; #M2200S, New England Biolabs)
Agarose plate containing ethidium bromide
Estimate the DNA concentration of purified insert and vector preparations by applying 1 µl to an agarose gel plate (+ethidium bromide) alongside a range of DNA standards and visualizing under UV light.
Prepare the ligation mix as follows:
XbaI/SalI digested pAdtrackCMV 50 ng
XbaI/SalI digested insert 17 ng
Add water up to 10 µl total volume.
Add 10 µl of 2X Reaction Buffer and mix.
Add 1 µl of DNA ligase and mix.
Microcentrifuge briefly to settle liquid to the bottom of the tube and incubate at 25°C for 5 min.
Place on ice* and transform into desired bacterial strain.
Tips and Tricks for DNA Ligation
This reaction setup is using a digested insert to vector DNA molar ratio of 3:1. Inserts of different sizes will require a different amount to be added. Important ligation control reactions to include are (1) digested vector only and (2) digested insert only.
Ligation reactions can be stored at -20°C for future use
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.
We all know that DNA is the basic building block of biology. So, how can we make use of DNA to change cell biology? Well, today’s blog post will focus on “gene cloning” — making plasmids (circular DNA strands) so that we can introduce them into bacteria using our previous bacterial transformation method. With a plasmid inside the bacteria, you can a) use bacteria to make copies of the plasmid, b) make new proteins with the transformed bacteria and c) do the same inside mammalian cells using the Calcium Phosphate transient transfection method that we developed earlier. With molecular cloning techniques, we can control biology and make cells do some really cool stuff! Note: this is an overview post and does not have a step-by-step protocol associated with it. I’ll tease apart the different steps in future blog posts.
Molecular Cloning of Plasmids: Primer Design
“Cloning” refers to the process of making a copy of a gene so that we can modify it and see what happens. Remember, if you modify genes, your cells start producing new proteins; these proteins could be therapeutic and/or give your cells some new skills. To start, you’ll probably want to review the PCR protocol & guide to remember how PCR works. Now, let’s say we have a gene that we want to clone already available. The next most important part of PCR based gene cloning is the primer…so to design a primer, we need the following:
Hybridization sequence: A series of bases that compliment the bases right before your “target gene” or gene of interest.
Leader sequence: A few extra bases for our restriction enzymes to make efficient cuts that don’t overlap with our gene of interest.
Restriction sites: Places that we will cut so that we can make the plasmid circular.
Take a look at this image to understand the above plasmid design:
Be Careful Designing Plasmid Primers for Gene Cloning
Based on the above image, you can tell that if an enzyme’s restriction site is inside your gene of interest, you cannot use that restriction enzyme because you’ll cut your gene. Also, you’ll be putting this gene into a new plasmid. Make sure that the restriction enzyme you use is compatible with the “multiple cloning site” within this new plasmid. If you end up inserting this gene in random locations, the probability that this plasmid will be incorporated into the bacteria or expanded will be significantly decreased.
Look at the image below to understand these tips:
Gene cloning with PCR
With the primer already designed, we are ready to clone our gene. The rest of the steps in the gene cloning process are:
Use restriction enzymes to digest the PCR product
Use Gel Electrophoresis to purify the insert and the “vector” (recipient plasmid)
Ligate the plasmid
Transform bacterial cells
Isolate our plasmid for future use
Analyze the PCR products
Since we already know how to do PCR from our previous blog post, let’s focus on the other stuff. The first step listed is to digest the PCR product. For this, we will use restriction enzymes and incubate them with the PCR products. If everything was designed properly, we would know exactly where the restriction enzymes will cut the DNA in both the “vector” and the “insert”. Next, we will run these restriction digests on a gel and pick out the bands corresponding to our vector and insert (which we already know the size of). Any other “junk” PCR products will be removed in this step. The vector and insert DNA will then be “ligated” to form our new plasmid. To confirm our gene is in this plasmid, we will transform some bacteria with it on a petri dish. Try to make dilutions of your bacteria so that you can grow colonies of bacteria and pick out colonies later on. With the colonies that you pick out, you’ll want to isolate their DNA and digest it to see if your vector and insert are inside. We’ve already isolated the vector and insert in the past, so it’s simple to find out if our insert is inside the bacteria. Finally, as another confirmation, we will sequence the DNA from the bacteria and confirm that everything exists. We will write more about each of these steps in the future, but we wanted you to see them together, as an overview, in this blog post.
Take a look at the steps below:
Tips and Tricks with this methodology
: Make sure you choose the melting temperature to match the part of the primer that binds the “open reading frame” (your gene of interest). If you choose the wrong melting temperature, you might get the wrong PCR products because either a) your Tm was too low and you didn’t split the ORF or b) your temp was too high and you got lots of non specific binding.
: Make sure DNA digestion occurs for a long time, preferable overnight, to make sure all your vector and insert products were cut and maximize your ligation in the next step. You may need to use alkaline phosphatase in this step. I’ll speak more about that in the future.
: During gel electrophoresis make sure that you run the correct controls and *know* what wells relate to each of the digested products. Also, make sure you skip lanes to make cutting the wells easier. After this, you’ll need to quantify your DNA so you have enough for the ligation step. You can use a UV spectrometer for this step.
: Ligation also requires you to have several controls. For example, you need a ligation reaction without any insert. This will tell you how much background self-ligation your recipient plasmid has. You also need a ligation with some of the other bands you see during your gel electrophoresis. This will tell you how much contaminant DNA there was in your ligation.
“TA cloning is another approach if cloning doesn’t work in systematic way” –Swapnil Oke on Linked In
“I think for completeness I think it would be valuable to also mention a few other plasmid features that are important. I didn’t see mention of ribosomal binding sites (RBS) or origin of replication, etc” – Michael Kim on Linked In. — I plan on write about more details regarding plasmid design and purification in the future. For now, please don’t use the above blog post as a comprehensive guide…more like an overview 🙂
Bacterial Transformation using Competent Cells: Summary
Since we have already learned Calcium Phosphate Transfection with mammalian cells, let’s now focus on bacterial transformation of DNA with competent cells. In general, bacterial cells take up naked DNA molecules or plasmids via a process called transformation. Usually, this happens at a slow rate, but when bacterial cells die in close proximity to others, or when they are stressed, the transformation process occurs at a much higher rate. However, not all bacterial cells can be transformed, so biologists use ‘Competent Cells’ which are more inclined to take up DNA. The end goal of transformation is to get bacteria that have your genes of interest so that they will replicate your genes along with their own. If the bacteria contain your genes of interest, you can use them to mass produce proteins, or just store them for extended periods of time because bacteria are so hardy. A good way to test whether your genes of interest were transformed is to include antibiotic resistance in your plasmid. This way, you can be fairly certain that if your bacteria are resistant to antibiotics, they are also carrying genes of interest to you.
Take a look at how natural transformation works:
Transformation Biology in Bacteria
For bacteria, survival is key and transformation is one of their survival mechanisms. As biologists, we can make use of this survival mechanism for our benefit as well. To do this, we first incubate our competent bacteria with our plasmid and calcium chloride. Bacterial membranes are permeable to chloride ions, but not to calcium. So, as chloride ions enter the cell, the bacteria tend to swell (because they also intake water with chloride ions). Then we heat the bacteria in a process called ‘heat shock’ such that they turn on their survival genes. This causes the bacteria to uptake the surrounding plasmids. With the right design, this plasmid will then be recognized by bacterial DNA polymerases (remember our PCR Guide ?) and it will be expressed/replicated along with the bacteria’s normal DNA.
Take a look below to understand how biologists transform cells:
Selecting for Transformed Bacteria with the Lac Z Operon
Once your target plasmid is inside the bacteria, you still need to separate transformed cells from those that are not transformed. Another key challenge is that the transformation process may lead to some DNA being recombined so that your gene of interest is no longer functional. How do you select for cells that only contain functional target DNA that hasn’t been recombined? The trick is to use both antibiotic resistance and a Lac Z operon. By cloning your plasmid along with a Lac Z operon, you give your cells the ability to make a galactosidase protein. If cells have the galactosidase and you feed them X-Gal, they turn blue; cells without this operon are white. So, you first transform all your cells. Then you feed them IPTG to activate the Lac Z operon and cause cells to produce the galactosidase. Then you add in X-Gal and just pick out the bacteria that have functional Lac Z because the useful cells will be a bright blue!
Check out the figure below:
Bacterial Transformation Protocol
Transformation describes the uptake and incorporation of plasmid DNA into bacteria. Antibiotic resistance genes carried on plasmids allow selection of transformants. This protocol describes the transformation of DH5α E. coli with pAdtrackCMV (a vector carrying kanamycin resistance).
Materials for Bacterial Transformation
Ligation mix (20 µl) – insert ligated into pAdTrack-CMV shuttle vector (Plasmid #16405, Addgene)
DH5α competent cells (includes pUC19 DNA control; #18265017, ThermoFisher Scientific)
LB broth (#10855-021, ThermoFisher Scientific)
LB Agar selective plates (prepare from #22700025, ThermoFisher Scientific) with 50 µg/ml kanamycin (#15160054, ThermoFisher Scientific)
Step by Step Transformation Protocol
Thaw competent cells on ice. Aliquot 50 µl into cooled Eppendorf tubes for each transformation reaction.*
Add 5 µl of ligation mix to each tube.*
Incubate on ice for 30 min.
Heat-shock the cells for 20 sec in a 42°C waterbath.
Place on ice for 2 min.
Add 950 µl of warm LB broth per tube.
Allow cells to recover at 37°C for 1 hour with gentle shaking.
Spread 200 µl and 20 µl of each transformation mix onto warm selective plates.*
Incubate plates overnight at 37°C.
Notes on this methodology
We will talk about “Ligation” in another future blog post
Step 1. Unused cells can be refrozen and stored at -80°C for future use.
Step 2. As a transformation control, add 1 µl of pUC19 plasmid to one aliquot of cells (pUC19 confers resistance to ampicillin so will need to be seeded onto different selective plates).
Step 8. Transformation mix can be stored at 4°C and plated the next day if required.
Thymidine and BrdU, Cell Proliferation Assay Summary
How do you know if your cultured cells are growing? Does your new cancer drug affect cell proliferation? What’s the effect of VEGF on endothelial cells? As you can tell, knowing how to perform cell proliferation assays is an absolutely essential skill for anyone in biology, biochemistry, or pharmaceutics. Radioactive Thymidine cell proliferation assays have been used since for over 40 years to detect whether cells are growing. The principle is simple: cells will incorporate Thymidine into their DNA as they proliferate. However, dealing with radioactivity is painful and annoying, so new fluorescence-based, non-radioactive, BrdU and EdU cell proliferation assays have become the new mainstay technique. These molecules are both thymidine analogs and hence work using the same principle as radioactive thymidine. In today’s guide, we will learn Step-by-Step, the theory behind these assays and how to apply them in the lab. Combining our techniques of MTT Cell Viability assays and Flow Cytometry or FACS, we are really building up a great list of skills to analyze biological phenomena!
Principle of Cell proliferation assays with nucleotide analogs
3H-Thymidine is a radioactive version of the Thymine DNA base (thymine + the sugar backbone = thymidine). When cells are incubated with thymidine, they use the radiolabeled thymidine to synthesize DNA and incorporate it into their DNA backbone. So, thymidine is an excellent measure of DNA synthesis in cells that have undergone the S-Phase of cell replication. Similarly, BrdU is a Thymidine analog that lacks the radioactivity from tritium and it is used identically to Thymidine. Just incubate cells in the presence of BrdU. However, unlike radioactive thymidine, BrdU is detected with Anti-BrdU antibodies.
A quick summary picture is shown below.
Using Thymidine vs. BrdU. Cell Proliferation Assay Tips and Tricks
Taking things with a grain of salt: Note that DNA replication can happen even when cells are not proliferating. For example, if you have damaged DNA (ie. DNA repair is taking place). So, Thymidine and BrdU assays are really DNA replication assays and not perfect cell proliferation assays. But, for the most part, they are the gold-standard when looking for cell proliferation.
Thick tissue sections? Choose your cell proliferation assay wisely: The 3H-Thymidine assay uses radioactivity. And the beta particles that are generated by this method cannot penetrate very deep into tissue. So, if you’re labeling tissue sections, make sure they are extremely thin! In these cases, BrdU is a great option because it penetrates deep into tissue and can be detected even from 50 um thick slices. This is illustrated below in the picture.
Not enough signal? Add more: Since cells are substituting the radioactive thymidine into their DNA. Adding more thymidine means you’ll get more incorporation. And, more incorporation means you’ll get more signal! So, if your signal is low, just add more of the nucleotide. BrdU, however, doesn’t behave this way. There is a limit at which adding BrdU doesn’t increase your signal.
Want to preserve your tissue? Use 3H-Thymidine: BrdU immunohistochemistry requires you to digest and disrupt tissue for visualization because the antibodies that detect BrdU need to access all the tissue. Detecting radioactivity just needs you to use a scintillation counter.
Other Methods: EdU Cell Proliferation Assay
BrdU immunohistochemistry has the disadvantage that you need antibodies to detect it. Because of this, you need to disrupt the tissue you are staining. EdU is a new version of BrdU which has an azide functional group. This can then easily be detected with a “click” fluorophore. This is shown below:
Step by Step Guide to BrdU Cell Proliferation assay in vivo
Here is how you can label proliferating cells in a mouse Materials for BrdU Assay
BrdU labeling reagent (Invitrogen, #000103)
BrdU staining kit (Invitrogen, #933943)
Opaque dark container for storing stained tissue
16% Paraformaldehyde in Water (#47608, Sigma Aldrich)
1% Paraformaldehyde with 0.5 M EDTA, pH 8 [Demineralization solution]
Histo-clear (#50-329-51 Fisher Scientific)
30% Hydrogen Peroxide (H2O2) in Water
Petri dish with wet paper towels and paperclips for humidifying tissue (see image)
Cell Viability is a common technique used by biochemists who are studying oncology and pharmaceutics. The most common use for cell viability studies is when determining the IC50 for a cytotoxic compound in cell culture. However, as you can expect, there are a lot of different times when you need to know if your cells are alive. In larger pharmaceutical companies, MTT Cell Viability studies for Cytotoxic compounds are performed as a high throughput method because companies routinely screen MASSIVE libraries of small molecule drugs. To measure cell viability, researchers typically use an MTT assay, Cell Titer Blue, Trypan blue exclusion, or ATP assay. In this method guide, we will walk through the theory behind all these methods and then end with a protocol for the MTT assay. It would be a great test of your skills if you could use our High Performance Liquid Chromatography (HPLC) Method Guide to detect the products of the MTT assay.
Using an MTT Assay to measure Cytotoxicity
In general, to measure cell viability, you need to incubate cells with a reagent and measure the conversion of your reagent into a product. If lots of cells are alive, most of your reagent will be converted. If lots of cells are dead, then your reagent will only be partially converted. For the MTT assay, the reagent used is
(3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) tetrazolium. This is a positively charged small molecule that undergoes NADPH-mediated conversion over to Formazan. Because of its positive charge, MTT can enter viable cells and non-viable cells with ease. Upon conversion, the Formazan product precipitates inside cells near the cell surface and can be detected using a spectrophotometer. Note: MTT only needs an intact and functioning mitochondria to be converted so it is a metabolic assay and not a proliferation assay. I’ll discuss cell proliferation assays in the future. The assay technique is very simple:
Grow an equal number of cells in different wells of a microplate
Add your cytotoxic compound and incubate
Then replace the media and add the MTT, let the cells convert the MTT (blue) into Formazan (purple)
Use SDS along with DMF or DMSO to resolubilize the formazan and to kill cells (stop them from converting any more reagent)
Then measure how much formazan was created using a spectrophotometer.
Take a look below to understand these steps:
Some researchers have even combined the use of the MTT assay with Flow Cytometry (FACS) to sort viable cells from non-viable cells. However, this is uncommon and there are much better stains for FACS such as Propidium iodide (PI).
Cell Titer Blue, Trypan Blue and ATP Assays
As noted above, the MTT assay is really a metabolic assay because the MTT molecule needs to enter a cell and get converted to Formazan using NADPH. While the exact mechanism of MTT’s metabolism isn’t clear, this means the mitochondria needs to be intact and functioning. So, if you add a cytotoxic material which reduces mitochondrial efficiency, you might get weird results. In this case, it’s useful to also know other live/dead assays. The other major cell viability assays that are used in research include:
Cell Titer Blue: Similar to the MTT Assay, this assay involves incubating cells with resazurin (blue) and forming resorfurin (pink) after the cells metabolize it. Generally the metabolism takes 1-4 hours but it is much more sensitive than the MTT assay because you can measure the product via fluorescence (Ex/Em 560 nm/590 nm). The main advantage of this assay is that you don’t need to resolubilize the product in DMF/SDS so it’s much simpler. This is also a great high throughput assay!
Trypan Blue Exclusion Assay: If you don’t have a spectrophotometer, then it’s simple to use the trypan blue staining method along with a microscope. Because trypan blue is a charged dye, it cannot permeate through living cells. So, simply incubating cells with trypan blue and looking under a microscope allows you to visually determine the # of viable cells (unlabeled), # of non-viable cells (blue), and the # of damaged cells (slightly blue). Count the number of cells in different fields of view and you’re done! Viability is just the ratio of live cells divided by total number of cells. The disadvantage with this method is that all you test is the membrane integrity of the cells. You don’t know if the cells are truly non-viable or just damaged a little bit.
ATP Assays: When cells are non-viable, they cannot make any more ATP whereas viable and happy cells can make ATP. Additionally, as soon as cells die, ATPases rapidly break down ATP. Using these bits of information, it’s easy to see why an ATP based assay would work really well. The theory is simple – lyse cells, stop ATPases from hydrolyzing ATP, add in Luciferase and Luciferin. You’ll get excellent luminescence signal for hours!
Note: there are several other MTT-like molecules which are also used in cell viability assays: MTS, XTT, WST-1. The general principle however is all the same. The only note-worthy difference is that some of these molecules don’t penetrate live-cells, so they give you the reverse signal (how many dead cells there are).
Here are some images describing the above methods:
Cell Viability with MTT Assay Protocol
Materials for MTT Assay
MTT Solution (5 mg/ml MTT in PBS, pH 7.4, #M2128 Sigma Aldrich)
Solubilization solution, recipe here:
40% v/v Dimethylformamide #D4551 Sigma Aldrich
2% Glacial Acetic Acid #320099 Sigma Aldrich
16% Sodium Dodecyl Sulfate #436143 Sigma Aldrich
pH 4.7 & 37oC
96 well plate
Hep G2 cells
Complete DMEM (indicator-free, no phenol-red) with 10% Fetal Bovine Serum
Cytotoxic compound (ex: Doxorubicin)
Step-by-Step Cell Viability MTT Assay
Make the above solutions. Store MTT solution protected from light at 4oC and make sure there is no precipitate in the Solubilization solution.
Seed 25 x 103 Hep G2 cells in a 96 well plate with 250 ul of DMEM.
Add your cytotoxic compound (5 uM for Doxorubicin). Incubate for a desired time period (24 hours for Doxorubicin).
Aspirate media and wash 3x with PBS.
Add 125 ul of DMEM with 25 ul of MTT Solution. Incubate for 2 hours at 37oC.
Add in 100 ul of solubilization solution.
Pipette gently to mix without creating bubbles.
Measure via absorbance at 570 nm using spectrophotometer.
MTT Assay Notes, Tips, and Tricks
Always set up positive and negative controls! For positive controls have cells untreated with any cytotoxic compound as part of your wells. For negative controls have cells treated with 3% SDS as part of your wells. Also, make sure to have wells that have no cells, only media.
Increasing the number of cells also increases your signal
Too much MTT forms Formazan crystals which will damage cells so you might see the cells changing morphology.
This is an end-point assay because the precipitate inside the cells will kill them. Don’t plan on keeping your cells alive for any further studies after you add the MTT.
Having thiol-containing compounds in solution will convert MTT over to Formazan, so you’ll get false-positive data.
Having phenol-red in your medium may also convolute your results. Dye-free media is important to use.
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.
RNA extraction and isolation is a precursor for many methods in molecular biology including Northern Blotting, RT-PCR, and Microarray analysis. This blog post will focus on this precursor method as opposed to other biology techniques on the Scigine blog where I’ve focused on direct analytical techniques. Nonetheless, there is a lot to learn about RNA isolation and plenty of theory that might be applicable in your research.
Important aspects related to RNA Extraction
While everyone knows DNA is double stranded and helical, few people know that RNA is typically a single stranded polymer. However, secondary structures do emerge within RNA due to complementary base pairing and structures such as tRNA can be double stranded. Also, due to secondary structures, flexible regions of RNA can actually catalyze the cleavage of phosphodiester linkages in nearby RNA chains.
The base polymer looks like this:
The RNA polymer has several ionic groups, inter-chain hydrogen bonds, and it is extremely hydrophilic. All of these forces need to be overcome in order to purify RNA from DNA, carbohydrates, and lipids which have similar structures and physical properties. Polymers of RNA can be short or long, but generally smaller strands that are less than 100 nu cleotide bases are hard to purify because they don’t separate well. Additionally, there are several RNAses present in cells and tissues that can catalyze cleavage of RNA chains, so extreme care needs to be taken to prevent degradation of RNA during RNA isolation and purification procedures.
RNA Extraction Method Guide
Typically RNA Extraction procedures start with cell lysis. A buffer that includes Guanidine Thiocyanate or other chaotropes is necessary to mask charges on RNA and water so that the polymer can be purified using solid-phase techniques. Chaotropes linearize the RNA polymer, disrupt hydrogen bonding and destroy the activity of any RNAses present in the cell lysate. Due to their ability to disrupt hydrogen bonding, they also lyse cells by disbanding the phospholipid bilayer.
To facilitate homogenization, a homogenization column (imaged below) may be used. By passing at high speeds through resins, viscous polymers such as DNA and lipids can be separated and the cell lysate will flow more easily.
Ethanol is then added to: reduce the overall water concentration in the sample and precipitate proteins. As you would expect, RNA is highly soluble in water. A spin column (solid-phase support, imaged below) is then used to bind the RNA from the cell lysate.
Initially a low chaotrope concentration wash buffer is used to clean the RNA sample and remove proteins while RNA remains attached to the column.
Next, an ethanol wash removes some of the chaotropic salts that were left over from previous washes.
Finally, without chaotropic salts present, RNAse-free water can be used to elute the RNA sample from the column
RNA Extraction and Purification: Step by Step
Here’s a technique for RNA Isolation and cDNA preparation from Pancreatic Islets. We can use this technique for analysis of gene expression later on. Our strategy is to use solid-phase extraction of nucleic acids from complex cell lysate samples and then we can prepare the cDNA to examine the expression of genes such as insulin.
Materials for RNA Extraction and cDNA preparation:
Isolated pancreatic islets
RNeasy minikit (#74104 Qiagen)
QIAShredder columns (#79654 Qiagen)
Omniscript reverse transcription kit (#205110 Qiagen)
Oligo dT primers (dilute to 10 µM; #18418012 ThermoFisher Scientific)
Random hexamer primers (dilute to 20 µM; #N8080127 ThermoFisher Scientific)
RNase inhibitor (dilute to 10 units/µl; #N211 Promega)
RNA Extraction/Purification Procedure:
Collect up to 100 islets in an Eppendorf tube and add 350 µl RLT buffer* to disrupt cells.
Vortex thoroughly and add to QIAShredder column with collection tube attached. Spin for 2 min at full speed in microcentrifuge to homogenize the sample.
Add 350 µl 70% ethanol to the lysate and pipette repeatedly to mix.
Transfer to an RNeasy spin column and spin at ≥8000xg for 15 secs to bind the RNA to the column. Discard the flow-through.
Wash the column with 700 µl RW1 buffer and spin at ≥8000xg for 15 secs. Discard the flow-through.
Wash the column with 500 µl RPE buffer* and spin at ≥8000xg for 15 secs. Discard the flow-through.
Repeat column wash with 500 µl RPE buffer and spin at ≥8000xg for 2 min. Discard the flow-through.
To elute the bound RNA, transfer column into a fresh Eppendorf tube and add 30 µl RNase-free water to membrane. Spin at ≥8000xg for 1 min.
To ensure a high RNA concentration, use the eluate to repeat the elution by reapplying to the membrane and spinning at ≥8000xg for 1 min.*
Determine the concentration and quality of the RNA sample.*
To reverse transcribe template RNA into cDNA, prepare the following reaction in one eppendorf:
Up to 2 µg RNA*
4 µl of 10X Buffer RT
2 µl Oligo dT primer
2 µl random hexamer primer
1 µl RNase inhibitor
1 µl Reverse Transcriptase
Add RNase-free water to take reaction volume to 40 µl.
Mix thoroughly by briefly vortexing and centrifuge briefly to collect reaction to the bottom of the tube
Incubate at 37oC for 60 min.
Place reactions on ice and use up to 1 µl per 10 µl PCR reaction*.
Notes for this RNA Isolation Procedure:
Step 1. On the day of RNA preparation, add 10 µl of 14.3 M 2-mercaptoethanol per 1ml RLT buffer prior to use. This helps disable RNAses.
Step 6. Ensure ethanol is used to dilute RPE buffer concentrate (provided in kit) prior to first use
Step 9. Store RNA samples at -80°C if required to prevent degradation
Step 10. RNA analysis chip techniques such as Experion (BioRad) use a small amount of sample to provide accurate quantification and assessment of RNA quality
Step 11. If RNA has been frozen, thaw on ice to avoid RNA degradation
Step 14. Reactions can be stored at -20oC prior to PCR if required
Notes from our readers:
Mr. Young on Google + states:
1. [It is important to] …keep… an amplicon-free, RSase- free, clean environment
2. [Also have] a dedicated space for nucleic acid extraction separate from post amp and Master mix prep
Dr. Carina Jorgensen from Linked In states:
[Check the] integrity of … [your] … RNA … – that is whether or not its degraded – before …[you]…continue with downstream applications…You need to know the quality of RNA to choose your primer for cDNA synthesis, and if you want to continue with real-time qPCR you need to have a certain minimum level when it comes to the quality of your RNA, if you want to be sure that you can trust the following qPCR results. If you want to perform microchip analysis the demand to quality is even higher comparer to qPCR.
Guides for Specific Applications of RNA Extraction on Scigine
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