Guide: Measure Cell Proliferation with Thymidine and BrdU

Cell Proliferation, BrdU, Thymidine, EdU

Thymidine and BrdU, Cell Proliferation Assay Summary

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

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Principle of Cell proliferation assays with nucleotide analogs

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

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

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

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

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

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

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

Thymidine Cell Proliferation Assay vs BrdU Assay

Other Methods: EdU Cell Proliferation Assay

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

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

Step by Step Guide to BrdU Cell Proliferation assay in vivo

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

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

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

Notes on this BrdU Cell Proliferation Assay Methodology

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

Checking Cell Proliferation in Zebrafish: Video

Applications of BrdU, EdU, and Thymidine Assays on SciGine

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


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

Cytotoxicity & Cell Viability with MTT Assay Protocol

Cell Viability with MTT Assay and Cytotoxic Compounds

Cell Viability with MTT Assay Summary

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.

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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:
Cell Viability with Doxorubicin Cytotoxicity
Cell Viability MTT Assay Steps
Trypan Blue and MTT Assay are similar

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:
Cytotoxicity measured with Cell Titer Blue
Trypan Blue stains cells with Cytotoxic compounds
ATP Assay with Cell Viability or Cell Proliferation

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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

  1. Make the above solutions. Store MTT solution protected from light at 4oC and make sure there is no precipitate in the Solubilization solution.
  2. Seed 25 x 103 Hep G2 cells in a 96 well plate with 250 ul of DMEM.
  3. Add your cytotoxic compound (5 uM for Doxorubicin). Incubate for a desired time period (24 hours for Doxorubicin).
  4. Aspirate media and wash 3x with PBS.
  5. Add 125 ul of DMEM with 25 ul of MTT Solution. Incubate for 2 hours at 37oC.
  6. Add in 100 ul of solubilization solution.
  7. Pipette gently to mix without creating bubbles.
  8. 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.

Cell Viability Protocols on SciGine

Checking IC50/Cell Viability via MTT Assay with Cytotoxic compound: Video


Excellent NIH guide related to Cell Viability
Trypan Blue Exclusion information
Discussing effectiveness of MTT assay for leukemia research

Calcium Phosphate Transient Transfection Protocol & Guide

Calcium phosphate Transient Transfection Protocol and Guide

Transfection with Calcium Phosphate: General Summary

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

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Transient vs. Stable Transfection

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

Calcium phosphate Mammalian Transfection

DNA Transfection guide

Transient Transfection vs Stable transfection

Calcium Phosphate Transient Transfection

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

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

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

The general technique is shown below:

Calcium phosphate Nanoparticles and Aggregates

Phagocytosis of Calcium Phosphate leads to Transfection

Selection Media Confirms Stable Transfection with Calcium Phosphate Protocol

Tips and Tricks when optimizing your Calcium Phosphate Transient Transfection Protocol

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

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

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

Calcium Phosphate Transient Transfection Protocol

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

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

Transfection Protocol Steps

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

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BBS High Efficiency Transfection Protocol Steps

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

Notes on this transfection methodology

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

Applications of Transfection on SciGine

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

Transient and Stable Transfection Video Tutorial


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

Immunofluorescence Microscopy Protocol and Method Guide

Immunofluorescence Microscopy Protocol and Method Guide

Immunofluorescence Microscopy Overview & Theory

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

Related articles:

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

Advantages and Disadvantages of Direct vs. Indirect Immunofluorescence Microscopy

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

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

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

Dealing with the weaknesses of Fluorophores in Immunofluorescence Microscopy

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

How Fluorescence works:

Immunofluorescence Microscopy Theory of Fluorescence and Electron Energy

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

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

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

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

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

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

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

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

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

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

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

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

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Immunofluorescence Microscopy Step-By-Step Guide

Example Immunofluorescence of Pancreatic Sections

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

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

Experimental procedure:

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

Procedural notes

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

SciGine Immunoprecipitation Microscopy Protocols and Methods

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


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

Immunofluorescence Theory Video by the HHMI

Southern Blot Scientific Method Theory and Guide

Southern blot scientific method title

Southern Blots: Overview & Theory

Southern Blotting is a technique that is used to detect whether you have a specific DNA sequence available in your sample. Most commonly, you will be testing for DNA that you get from cell lysates or after creating your own plasmids. You can also do some amazing things like making transgenic mice and proving that you have selected for certain genes of interest to you! All you have to do is Southern Blot any set of DNA from the mice and you’ll have concrete proof. As you probably understand, this is one of the most common techniques that geneticists and molecular biologists know because everything they do needs to be proved by a southern blot.

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

To execute a southern blot, first collect some DNA. This can be from a cell lysate or from tissue samples, etc. Next, digest the DNA using DNAse and run these fragments on an agarose gel. This will separate the DNA by size and you’ll know which well in the gel corresponds with which sample. Next, in a similar fashion to Western blots, this DNA is then transferred onto blotting paper (typically made of nitrocellulose or nylon). After the “blotting” process is complete, the DNA is then “probed” using a radioactive (or fluorescent) DNA sequence that is complementary to the sequence that you want to detect. Finally, this “probed” radioactive blot is then imaged using an autoradiograph. The steps for this process are described in the following image:
Southern Blot Scientific Method
Southern Blot Step by Step
Southern Blot Tips and Tricks
Southern Blotting Method

Nuances of Southern Blotting theory

Why is there is a second denaturing step for DNA that’s 15 KB or larger?
It’s because these large pieces of DNA don’t transfer very well to the blotting paper. As you can imagine, the larger a piece of DNA is, the slower it will migrate through an agarose gel. Even after leaving the blotting paper overnight, the transfer may not be complete!

How does the DNA actually go from the gel into the blot?
Through capillary action and wicking! Unlike a western blot where a voltage gradient is utilized to pull proteins into the blotting paper, in Southern Blots, the DNA merely moves over to the blotting paper overnight without much force at all. That’s why it is incredibly important to make sure that the blotting paper and the gel are in close contact. It’s also very important to make sure that the glass plate on top is heavy enough so that it forces the gel and the blot together.

Why is there a UV step?
By using UV after the transfer step, the DNA (which has some free aldehyde groups due to depurination) can react with the nitrocellulose/nylon membrane to form covalent bonds.

What do the NaOH and the HCl do during the denaturation step?
Essentially HCl removes some/all of the purine bases from the DNA and makes the two DNA strands less sticky to each other (because there is less hydrogen bonding). This process is called depurination. NaOH also prevents the two strands from forming hydrogen bonds due to deprotonation of all bases.

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

How does radiolabeling with P32 work?
Small amounts of DNAse introduce nicks into the single stranded probe DNA. DNA polymerase then utilizes the dATP32 from solution to repair these nicks and incorporates these radioactive phosphates into the backbone of the DNA.

Southern blot Step-By-Step Guide

Materials for Southern Blot of Mouse Tail DNA

Denhardt’s Solution 50X (5g Ficoll 70000, 5g Polyvinyl pyrrolidone, 5g BSA Fraction V in 500 ml water)
Hybridization cocktail (25 ml 50% dextran sulfate, 25 ml 20X SSC, 50 ml formamide, 1 ml Tris 1M, 2 ml Denhardt’s Solution 50X, 1ml 10% SDS) – prefilter before use
TE buffer (10 mM Tris HCl pH 7.5, 1 mM EDTA)
BamHI enzyme and buffer (#R0136S New England BioLabs)
100X BSA (B9000S New England BioLabs)
Gel loading dye (#G2526 Sigma)
TBE buffer 10X (#93290 Sigma)
SSC buffer 20X (#S6639 Sigma)
[α32P]dCTP (3000 Ci/mmol; #NHG013H250UC Perkin Elmer)
Sephadex G-50 (#G50150 Sigma)
Ready-To-Go DNA Labeling Beads (-dCTP) (#27-9240-01 GE Lifesciences)
Whatman paper (#WHA10427810 Sigma)
Salmon Sperm DNA (#15632011 ThermoFisher)

Southern Blot Protocol
DNA digest and gel electrophoresis

    1. Setup DNA digest reactions from collected tail samples as follows:

10 µl DNA (12 ug)
6 µl 10X BamHI buffer
0.6 µl 100X BSA
1 µl BamHI
42.4 µl H2O
Incubate at 37°C for at least 5 hours.

  1. Add 6 µl gel loading dye to each sample.
  2. Run samples of a 0.8% agarose gel in TBE for 16 h at 25V.
  3. Stain gel in 1 µg/ml ethidium bromide for 30 min.
  4. Take a photo of the gel.*


    1. Soak gel in 0.2 N HCl for 10 min with shaking to depurinate (remove purines from DNA). Rinse with water.
    2. Soak gel in two washes of 500 ml 0.4 M NaOH/1.5 M NaCl solution for 30 min each time.
    3. Soak gel in two washes of 500 ml 1 M Tris/1.5 M NaCl solution for 30 min each time.
    4. Setup transfer stack by placing 2 layers of Whatman paper on glass plate (wet before laying down) with soaked gel on top (flipped over). Cover with plastic wrap.
    5. Using a razor blade, cut out plastic over gel and place wetted membrane on gel. Place two layers of Whatman paper on top (wet before laying down).
    6. Place a stack of paper towels on the top and cover with a glass plate and transfer in 6X SSC overnight.
    7. Dismantle and air-dry membrane.
    8. Expose membrane to UV for 90 s (DNA side down). Bake between Whatman paper at 80°C for at least 1 h.

Labelling the probe

      1. Denature probe by adding 2 µl probe (25-50 ng) to 44 µl TE and incubating at 95°C for 4 min). Immediately place on ice.
      2. Add 46 µl denatured probe and 4 µl of [α32P]dCTP to Ready-to-Go labelling bead tube. Flick to mix.
      3. Incubate at 37°C for 15-30 min.
      4. Add 50 µl TE and run through a G50 sephadex column

Probe Hybridization and Autoradiograph

      1. Add 50 ml hybridization cocktail to the membrane.
      2. Add 300 µl of 10 mg/ml salmon sperm DNA to labelled probe and denature at 95°C for 5 min. Immediately place on ice. Add to membrane and cocktail solution.
      3. Hybridize overnight at 42°C.
      4. Wash briefly with 50 ml wash solution (0.2X SSC/0.1% SDS).
      5. Wash for 30 min with 100 ml wash solution at 50°C. Repeat.
      6. Check blot with Geiger counter to estimate cpm.
      7. Expose membrane to film and store at -70°C.
      8. Develop after 2-5 days.

Notes on this Southern Blot method

      • During DNA digest Step 5 you can note the position of the marker bands by using a fluorescent ruler or marking bands directly on the gel by punching small holes with a needle.
      • This method uses dCTP32 and not dATP32 as we talked about in the theory section
      • After undergoing HCl and NaOH treatment for the denaturation step, it is very important to neutralize the gel once again.
      • If you don’t want to buy the labelling beads, an alternative strategy is presented here at MIT Cores

Application of Southern Blots on SciGine, a Scientific Method Search Engine


ASU Southern Blots Guide
MIT Core Guide
DNA Blot from Davidson College

Southern Blot Video by Shomu’s Biology

Flow Cytometry and FACS: Method Guide

FACS & Flow Cytometry for Cell Population Analysis

FACS Method Overview: Using Fluorescence to Understand Cells and Cell Populations.

FACS, or Fluorescence Assisted Cell Sorting, is a type of technique that enables you to understand cells by tagging them with fluorescent markers. There are a number of things that FACS allows you to do:

  • Separate one type of cell from a mixture of cells
  • Count how many cells are in a mixture
  • Detect what kind of biomarkers a cell might have (is it cancerous? Just a regular lymphocyte? Etc.)
  • Find out if your cell is making a protein of choice (similar to biomarker detection)
  • And more.

It’s an amazing and powerful technique that you should always keep as part of your skills. Most Bio-Pharmaceutical companies have FACS experts in-house because it is so versatile!

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

The principle of FACS is simple:

  1. Label cells of interest with a marker such as an antibody
  2. Pass the cells through a laser and detect which ones have the antibody
  3. Separate those cells from the rest

Here’s what this looks like in a simple picture:
FACS and Flow Cytometry Method Overall

What does the FACS instrument detect?

  • Forward Scatter Light (FSC): When you shine a laser on an object, the object blocks some of it and the rest continues to go forward towards the detector. This is called forward scattered light.
  • Side Scatter Light (SSC): An object might also bounce some of the laser in an alternate direction. There is a detector that is orthogonal to the light direction and detects this scattered light.
  • Fluorescence Emission Signal: Excite your fluorophores with a certain wavelength of light and they’ll emit a different wavelength depending on the type of fluorophore. Fluorescence detectors in the instrument will detect this emitted light.

Take a look at this illustration:
FACS Detection of Molecules and Cells
Based on the previous section, FACS provides data on forward scattered light, side scattered light, and fluorescence intensity. By graphing these different values together, you can get an idea of what your cell sample is like. Here’s the kind of data you would get:

  • Plot of FSC vs. SSC:
    • Cell debris in your solution is usually small, so it would have low FSC and SSC values.
    • Normal cells have medium sized FSC and SSC values
    • Large cells like Granulocytes have large FSC and SSC values
  • Plot of Fluorescence intensity:
    • After cleaning your cell solution and removing any non-specific fluroescence, your cells will be the only thing with large fluorescence intensity
    • A plot of intensity provides you a histogram telling you how much fluorescence is in the solution
    • You can use this to find out how many cells are there, how many antibodies bound to each cell, how much of a certain biomarker is there, etc.
    • You can compare different fluorescence histograms to compare cell types as well

Take a look at these images:
FACS Understanding Dot Plots and Histograms

Gating: How to measure cells of only one population and ignore debris

In the above images you can see that your figures may provide information on several types of particles: debris, small cells, large cells, particles with high and low fluorescence, etc. In order to narrow down your results, most instruments allow you to “Gate” your results so that the instrument provides you with a new graph within limits that you set. As an example, you may plot FSC vs. SSC and then gate only for larger particles. Then using that gate you may look at the fluorescence intensity in Red, Green, and Blue. This way you can analyze cells that have red biomarkers, or cells that have blue biomarkers specifically.

As an example, what if you wanted to only collect cells that highly express one cancer-related cellular protein (ex: folate receptor) ?

Using FACS, this would be easy to analyze! Just mark your cells with an anti-folate receptor antibody with GFP. Then, use FACS to separate out the cells that express the receptor in your sample. Take a look at this image to understand this concept.
Gating in Flow Cytometry

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

Complete Step by Step Method Guide

Unfortunately, this is such a highly instrument-oriented technique that it’s difficult for me to provide details on how to operate a FACS machine. Nonetheless, take a look here to get an idea.

Tissue Culture Flask
With cells such as HT-29
Trypsin such as Tryp LE from Invitrogen
Serological pipette
Without pH indicator so that there isn’t any extraneous fluorescence


  1. Grow HT-29 to confluence in a T-75 flask in 7 ml of DMEM
  2. Use lab protocols to mark them with any antibodies that you’d like such as an anti-folate receptor antibody
  3. Aspirate the DMEM and add PBS 1x
  4. Swirl around and remove PBS
  5. Wash once again with PBS by following the previous 2 steps
  6. Add 2 ml of Trypsin (TrypLE) and wait until cells detach for approx 10 minutes
  7. Collect cells into a falcon tube by using a serological pipette
  8. Use a hemacytometer to dilute cells until they are at a concentration of 1×106 cells/ml using DMEM
  9. Load cells into the FACS using using appropriate 5 ml tubes
  10. Make sure the instrument is running well by using control samples or any extra cells that you have
  11. Replace cell collection tubes and make sure they are sterile if you plan on culturing cells that are collected
  12. Run the FACS instrument 🙂

Notes on this FACS/Flow Cytometry Methodology

  • FACS may also be referred to as Flow Cytometry on Job Postings. This is because FACS is a part of the overall group of techniques called Flow Cytometry. In biology, however, it is unlikely that you will use any other techniques besides this one.
  • The types of tubes that are necessary for loading a FACS unit vary between instruments
  • Some cells clump up and make data analysis harder. Try diluting them more or mixing them more using your serological pipette
  • It’s important to set the instrument’s gain settings to maximize/minimize signals as appropriate
  • Make sure to also set the correct excitation/emission levels for your various fluorescence markers. You will also need to set your thresholds such that your signals don’t overlap.
  • If you plan on using multiple fluorescence markers at the same time make sure you consult a fluorescence ex/em table like this fluorescence spectra table. You need to make sure that your fluorescent signals don’t overlap.

Applications of FACS on SciGine

Analysis of cell cycle and viability with a FACScan
Analysis of a GFP reporter gene using FACS
Analysis of gene disruption in diploids
Using FACS to analyze camptothecin induced apoptosis
FACS analysis of cell cycle using Propidium Iodide


Excellent Antibodies-Online FACS reference
Thermofisher FACS manual
What is FACS from UMass