Fluorescent in situ hybridization is a very powerful method to optically detect specific nucleic acid sequences in fixed but otherwise intact cells. FISH can be used to detect specific DNA or RNA sequences. DNA FISH can yield interesting things (for instance to determine the number of gene copies, the location of the gene in the nucleus when it is turned on/off, DNA repair and replication studies). However, I will discuss here about RNA FISH, which has become a very powerful tool to study gene expression and RNA biology. I described an RNA FISH application in a previous post.
RNA FISH is a quantifiable method that can determine the localization of single mRNA molecules in the cell, their number, their transcription and degradation rates and more. I will dicuss these applications in other posts.
Recently, I started FISHing myself. Obviously, I ran into some difficulties (but also success). So I’ll start by describing the method and protocol, some tips and problems to avoid, how I analyze the data, different types of dyes etc… Hopefully, I’ll also learn something from all this writing.
The idea of FISH (or rather, ISH; patented by my PI almost quarter of a century ago) is that nucleic acids with complementary sequences tend to form a double helix. It can be DNA:DNA, RNA:RNA or RNA:DNA. Therefore, one can synthesize a short DNA probe that carries some sort of tag. In the olden days, it was radioactivity, or biotin – that would be bound by avidin. Today, the F in FISH stands for a fluorescent dye that is chemically bound to the DNA probe.
The length of the probe is significant. The longer it is, the more specific it will become. However, we will need harsher conditions for hybridizations and also it is more difficult to synthesize. Commonly, people use either 50 bases or 20 bases (20-mer) probes.
Tagging of the probe is also quite variable. The probe can be tagged at one or both ends or one or more times in the middle. The chemistry of end-labeling and middle labeling is different, and so is the synthesis process.
The most common fluorophores that people use are the cyanine dyes (known as the Cy dyes). But there are other types such as the Alexa series, Quasar dyes and more. The choice depends on required ex/em wavelengths, availability and funds. Today, many companies design and produce the probes for the researcher, but some companies use their own dye brand.
A major problem is that of signal intensity. We need multiple dye molecules bound to the same mRNA molecule in order to be able to detect it above the background. For this reason, it is recommended to have either multiple end-labeled probes (preferably >20 probes) or at least a few long multi-labeled probes (to get a minimum of twenty dye molecules or more per mRNA).
Here is a short version of the protocol (for adherent mammalian cells):
First, we need to grow the cells on a thin glass coverslip. This is important – the cells cannot be transferred from culture dish to the glass once the cells are fixed.
Next, we fix the cells (kill cells and immobilize all biomolecules) using 4% paraformaldehyde in buffer. This chemical creates multiple cross-linking bonds between all bio-molecules, thus “fixing” them in place relative to other biomolecules. This process keeps the shape of the cell and all the sub-cellular structures.
The fixation step needs to be calibrated – too much or too little can affect the hybridization efficiency. The protocol I got uses 10 min for fixation.
The cross linker is then removed, and the cells are permeabilized. There are two major protocols for permeabilization. One uses 70% ethanol at 4°C over-night. The other uses a low % solution of detergent (I use 0.1% triton X100) for a few minutes. From my brief experience, the triton works better (I’ll show images later).
The cells are then washed again, and placed in pre-hybridization buffer (2xSSC) that contains 10% formamide (for 20-mer probes) or higher (50% for 50-mer probes). The formamide denatures DNA and RNA secondary structures and assists in hybridization. SSC stands for saline-sodium citrate. The salts increase the stringency of the hybridization.
Hybridization is performed using a similar buffer, but with the addition of competitor RNA (usually tRNA) and competitor protein (BSA) to reduce background. The protocol also requires dextran sulfate (accelerates hybridization rate) and vanadyl ribonucleoside complex, a potent ribonuclease inhibitor (because we don’t want any rogue RNAses digesting the RNA in the cells).
Usually, a small drop (40-50µl) of the hyb buffer + probe is deposited on a clean surface in a humid chamber. The glass coverslip is them placed face down on the drop (i.e. cells facing the liquid).
The chamber is closed and placed at 37°C for at least 3 hours up to over-night.
We continue with several rounds of washing (which also includes an optional DAPI staining step) and finishing with mounting the coverslip onto a microscope slide using an anti-fade mounting medium.
Then, we go to the microscope.
- FISH for MEF cells expressing beta-actin-MBS, with MBS 20mer probe mix labeled with Cy3. Image is maximum projection merge of three channels using ImageJ. Blue- DAPI stain; green – GFP (cells are GFP negative); red – Cy3.
It is important to have a negative control (cells not expressing the RNA in question) for further analysis.
Here’s an image of negative cells:
FISH for wild type MEFs expressing GFP but not expressing beta-actin-MBS. Probe details and image processing as the previous figure.
I will discuss spot analysis at the next post.
Sources for protocols:
Singer lab website (these are protocols for 50-mer or longer probes).
A more recent published protocol of 50-mer (for yeast)
Stellaris protocols (Biosearch technology company) for 20mer probes. [ I do not use their mounting method, but use Prolong gold from Invitrogen. It works very well].
Protocols from Arjun Raj’s lab (includes their “Turbo FISH” – FISH in 5 minutes. Never tried it myself, yet).