Tag Archives: my pics

Does bound MS2 coat protein inhibit mRNA decay?

Roy Parker recently sent a  “Letter to the Editor“, published in RNA journal, in which he suggested that the MS2 system might not be best suited for live imaging of mRNA in budding yeast. According to Parker, the MS2 system inhibits the function of Xrn1, the major cytoplasmic  5′ to 3′ RNA exonuclease in budding yeast, causing us to image mostly the remaining 3’UTR fragments. Thus, he claims, it is possible that interpertation of mRNA localization data using this system in yeast can be faulty. We wrote a response to his letter which just opened the debate even further.

But lets start with his Letter:

Continue reading

My 2nd grade science project

My daughter’s school has a tradition for 2nd grade, to celebrate 100 days of school.

So my daughter had to prepare something relating to the number “100”. At 1st grade they celebrated 50 days of school and my wife made a cake shaped like the number 50. So this year my daughter decided to spare her mother and not bring a cake (as several other kids did).

We pitched several ideas, and she chose my idea of using the microscope to enlarge 100x certain objects (hence, the relevance to this blog 🙂 ).

A few years ago I bought a microscope at AmScope to use at home with the kids. It is a 40x-400x compound microscope, and it has a digital camera that you can connect with a USB to your laptop. This allowed us to take really nice images of the objects we examined.

I brought E. coli and yeast samples from the lab and we imaged them 100x and 400x. We then imaged a single hair from her head. She got really excited about that and also took a hair from her dog, to compare.

She then brought me salt which gave a pretty picture of the cube-shaped crystal. Last, we imaged a leaf of strawberry and mold that grew on a cucumber.

We made a nice poster:

final poster

 

This was a great experience for both of us. She was really excited and we imaged for over an hour, late in the evening. She’s interested in science, particularly medicine. She’s rational, she’s smart and clever. She will do great things when she grows up.

 

Corrigendum

It takes a lot of work to publish a paper. There’s the research itself, of course. Then there’s the writing and preparation of the figures for the paper, that are submitted to the journal for publication. Typically, most of the people who contributed to the research are involved in that, so the draft is circulated between at least 2 people, usually many more. There is one author (typically the 1st author, or the PI who is leading the research) who also needs to combine all the comments & changes into one coherent text.

The paper then goes through several rounds of  changes, following reviewers’ comments, editor’s comments, final proof-reading etc…

The accepted manuscript (MS) should be “perfect”. But mistakes can happen. Mistakes did happen for three of my recent papers.

For my Cell paper, I have prepared a nice schematic model to summarize the main highlights. It was nice, but we decided it is worth spending money on a professional artist to make a nicer one. Somehow, my scheme was uploaded for the accepted MS, instead of the more artistic one, and this is the one that was eventually  published  (you can see the artistic version in the blog post about the paper). I found that out after the fact, but since it was just a model, and the only difference was the art, we decided that publishing a correction just wasn’t worth the trouble.

Then, we published a paper in PLOS One. Only after it was published, I noticed that something happened to figure 4. Here it is:

Can you find the error?

Can you find the error?

Here, we had no choice but to publish a correction.

Recently we published a review paper, in Nature Reviews Molecular Cell biology. We worked very hard with lots of proof-reading and integrating each other’s changes to make sure it is perfect. Then, a few weeks ago, I was looking for some papers which I knew we referenced in that review. Strangely enough, those papers were mis-quoted in our review. Maybe its silly to publish a corrigendum just for a tiny mistake in references. But I think that it is important to keep science as mistake-free as possible, even with those tiny seemingly unimportant mistakes. So we published a corrigendum.

Here is also the place to thank the editors for handling these corrections pleasantly and efficiently.

I’m looking forward for my next corrigendum. What will that be?

Tracking membranes by imaging – mCLING and surface glycans

Living cells exhibit many types of membranes which participate in most biological precesses, one way or another. Imaging membranes is usually acheived by two types of reagents: chemical dyes or fluorescent proteins that are targeted to the membrane itself or inside an organelle.

The chemical dyes are usually targeted to an organelle based on a specific chemical property of that organelle.

For example:

Rhodamine 123, tetramethylrosamine, and Mitotracker  are dyes that preferentially target mitochondria, due to its membrane potential. Mitotracker has thiol groups that allow it to bind to matrix proteins, thus making it more resistant to disruption of the membrane potential (e.g. by fixation).

Lysotracker are lypophilic, mildly basic dyes, which accumulate in the acidic lysosomes.

ER-tracker is a BODIPY (boron-dipyrromethene; a group of relatively pH insensitive dyes that are almost all water insoluble) based dyes which are linked to glibenclamide – a sulfonylurease – which binds to ATP sensitive Potassium channels exclusively resident in the ER membrane.

Long chain carbocyanines like DiL, DiO and DiD are lipophylic fluorescent molecules, which are weakly fluorescent in water, but highly fluorescent when incorporetaed into membranes, particularly the plasma membrane.

FM lipophylic styryl dyes bind the plasma membranes in a reversible manner and are also incorporated into internal vesicles.

On the other hand, fluorescent proteins (FP) are targeted to membranes or organelles by fusing them to either whole proteins that localize to a specific organelle, or to short peptides that carry a localization signal. Thus, a nuclear localization signal (NLS) targets the to the nucleus, mitochondrial targeting signal (MTS) to the mitochondria and a palmitoylation signal to the plasma membrane and endocytic vesicle.

There are advantages and disadvantages to each system, relating to ease of use, specificity, photostability etc… I do not want to go into that.

Here, I would like to mention two new methods to image the plasma membrane.

Continue reading

Don’t eat this FISH-STIC

single molecule FISH (smFISH) is a great way to detect single RNA molecules in fixed cells. The “traditional” FISH uses fluorescently labeled oligonucleotides which directly hybridize with the target RNA sequence. The two most common approaches are the use of 1-5 50-mer oligos, that are labeled with 5 fluorophores, or the use of more than 20 20-mer oligos labeled at one or both ends.
A more recent approach is nicknamed “Christmas tree”, in which 2-3 rounds of hybridizations are done, with only the last round uses fluorescently labeled probed. I’ve expanded on that when I discussed RNAScope.  Other people have independently developed similar protocols (e.g. branched DNA FISH (bDNA), which was recently used to get smFISH transcriptomics in human cells).

Now, a new paper came out with a very similar method whish they call FISH-STIC (Fluorescence In Situ Hybridization with Sequential Tethered and Intertwined ODN Complexes).

However, their images do not compare to the quality shown by the RNAScope or bDNA papers. Maybe its just bad imaging and not the FISH itself, but their images look blurry and doesn’t seem to be up to modern FISH standards. I used 20-mer FISH and can get nicer images.

They also get a few high-intensity spots which the call “mRNA-independent” (since they can also see it outside the cell). To me it means that their protocol was just not optimized – either the washes, or the oligo sequences which should not make complexes like that.

They also seem to have higher background fluorescence compared to RNAScope.

Simultaneous detection of Actb and Actg mRNAs with FISH STIC probes. Source: Sinnamon & Czaplinski (2014) RNA 20:260-266.

Simultaneous detection of Actb and Actg mRNAs with FISH STIC probes. Source: Sinnamon & Czaplinski (2014) RNA 20:260-266.

On the other hand, their beta-actin FISH-STIC looks better than the RNAScope-FISH. beta-actin mRNA is found in 500-2000 copies per cell, so it is not easy to get smFISH. Still, 20-mer FISH that I do looks better than both.

20-mer FISH for beta-actin mRNA in immortalized MEFs. Source: myself.

20-mer FISH for beta-actin mRNA in immortalized MEFs. Source: myself.

Notice also in my image the transcription sites (TS) are clrearly visible, whereas I did not see any TS in their images, which is weird (unless they chose cells without TS on purpose. Don’t know why, since smFISH is a powerful tool to quantify transcription 1, 2).

They did not test FISH-STIC on a less prevalent mRNA, so I cannot comment on how it would look like compared to RNAScope or 20-mer FISH.

Anyway, the methods seem similar, but I think that the RNAScope demonstrates better results than the FISH-STIC. However, I haven’t tried this approach myself. I continue to do FISH with 20-mer probes and so far pleased with my results.

ResearchBlogging.orgSinnamon JR, & Czaplinski K (2014). RNA detection in situ with FISH-STICs. RNA (New York, N.Y.), 20, 260-266 PMID: 24345395
Battich N, Stoeger T, & Pelkmans L (2013). Image-based transcriptomics in thousands of single human cells at single-molecule resolution. Nature methods, 10 (11), 1127-33 PMID: 24097269

If you chew that mRNA, you must make a new one!

Gene expression is very complex.  My paper, which was published in Cell today, just shows that it is more complicated than previously realized.

Traditionally, eukaryotic gene expression is divided into five steps:

Continue reading

FISHing RNA for beginners

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).