Tag Archives: GFP

Poking holes into membranes to label proteins for live imaging

There are two major way to label inner proteins, structures or organnelles for live cell imaging. The most common method is fusing the studied protein to a fluorescent protein. A second approach is the addition of labeling agents from outside the cells. However, many labels cannot penetrate through the cell membrane. This is true to some, but not all dyes, but more importantly, to larger agents, such as antibodies or DNA/RNA oligos. To allow these agents to enter cells, researchers can use microinjection, electroporation, bead-loading, or transfection (e.g. of short oligos).

In a paper just published in eLife, a new technique is described to form temporary holed in the cell membrane. These holes allow delivery of any labeling agent into cells. Continue reading

Roger Tsien – the scientist that colored our research

Roger Tsien died a few days ago, at the relatively young age of 64. He was a UCSD scientist, a Nobel laureate and he was one of the first to see the significance and usefulness of GFP.

I’ve never met him. But, I guess, this blogs owes him its existence.

I don’t want to discuss his body of work, his achievements, or awards he won (e.g. the Nobel award). Many wrote nice things about him, such as here, here or here and all over the internet, with nice pictures of fluorescent proteins used in research.

I thought it will be nice to look back at his first GFP paper.


His goal in this paper was to investigate the formation of the fluorophore of GFP. Specifically, he asked:

“What is the mechanism of fluorophore formation? How does fluorescence relate to protein structure? Can its fluorescence properties be tailored and improved-in particular, to provide a second distinguishable color for comparison of independent proteins and gene expression events?”

Already here he looked to utilize GFP – to improve it, to change it, so it can be useful for fluorescent studies in biology.

He used random mutagenesis of the GFP cDNA to screen for mutants with altered brightness and emission. A simple yet powerful method, still used today, to find new FPs with exciting and useful properties.

Here is an ex/em spectral analysis of some of the mutants:


One mutant, I167T, proved to be almost twice as bright as the WT GFP protein.

But the most exciting was the finding of a blue FP (Y66H):

blue mutant

To sum up in his words:

“The availability of several forms of GFP with such different excitation and emission maxima [the most distinguishable pair being mutant P4 (Y66H) vs. mutant Pll (I167T)] should facilitate two-color assessment of differential gene expression, developmental fate, or protein trafficking. It may also be possible to use these GFP variants analogously to fluorescein and rhodamine to tag interacting proteins or subunits whose association could then be monitored dynamically in intact cells by fluorescence resonance energy transfer (19, 20). Such fluorescence labeling via gene fusion would be site-specific and would eliminate the present need to purify and label proteins in vitro and microinject them into cells.”

He saw the future, and it was bright green.

ResearchBlogging.orgHeim, R., Prasher, D., & Tsien, R. (1994). Wavelength mutations and posttranslational autoxidation of green fluorescent protein. Proceedings of the National Academy of Sciences, 91 (26), 12501-12504 DOI: 10.1073/pnas.91.26.12501

Design guidlines for tandem fluorescent timers

Almost 4 years ago, I wrote a post on tandem fluorescent timers (tFTs). The idea is to have two different fluorescent proteins fused together to the protein of interest. In the paper from 4 years ago, it was superfolder GFP (sfGFP) and mCherry. sfGFP matures very fast (within minutes) and mCherry  matures more slowly (t1/2 ~40min). The ratio beween green to red fluorescent signal indicates the percentage of new vs old proteins, thus acts as a “timer”.  This latests paper on tFTs from the same group of Michael Knop’s lab, found that analyzing tFTs might be more complicated due to some possible problems of this system.

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Imaging translation of single mRNAs in live cells

Translating the information encoded in mRNAs into proteins is one of the most basic processes in biology. The mechanism requires a machinery (i.e. ribosomes) and components (mRNA template, charged tRNAs, regulatory factors, energy) that are shared by all organisms on Earth. We’ve learned a great deal about translation over the last century. We know how it works, how it is being regulated at many levels and under varuious conditions. We know the structures of the components. We have drugs that can inhibit translation. With the emergance of next-gen sequencing, we can now perform ribosome profiling and determine exatly which mRNAs are being translated, how many ribosomes occupay each mRNA species and where these ribosomes “sit” on the mRNA, on average. New biochemical approaches like SILAC and PUNCH-P can quantifiy newly synthesized proteins & peptides. Yet, all of that information comes from population studies, typically whole cell populations. Rarely, whole transcriptome/ribosome analysis of a single cell is performed. Still, there is no dynamic information of translation, since cells are fixed and/or lysed. And there is no spatial information regarding where in the cell translation occurs (poor spatial information can be determined if cell fractionation is performed, which is never a perfect separation of organelles/regions and we are still not at the stage of single organelle sequencing).

Imaging translation in single cells is intended to provide both spatial and dynamic information on translation at the single cell and, hopefully, single mRNA molecule resolution. Recently, four papers were published (on the same day!) providing information on translation of single mRNAs. Here is a summary of these papers.

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FISEB 2014 meeting -day 1

FISEB meeting happens every three years, and it includes participants from 28 different experimental biology societies in Israel. It is the best meeting to learn about biological-medical research performed in Israel at all fields and doctrines.

4 days, 8-10 parallel sessions, hundreds of lectures, >1000 posters, >2200 participants.

The first day started by a plenary lecture by Aryeh Warshel, Nobel lauret. He is really far from my field, and his lecture was very much confusing to me. But he has nice cartoons 🙂 The bottom line – enzymes are able to catalyze reactions due to electrostatic connections that are maintained stable (unlike in water).

From the afternoon sessions, I chose “signaling pathways & networks”. Relevant to this blog:

Yoav Henis from Tel-Aviv Uni. talked about oligomerization of TGF-beta receptors. he used a method he calls “co-patching”, which is essentially IF with two different antibodies for two receptor subunits. homodimerization will yield single color “patch” whereas heterodimerization will yield an overlap of both colors (co-patch). He then looked at the % of co-patch with different receptor subunits with/without ligand, or with mutants.

Maya Schulinder from Weizmann Institute talked about the contacts between mitochondria and other organelles (ER, vacuole) in yeast. These contacts are important for lipid metabolism. She new about the mito-ER contact but found there must be a second contact (bypass mechanism). She used an interesting screen method to find the bypass mechanism to the mito-ER contact: she expressed one of the contact protein as a GFP fusion. She expected that if the bypass mechanism and the mito-ER contact “share the load” of lipid metabolism, then deletion of the bypass will increase the number of the mito-ER contacts to compensate. Using automation, she imaged 6200 deletion mutants (from the yeast deletion library) each expressing this GFP fusion. As expected, she found 4 candidates which turned out to be very interesting.

Roni Seger from Weizmann showed that targeting the nuclear localization signal of ERK can be a novel cure for certain pathologies, including certain types of cancer.

On the other hand, Maya Zigler from the Hebrew Uni. suggested another new idea to cure cancer – by inducing the surrounding immune cells to destroy the tumor.

Ido Amit from Weizmann as well told us that we may not really know all the different types of cells that exist. What most people do, particularly in immunology, is rely on one or two known “markers” and use FACS or other methods to sort the cells based on these markers. However, some of the markers overlap. and there may be cells for which we do not have any markers and they “disappear” in the crowd of unsorted cells. or, the could be further sub-types we do not know about. So he approached the problem in an unbiased way – he took all the cells in the spleen, and did single cell RNA seq to individual cells from the spleen. Thus, each cell type has several hundred/thousand “markers” based on gene expression profiles. Not only did this method agree with the common FACS sorting markers, but he identified several sub-types unknown before.  Expect his paper this month in Science. His paper just got published in Science.

Finally, Yaron Shav-tal from Bar-Ilan Uni. used the MS2 system to study how perturbing the signaling pathway of serum stimulation affects transcription of beta-actin gene. As per usual – very neat job and interesting results.

High res imaging of DNA double strand breaks: Clearing the nucleus or marking the spot

DNA can be damaged in many ways. Consequently, there are numerous mechanisms to repair it. It is a fascinating field full of innovative concepts (“DNA repair” was my favorite course during my undergrad studies).  Double strand breaks (DSBs) are considered the most genotoxic, which is why many DNA damaging drugs and treatments intended to treat cancer are intended to create DSBs. On the other hand, DSBs can lead to chromosome translocations which can promote cancer, and can actually be viewed as a hallmark of cancer cells. DSBs also occur naturally during recombination events at Meiosis, and are important intermediates in immune system development.

DSBs are recognized by repair complexes that act to mark and repair the damage. Though the sequence of event has been studied biochemically, knowledge on the in vivo temporal and spatial arrangement has been limited due to lack of good high-res methods to visualize DSBs and repair proteins at DBS sites.

Two papers recently published take two different approaches to get  high-resolution images of events at DSBs.

The first paper, from Stephen Jackson’s lab, implemented a simple method to reduce the background fluorescence in the nucleus, thus increasing the signal/noise ratio. They study the non-homologous end joining (NHEJ) process which initiates with the Ku complex recognizing the DSB, followed by recruitment of the DNA dependent protein kinase (DNA-PK) and ending with ligation by XRCC4. DNA-PK phosphorylates the histone H2AX near DSB (gamma-H2AX). This is a known marker for DSB.

They used laser illumination to create a streak of DSB, and immunofluorescence (IF) against gamma-H2AX and Ku. They used a buffer called CSK that is known to release soluble proteins, thus removing background. Though the gamma-H2AX appears as a clear streak, Ku is all over the nucleus. Ku is also an RNA binding protein. So they added an RNase to the CSK wash. Miraculously, only DNA-bound Ku was now visible.

Look at the image – it looks amazing!

Hi-res image of Ku at DSB

Addition of RNase to the CSK buffer during IF allows for high-resolution imaging of DNA repair proteins. Source: Britton et al (2013) JCB vol 202(3):579-595.

Using this simple upgrade to the protocol, which removed most of the background, and combined with 3D structured illumination (3D-SIM) super resolution microscopy*, they were able to study co-localization (here’s a video), measure distances and study composition of different components of the repair complex. They also used Ku foci count to asses the effect of different drugs on DSBs repair efficiency.

I found this paper important mostly because  this simple improvement in their protocol yielded a vast improvement in imaging. This technique could be useful for studying many other nuclear processes which involve proteins that associate with both DNA and RNA (e.g. transcription complexes).

The second paper, from Tom Misteli’s lab, took a very different approach. They wanted to see sites of chromosomal translocations  following DSBs. First, instead of getting random, or multiple DSBs using drugs, radiation or laser (as in Jackson’s paper), they introduced restriction sites for the rare-cutting IsceI restriction enzyme. One site, on chromosome 7, was surrounded by multiple repeats (256) of the LacO sequence. Other sites, on chromosomes 1 & 10, were surrounded by 96 TetO repeats. The LacO and TetO arrays can be viewed by fusion of GFP to the Lac repressor (which binds LacO) and mCherry to the Tet repressor which binds TetO. Thus, each potential DSB site is marked by a fluorescent array in distinct colors. Expression of ISceI enzyme from a plasmid induces DSB. They then followed the temporal position of the green and red spots, until they get co-localization of the colors – meaning chromosomal translocation has occurred.

Time-Lapse microscopy of cells after transfection with ISceI enzyme, showing the translocation of two distinct sites (marked by GFP and mCherry) into a co-localized spot. Source: Roukus et al. (2013) Science vol 341:660-664.

Time-Lapse microscopy of cells after transfection with ISceI enzyme, showing the translocation of two distinct sites (marked by GFP and mCherry) into a co-localized spot. Source: Roukus et al. (2013) Science vol 341:660-664.

Like in Jackson’s paper, they used this system to look at temporal and spatial events, including recruitment of proteins (tagged with a third color: BFP), and to study the effect of different mechanisms that perturb the DSB repair machinery.

In conclusion: both approaches yielded beautiful and informative images about DSBs behavior and repair. Jackson’s method has a more global and immediate application, since it can be used to study other nuclear processes. Misteli’s system requires a lot of “genetic engineering” but can provide a more precise temporal resolution (since it enables live-cell imaging, unlike Jackson’s modified IF which only uses fixed cells). Misteli’s method can also provide insight into specificity (studying DSBs at specific genomic locations). These arrays can also be used to study chromosome dynamics (i.e. movements within the nucleus, during cell division, or transport of specific genomic regions during transcription induction e.g. to the nuclear pore complex). I am actually using such an array myself, to mark the location of my gene of interest.

* 3D-SIM uses laser light that passes through an optical grating. A series of images created by these striped patterns, generated at high spatial frequency, can then be processed by a computer algorithm into a high resolution imaging. See ref. below for more details.

ResearchBlogging.orgBritton S, Coates J, & Jackson SP (2013). A new method for high-resolution imaging of Ku foci to decipher mechanisms of DNA double-strand break repair. The Journal of cell biology, 202 (3), 579-95 PMID: 23897892
Roukos V, Voss TC, Schmidt CK, Lee S, Wangsa D, & Misteli T (2013). Spatial dynamics of chromosome translocations in living cells. Science (New York, N.Y.), 341 (6146), 660-4 PMID: 23929981
Schermelleh L, Heintzmann R, & Leonhardt H (2010). A guide to super-resolution fluorescence microscopy. The Journal of cell biology, 190 (2), 165-75 PMID: 20643879

Green Fluorescent sushi

Fluorescent proteins have been isolated from invertebrate species only, until now.  A group of researchers from Japan isolated a green fluorescent protein from the freshwater eel called Unagi (yes, the same Unagi used for sushi).

The protein, named UnaG, is smaller than GFP (139 amino acids compared to 237 of GFP), excited at 498nm (after bilirubin binding) and emits light at 527nm.

UnaG is a green fluorescent protein found in eel muscle. Source: Kumagai et al. (2013) Cell 153(7):1602-1611

UnaG is a green fluorescent protein found in eel muscle. Source: Kumagai et al. (2013) Cell 153(7):1602-1611

UnaG, glows in green upon noncovalent binding to bilirubin – a membrane permeable heme metabolite.  This is a major advantage, since this mechanism can be utilized as a fluorescent switch: add bilirubin–> get fluorescence; remove bilirubin–>remove fluorescent.

This unique characteristic of UnaG prompted the researchers to develop a sensitive assay to measure bilirubin levels in blood serum – a known biomarker for several human diseases. Their assay sensitivity is 100-fold better than current clinical assays, they claim.

Another big advantage of this protein is that its fluorescence is independent of oxygen (unlike GFP-based FPs).  UnaG can therefore be used under anaerobic conditions.

UnaG fluorescence depends on bilirubin, but not on oxygen. Source: Kumagai et al. (2013) Cell 153(7):1602-1611

UnaG fluorescence depends on bilirubin, but not on oxygen. Source: Kumagai et al. (2013) Cell 153(7):1602-1611

The biological role of UnaG is still unknown, but it is suggested to have a function in oxidative stress.

I think that this is just the opening shot for the search for more vertebrate fluorescent proteins…

Read a research highlight from Nature Methods.

Unagi, from “Friends”:

ResearchBlogging.orgKumagai A, Ando R, Miyatake H, Greimel P, Kobayashi T, Hirabayashi Y, Shimogori T, & Miyawaki A (2013). A bilirubin-inducible fluorescent protein from eel muscle. Cell, 153 (7), 1602-11 PMID: 23768684