Category Archives: Super resolution

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

ASCB15 part 1

The ASCB meeting brings scientists from all levels to talk about cell biology, which is actually almost anything “biology”. But there’s also a full program dedicated to other matters, like science careers, science publishing, science communications and science policy. This is also a great venue for companies to show their products, and for organizations/institutions to recruit new members. If I remember the numbers correctly, there were over 550 oral presentations and over 2,700 posters. I overheard someone saying there were ~6000 people attending the meeting. I typically go to RNA meetings that are mostly in the lower 100’s of participants. So, to me, that’s a large meeting.


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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.

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Nobel prize in chemistry 2014

The Nobel prize in chemistry 2014 was awarded to Eric Betzig, Stefan W. Hell & William E. Moerner for the development of Super resolution fluorescent microscopy!

See here for details on the history of this discovery.

As readers of this blog know, Super-resolution microscopy has made a revolution in the field of fluorescent microscopy in a very short time.

This is a justified award.

Silicon nanocrystals – the next generation of fluorescent dyes?

Fluorescent microscopy is the only current method to follow biological structures and molecules in real-time in live specimens. Many advances were made but there are still a few problems with present fluorescent probes. Photobleaching (permanent disappearance of the fluorescent signal due to destruction of the fluorophore) is a major problem for all current fluorescent probes, be it chemical or biological. Photobleaching prevents prolong imaging.

A second problem is blinking of the fluorophore. Although blinking is actually useful in some super resolution techniques, it may still pose a problem for some applications.

Another problem is the size of some fluorescent tags such as fluorescent proteins and quantum dots. FP fusion to the studied protein can affect to protein function, simply by spatial interference.

Now, a new type of fluorophore was developed by the lab of Akihiro Kusumi from Kyoto university. Silicon nanocrystals (SiNCs) were apparently known for two decades as fluorescent, that are stable and can be used for biological applications. However, no serious effort was made to make SiNCs applicable in cell biology studies. This, he says, is due to the difficulty in producing SiNCs, manufacturing them in uniform size (and the size determines the fluorescent properties) and properties and conjugating them to biomolecules.

Kusumi developed a simplified protocol to produce mercaptosilane coated SiNC (mSiNC) that has a hydrodynamic diameter of 4.1 nm.  This was confirmed by three independent methods: gel filtration, FCS and dynamic light scattering (DLS). The hydrodynamic diameter of GFP is ~5.5nm and of Qdot655 – 19.5nm. They further measured by electron microscopy the actual size of the mSiNC core to be 3.2nm.

A & B - Elution traces of gel filtration experiments. The longer the molecule stays in the gel, the smaller it is. C- DLS analysis of mSiNC. D - FCS curves of the indicated molecules. E - image of mSiNCs obtained by electron microscopy (EM). F - distribution of measured diameter based on EM imaging.

A & B – Elution traces of gel filtration experiments. The longer the molecule stays in the gel, the smaller it is. C- DLS analysis of mSiNC. D – FCS curves of the indicated molecules. E – image of mSiNCs obtained by electron microscopy (EM). F – distribution of measured diameter based on EM imaging.

Characterization of the fluorescent properties showed that  the mSiNC fluorescent emission peak is at 655nm with a broad spectrum. Storage of dehydrated mSiNC did not affect the properties for over two months. However, fluorescence intensity decrease if stored in aqueous buffer at 37C (exponential decay time of 42 hours). Exposure of mSiNCs to excitation under fluorescent microscopy conditions showed that they are not blinking of photobleacing for over 300 minutes! Under similar conditions, GFP and the Cy3 dye photobleach within 3 seconds. the Qdot exhibited frequent blinking and fluctuations in fluorescent intensity compared to the mSiNC. mSiNCs fluorescence intensity is lower than Qdots. However, due to the instability of Qdot fluorescence, this difference diminishes over time.

After characterization came the actual test: they conjugated mSiNC to the transferrin protein (mSiNC-Tf) and applied it to cells, to follow the dynamics of cell surface transferrin receptors (TfR). whereas GFP-Tf allowed tracking for only 20 frames (0.67 sec), and Cy3-Tf allowed tracking of 150 frames (5sec), mSiNC-Tf allowed tracking for 3600 frames (120 seconds). This is quite an improvement.

A - a typical image of immobilized single mSiNC molecules. B - Distribution of fluorescence intensities of of single mSiNC spots (top) and single Qdot655 (bottom). C - Typical time-dependent changes of the fluorescence intensities of an mSiNC spot (top) and of a Qdot655 spot (bottom).

A – a typical image of immobilized single mSiNC molecules. B – Distribution of fluorescence intensities of of single mSiNC spots (top) and single Qdot655 (bottom). C – Typical time-dependent changes of the fluorescence intensities of an mSiNC spot (top) and of a Qdot655 spot (bottom).

Usually, due to blinking, single-molecule trajectories have gaps in them. These trajectories are later reconstructed based on the assumption that these gaps are due to blinking. With mSiNC-Tf, no gaps were observed (even of a single frame) for the whole 120 second (3600 frames) tracking. Thus, using mSiNC-Tf, they could follow the dynamics of TfR for long periods of time.

So, the advantages of mSiNC are very clear – they are extremely photostable, which allows for prolong imaging without loss of fluorescence. They are also small, allowing single molecule imaging with minimal perturbation. They can be conjugated to proteins.

Here are links to videos showing the lack of blinking; mSiNC-Tf on cell membrane (exhibiting Brownian motion) and what seems to be receptor internalization.

What are the disadvantages?

Well, first, the protocol to produce them, and verify their uniformity, though simplified, is yet not simple.

Second, the very wide emission spectra limits the use for multicolor imaging.

Last, similar to other dyes, it is excellent for live imaging of outer-membrane molecules, but may be more difficult for intra-cellular tagging for live imaging.

ResearchBlogging.orgNishimura H, Ritchie K, Kasai RS, Goto M, Morone N, Sugimura H, Tanaka K, Sase I, Yoshimura A, Nakano Y, Fujiwara TK, & Kusumi A (2013). Biocompatible fluorescent silicon nanocrystals for single-molecule tracking and fluorescence imaging. The Journal of cell biology, 202 (6), 967-83 PMID: 24043702

Imaging gene expression – methods & protocols

A new book in the “Methods in molecular biology” series, recently published, contains 23 imaging protocols in three major research areas: gene expression & RNA dynamics, genome & chromatin dynamics, and nuclear process & structures.

book cover: Imaging Gene Expression

This is a fairly good overview of the field and can help both beginners and researchers looking for new ideas.

The book can be freely downloaded.

Here’s the contents of the book:

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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