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Category Archives: Red protein
Communication between cells takes many forms. There could be communication by sending out microvesicles with important messages inside, by sending out free molecules (like hormones) or by special structures (e.g. synapses).
Sonic hedgehog (SHH) is a signaling protein that is important for the development of vertebrate limbs. It was thought to be release from a small group of cells at the posterior end of the limb bud, and is recognized by receptors on cells a long distance away.
A new paper publish in Nature from Maria Barna’s lab shows that SHH actually remains bound on the external side of the cells that produce it. The cells simply send very long thin protrusions (here named filopodia) that reach all the way to similar filopodia of the receiving cells.
I think that not only the story is very novel and interesting, but the images are very pretty.
Several “technical” issues:
In order to study the SHH signaling in live chick embryos, they designed a custom made live in ovo microscopy system: a temperature controlled plate; on it an egg container chamber, and an objective that is dipping into the yoke.
They show that standard fixation methods (e.g formaldehyde) destroy these filopodia. Also, a volume marker (in this case sfGFP) that just fills the cytoplasm does not give a strong enough signal to detect these filopodia (possibly since they are very thin, and packed with actin filaments and other proteins, so there’s very little free volume left).
So, they used palmitoylated fluorescent proteins, pmeGFP (green) and pmKate2 (red) that target them to the plasma membrane. This enabled them to visualize these very thin and long filopodia. Here’s a video movie from their paper.
They use a variety of cytoskeletal proteins fused to EGFP or to mKate2 to learn about the structure of these filopodia. Their conclusion is that these structures contain only a specialized form of actin filaments.
They show beautifully that SHH (fused to EGFP) travels to the tip of these filopodia:
They used a split GFP technology to show that SHH is actually found on the outside of the cell membrane. In split GFP, two fusion proteins are produced, each one is fused to “half” of a GFP protein (its not exactly half but let not go into that now). If the two fused proteins are in close proximity, the two halves associate to produce an GFP that fluoresce irreversibly. The two separate halves do no fluoresce. So one half was fused to SHH and the other was anchored to the extracellular leaflet of the plasma membrane, and when both were expressed, they got green fluorescence.
In total – a very nice and pretty paper.
Sanders TA, Llagostera E, & Barna M (2013). Specialized filopodia direct long-range transport of SHH during vertebrate tissue patterning. Nature, 497 (7451), 628-32 PMID: 23624372
Super-resolution microscopy can potentially allow imaging of single protein molecules. A new paper now tracks single Pol and Lig proteins in E. coli, as they repair DNA damage.
The researchers replaced the endogenous proteins with proteins tagged with a photoactivatable mCherry (PAmCherry). PAmCherry is non-fluorescent, unless activated by UV light (in this case, a 405 nm laser). By using very short pulses, they activate on average less than one molecule per cell at a time. This method is called photoactivated localization microscopy (PALM).
Using this method, they were able to follow single molecules (see this movie), count the number of Pol & Lig proteins per cell, to measure their binding rates to DNA (at optimal and DNA damage states) and importantly, to measure the timing and rates of the different steps of the base excision repair process (BER).
Their findings show that Pol & Lig diffuse by Brownian motion near undamaged DNA, but immobilize next to damaged DNA. Only a small fraction of the proteins (<5%) are involved in repair under normal conditions. But upon severe damage, the excess proteins come into play, to process >1000 DNA damage sites per minute!
The single proteins fix the damage at about 2 seconds per site. That’s pretty fast!
I think that as PALM and other super-resolution methods become more accessible and prevalent, we will be able to collect amazing data on the function of single proteins and other macro-molecules in vivo.
Uphoff S, Reyes-Lamothe R, Garza de Leon F, Sherratt DJ, & Kapanidis AN (2013). Single-molecule DNA repair in live bacteria. Proceedings of the National Academy of Sciences of the United States of America, 110 (20), 8063-8068 PMID: 23630273
So, I sat with the biophysicist who built the microscope that I intend to use. We discussed my proposed experiment and the different capabilities of that particular microscope.
Immediately we discovered that I will not be able to use LSSmKat1. If you recall, the emission peak of LSSmKate1 is at 624nm. However, the dichromatic mirrors set in that microscope for the red channels are from ~500 to 620 and from 650 to 750. So the 624 falls just in the waveband that I cannot detect.
So I remain with LSSmKate2, with em. Peak at 605.
Ok, now, the available lasers in this microscope are 407, 436, 488, 561 and 640nm.
GFP is excited at 488. That the peak and that’s good. We will use the 525/30 filter (i.e. from 510 to 540).
mCherry is excited at 587, so using the 561 laser we can get a fair signal (~60%). We will use the 593/40 filter (i.e. from 573 to 613).
This means that in both cases I collect only part of the emitted photons (the ones within the filter range, see fig 1).
LSSmKate2 can be excited with the 436 or 488 laser. The 488 will give a ~8% signal for mCherry. But the 436 will not. Although LSSmKate2 is 3-fold less bright than mCherry, and is less photostable, to object using this marker is only for time zero, when the GFP and mCherry signal should be low. However, this needs to be experimentally tested. It could be that the mCherry signal could be mathematically subtracted later.
The problem is that I would not be able to excite simultaneously at 488 and 561 to get simultaneous data of GFP and mCherry. This is because I would also have LSSmKate2 excited. However, for my purposes, I do not have to take the images simultaneously. The microscope can take images with the different lasers at 50-100ms time intervals. Since I expect to detect changes in the seconds to minutes range, a 0.1sec difference between the green and red channel is acceptable.
There is another option and that is to use a far-red protein, such as TagRFP657.
|FP||Max ex.||Max em.||QY||EC(M-1 cm-1)||Brightness||Relative brightness||Photostability|
The advantage is that we can use the 650-750 dichromatic, and use excitation at 640, which still gives a fair excitation, and it will not excite mCherry. The down side is that the 561 laser will excite TagRFP657 too. However, the TagRFP657 emission at the 593/40 filter is fairly low and with a less than maximum excitation to begin with, and a lower brightness and photostability (compared to mCherry), this should not be a big problem (see figure 2) . But again – should be tested experimentally.
So here is another lesson: when designing a multi-color experiment, you need to take into account both the FP properties, and the capabilities of your microscope (lasers, filters, dichromatics, speed etc…).
After further discussion, another LSS FP was suggested, LSSmOrange. I added its characteristics to the table above. It is brighter than TagRFP657, and the Stokes shift of LSSmOrange is better than LSSmKate2 in relation to mCherry (bluer excitation, which means NO excitation of mCherry.) So I will test all three, and hopefully have an answer which is better within 2-3 weeks.
This post continues the previous post.
I encountered a serious dilemma in choosing the right protein. This is also a great opportunity to learn about the many properties of fluorescent proteins.
Let us start with the obvious: excitation and emission maxima and spectra.
We all know that EGFP has an excitation maximum at 488nm. That is, EGFP protein that is excited with photons at 488nm will give its maximum emission intensity at its emission maximum, 509nm.
However, we must remember that this is the maximum emission. The emission spectra is much wider, and for GFP it goes from ~470nm up to ~630nm. Figure 1 shows the EGFP excitation (dashed line) and emission (full green) spectra.
And now, what if we want to look simultaneously at two colors, EGFP and dTomato?
Figure 2 (upper panel) shows you that excitation at 488nm (the max ex. of EGFP) also excite dTomato, leading to almost 30% emission at its peak of 581nm. Therefore, if we would collect all the emitted photons, we would detect the combined excitation of EGFP and dTomato.
We therefore use filters with a narrow band of wavelength (fig. 2, bottom panel). Thus, if we use the 510/20 filter, we would detect only photons emitted at the band, in this case only from EGFP. If we use the 580/30 filter, we would detect photons coming from dTomato and EGFP.
Now what happens if we add the long-stokes shift protein LSSmKate1 to the system? Figure 3 shows LSSmKate1 is maximally excited at 463nm (red dashed), and has max. emission at 624nm (full red) [note- I drew the LSSmKate spectra, based on prior publications]. However, 460nm also excites EGFP and dTomato. The EGFP signal, at 624nm is negligible. However, dTomato gives emission at 8-9% of its maximal emission. Is this negligible? We will soon learn.
Now let’s look at mCherry. With excitation laser at 460, we have virtually no excitation of mCherry. We would therefore prefer to use mCherry in our 3-color system, instead of dTomato.
This table summarizes the excitation & emission maxima:
|FP||Max ex.||Max em.|
This table shows the relative emission of the different FPs with different excitations (the LSS data is estimated based on publications, since the BD
spectrum viewer does not include LSSmKates in its database):
|% from max emission||% from max emission|
However, excitation & emission spectra are not the only considerations. There are other parameters to consider.
Brightness, QY and EC
Different fluorescent proteins have different brightness: some are very bright, some are dim. As a matter of history, we usually consider the relative brightness, compared to EGFP (which is set to 1.00). You can see in the table below the relative brightness of the proteins discussed above.
The brightness is determined by two parameters: quantum yield (QY) and Extinction coefficient (EC).
QY is simply the ration between the number of photons absorbed to the number of photons emitted. If QY=1, then for each absorbed photon you get one emitted photon. However, we never get 100% efficiency. The protein with the highest QY that I encountered is called ZsGreen with QY=0.91.
EC (or ε) relates to the formula A = εcl in which A is the absorbance, l is the path length (in cm, usually) and c the concentration in Molar units. ε, in M-1cm-1 units, is a measurement of the capability of a certain fluorescent protein to absorb light at a certain wavelength.
The formula ε*QY gives a measure of the brightness, so for EGFP, the brightness is 55,000*0.60=33,000
For ZsGreen, the EC is only 43,000 so the relative brightness is only 1.18. This example shows that a high QY does not necessarily mean a very high brightness.
This table summarizes al the brightness data:
|FP||QY||EC(M-1 cm-1)||Brightness||Relative brightness|
Let’s go back to our dilemma: dTomato or mCherry?
We now know that with a laser excitation of 460nm, we get 100% emission from LSSmKat1 at 624nm, and only 9% emission from dTomato at 624nm.
However, dTomato is 24-fold brighter than LSSmKate1. If we look only on the QY data, at 8% emission, dTomato will produce ~6 photons for each 100 photons (100*0.69*9%), whereas LSSmKat1 will produce 8 photons (100*0.08*100%). In other words, although we get very low emission of dTomato, relative to its maximum, it is almost as high as the LSSmKate1 emission in that wavelength. Since eventually, we just “see” the photons with no knowledge of their source, the signal that we will get will be ~40% from dTomato. This will create a problem if we wish to detect changes in fluorescent intensity over time due to changes in protein localization (or other changes) because we will not be able to know if the change is due to dTomato or LssmKate1.
That is why mCherry is a better choice than dTomato for this experiment.
Another parameter to take into consideration is the protein’s photostability.
Photostability is a measure (in seconds) of how long it takes for half of the number of proteins to bleach at the maximum excitation. This matter is important for any experiment involving fluorescent proteins (or dyes) but is crucial when doing live imaging, particularly for time-lapse experiments.
LSSmKate1 is more stable than LSSmKate2 (see table below), and therefore, it might be a better choice than LSSmKate2 for my time-lapse experiment.
There are other parameters to take into account, some of them I discussed in earlier posts.
pH stability (and the effect of pH on the absorption and emission spectra).
Maturation rate – how long does it take your fluorescent protein to fold correctly and create the chromophore? If you are using your protein to measure expression rate, and the maturation time is longer than the time frame of your experiment, then you will not get the information you are looking for. Maturation rate depends on protein characteristics, as well as oxygen levels and temperature. Several fast-folding FPs were developed in recent years.
Oligomerization state – many FPs are naturally monomeric. However some FPs particularly in the orange & red range) are dimeric or tetrameric. Since in many cases the FP is genetically encoded as a fusion with a protein of interest, fusing a dimeric FP may cause your protein of interest to dimerize, through the FP, thus altering its biology. If your protein is naturally oligomeric, fusing it to a dimeric or tetrameric FP may create large protein aggregates. One solution for dimeric FPs is to create a tandem fusion proteins (i.e. two FPs are fused one to the other and to the your protein).
Generally, the letter m at the beginning of the FP name indicates that it is monomeric (e.g. mCherry, mKate), d indicates it is dimeric (dTomato) and td indicates tandem dimer (tdTomato). However, there are many FPs with no indication of their oligomeric state in their name (e.g. TurboRFP is dimeric). EGFP and its many derivatives are monomeric (except at high concentrations when they form weak dimers).
Phototoxicity – although the FPs themselves do not emit harmful radiation, the excitation light can be harmful to the cell. For instance, UV light, that is used to excite BFPs, as well as some LSS FPs and photoactivatable FPs, can cause direct DNA damage, create reactive oxygen species that will cause an oxidative stress etc… This is one reason why BFPs are the least popular FPs, particularly for live imaging.
Nowadays, confocal microscopy is possibly the most widely used optical method in biological research. This methods creates better (and prettier) images than widefield microscopy (whether transmitted light or epifluorescence). The main advantages of confocal vs. widefield microscopy is the elimination of out-of-focus glare (thus increasing resolution and increasing signal-to-noise ratio) and the ability to collect serial optical sections of the specimen (z-sections).
The basic configuration of the optics is similar to that of the epifluorescnece microscope. The addition that created the confocal microscope, invented by Marvin Minsky in 1955, was to add two pinholes. The light produced by lamp (or laser) passes through the first pinhole on the way to the specimen. The light that is reflected (bright light) or emitted (fluorescent light) from the specimen passes through a second pinhole on the way to the detector (eyepiece, camera or any recording device). The two pinholes have the same focus – thus they are confocal. The light from other focal planes cannot go through the second pinhole, and this reduces the background “glare” of out-of-focus fluorescence seen in epifluorescence widefield microscopes.
Since biological samples usually have thickness of a few microns at least, one can get an image of a thin slice of the sample (e.g. 0.1 µm) without physically slicing the sample (optical section). We can then move the focus along the Z axis to get clear images of up or down sections. Thus, for a cell 3µm thick, we can have 30 hi-resolution images 0.1 µm thick from bottom to top (z-sections). These images can then be stacked one on top of the other (z-stacking) to create a single 2D image or to reconstruct a 3D image of the sample.
Here’s an example from an experiment I did last week (note that this is a widefield, not confocal microscope):
Above is a composite image of 31 Z sections of U2OS cells, create by the ImageJ program. The”pseudo-blue” represents the blue fluorescnce of a dye called DAPI (4′,6-diamidino-2-phenylindole) which intercalates into DNA, and is therefore a popular nuclear dye. When bound to DNA, it is excited by UV light (peak at 358nm) and emits blue/cyan light (peak at 461nm). The “pseudo-red” color represents the fluorescence of mCherry-ZBP1 fusion protein. mCherry is an RFP.
You can see in the image that the first and last few images in the series are out of focus. You can therefore choose the best or sharpest Z-section according to your needs.
However, most people do not show a single Z section since then we miss a lot of information that is found in other sections. The available programs today allow “stacking” the section to create a projection of all the sections into a single image.
Here is the maximum projection of the Z sections shown above:
Maximum projection means that the algorithm chooses, for each pixel, the highest value found in any of the 31 Z sections. However, since we chose all 31 sections, we can still see a “glare” or halo. This is a result of the “halos” from the out of focus sections.
I therefore choose only a few sections to create the next image:
This image is now sharper and better looking.
The program allows you other options besides maximu projection: you can choose minimum, average, median, and even standrad deviation, seen in the next image (DAPI channel only):
It looks very cool. I stacked the entire 31 sections (of a differnt field), so you can see the halo from the out-of-focus sections sorounding the “black” rim of the nucleus (black since it has the minimal standrd deviation value for all the images). The blue zones, with high SD, suggest a larger differnce in fluorescence between the differnt sections.
Above, I mentions that the blue and red are pseudo colors. What actually happened was that the images I took with the microscope at each channel (range of wavelengths) is actually maintained as a greyscale image.
Using the image analysis program you can then merge the images of the differnt channels (up to 4 in ImageJ) to create a color image. When creating the merged image, you determine what color to assign to each channel. Here is the same image, but with the colors reversed:
You should take that into account when you see pretty pictures in sceintific journals.
The program also allows to creade 3D representations of your Z stack. but I haven’t learned how to do that.
There are many other tools that one can use with the image analysis program besides creating the image. One important feture is the ability to measure the intensity of the fluorescent signal (actually, the pixels) in certain areas within the cell. You can measure distances and angles between objects and probably many moer that I still have to learn.
GFP was the first fluorescent protein to be dicovered, and subsequently used in biological research. However, by now, the biological community has found or developed an enormous number of fluorescent proteins of many colors.
According to my count (based on recent review papers) there are over 90(!) differnt fluorescent proteins. These proteins can be classified based on several charactereisitcs:
Emission color: is the most obivous classification. The classification generally goes by: Blue (424-457nm), Cyan (474-492nm), Green (499-509nm), Yellow (524-529nm), Orange (559-565nm), Red (584-610nm) and Far-Red (625-650nm).
Oligomerization: many of the FPs are monomeric (i.e. fluorece as single molecules). Others may be dimeric (two) or tetrameric (four).
Photoactivation/photoconversion: some proteins can switch there color when activated by a specific excitation wavelength. This means that the emission wavelength can change from green to red, for instance. In a few cases, the initial state of the protein is non-fluorescent, thus allowing very low background level of fluorescence. This group can be sub-divided into reversible and non-reversible photoactivatable proteins.
Fluorescnet timers – These protein change their color over time. Therefore, these can be used as “timers” for cellular processes following their activation.
Large Stokes shift (LSS): Stokes shift (named after George G. Stokes) is the shift in wavelength from excitation to emission. For most FPs, Stokes shift is less than 50nm (usually much less). For LSS proteins, the differnce is over 100nm (i.e. cells are excited by UV light or blue light and their emission is Green or Red light).
Natural vs. engineered: There is currently a lot of work invested in developing new colors and new activatable proteins by directed mutagenesis.
Three excellent review papers on the differnt kinds of FPs: