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Archive of Green Fluorescent Blog
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Category Archives: Yellow protein
Fluorescent imaging is all about the contrast between the signal and the background. For imaging to be successful, the signal should be clear above the background. Background fluorescence can come from free/non-specific fluorescent probe, autofluorescence, and out of focus fluorescence.
There are two major strategies to improve signal/background ratio.
The first is to increase the signal. We do that by choosing brighter fluorescent molecules, by increasing the number of fluorescent probes per target, by using more than one color per target, by having photoactivatable probes etc…
The second strategy is to reduce the background. The wash step in IF and FISH protocols is intended to remove excess, non-specific bound, probe. There are even more extensive wash protocols. We have many type of microscopes that are designed to reduce out-of-focus light (these include confocal, TIRF, multi-photon, and SPIM). In yeast imaging, we sometimes add an excess of adenine to the culture media, since many strains are defective in adenine biosynthesis, and accumulate a red intermediate molecule. In the field of single molecule live mRNA imaging, we usually add a nuclear localization signal (NLS) to the fluorescently tagged RNA binding protein, in order to reduce its cytoplasmic fluorescence.
Now, my lab-mate Bin Wu develop a system that he calls “Background free imaging of single mRNAs in live cells using split fluorescent proteins”.
The idea is to combine the two most common systems – the MS2 and the PP7 systems, so that the MS2 binding sequence (MBS) and PP7 binding sequence (PBS) will be in tandem. Then the MS2 coat protein (MCP) will be fused to one half of a fluorescent protein (Venus) and PCP will be tagged with the other half. Only when MCP-VenusN and PCP-VenusC are in very close proximity (e.g. bound to the MBS and PBS, respectively) the two halves can bind to form Venus, which fluoresce in bright green-yellow. Add 12 of these tandem repeats to the mRNA and you have 24 fluorescent proteins on the mRNA in the cytoplasm, with, theoretically, zero unbound fluorescent protein in the cytoplasm, hence “zero background”.
The system has some limitations. For one thing, the protein levels must be low, since the fluorescent protein halves can self-associate at high concentrations independent of interaction with the mRNA. Also, since it takes the fluorescent protein some time to mature, it is not useful to study short-lived mRNAs, or transcription in live cells, since by the time it matures, the mRNA has already left the nucleus.
Wu B, Chen J, & Singer RH (2014). Background free imaging of single mRNAs in live cells using split fluorescent proteins. Scientific reports, 4 PMID: 24402470
The conditions used for microscopy are often not “physiological” conditions. If we are talking about live imaging, then the cells are usually in culture, placed on a glass surface and grown in an artificial media. In many cases, we use genetically encoded fluorescent markers, that are rarely inert. These are acceptable and known limitations of the system.
However, when we think about microscopy, we do not often consider to effect of the light itself. The light we use can have deleterious effect. For instance, UV light is known to cause damage to cells, e.g. creating reactive oxygen species (ROS), creating thymidine dimers, cross-linking macromolecules and probably more. That is why people tend to limit the use of fluorescent proteins that are excited by UV light (such as blue fluorescent proteins or some photoactivated proteins).
Yet, visible light is considered relatively harmless. Now, a new study suggests that visible light, particularly blue-green light (which is used to excite green-yellow fluorescent proteins), can affect the metabolic state of the cell. How?
Well, all cells contain light absorbing molecules. Some molecules function as light sensors or light energy harvesting molecules: Chlorophyll, cryptochromes, phytochromes, photoreceptors, rhodopsins, and fluorescent proteins. Other molecules absorb light, e.g. pigments.
However, some cells do not have obvious light sensitive molecules. For example, the budding yeast. In this paper, Carl H. Johnson’s lab looked at the effect of visible light on yeast respiratory oscillations (YRO) and oxidative stress. Apparently, under some conditions, yeast show 1-6hr oscillations of oxygen consumption and metabolite productions. surprisingly, shining white, blue or green light (but not red) has shortened the cycle and decreased the amplitude of oxygen consumption. There was also increased expression of antioxidant enzymes, and increased light sensitivity to oxidative-stress mutants.
Based on several other assays, the authors suggests the blue-green light harms cytochromes in the respiratory electron transport process. Photoinhibition of the electron transport causes accumulation of ROS and oxidative stress response. It is hypothesized that the shortened oscillation periods and reduced amplitude of oxygen consumption (= less respiration) is a cellular mechanism to reduce the photoinhibition to electron transport, thus reduce oxidative damage.
What does this mean to microscopists?
It only suggests that long term exposure to light (either in prolonged live imaging, or just the incubation conditions) can affect the metabolic state of the cell and therefor may have either deleterious effect on the health of the cell, or create false positive or false negative results, particularly in experiments that are designed to study respiration and oxidative stress. Obviously, if you have light sensors or light harvesting molecules, their absorbance wavelengths need to be taken into account.
Practically, it is always a good advice to minimize the exposure to light (which we do anyway to reduce photobleaching).
Robertson JB, Davis CR, & Johnson CH (2013). Visible light alters yeast metabolic rhythms by inhibiting respiration. Proceedings of the National Academy of Sciences of the United States of America, 110 (52), 21130-5 PMID: 24297928
A new and improved green fluorescent protein, named mNeonGreen, was developed.
It was engineered from a Yellow fluorescent protein (LanYFP) that was isolated from the cephalochordate Branchiostoma lanceolatum. Therefore, LanYFP is genetically unrelated to the commonly used Aequorea victoria GFP.
Directed evolution to make it a monomer produced the new, monomeric protein, mNeonGreen. The ex/em of mNeonGreen is slightly shifted to the yellow compared to EGFP (509/516 compared to 488/509), making it a better choice to separate from CFP emission.
Though slightly less brighter than its parent protein LanYFP, mNeonGFP is 2-3 times brighter than EGFP and actually brighter than most green & yellow proteins.
Another great advantage of this new protein is that is it fast folding – the authors claim it is <10 min at 37C. This is fairly close to the superfolder GFP.
It is also very photostable (comparable to EGFP), performs well as a fusion construct at N & C termini many tested proteins, performs 4-times better that EGFP in stochastic single-molecule superresolution imaging and is a better FRET partner (both as acceptor and donor) than other proteins.
In short, this may very well be the “next generation” of fluorescent proteins. It has all the good qualities, and seems to have none of the bad ones. It performs better than most, if not all fluorescent proteins in every tested parameter.
Only its name is rather plain. I would call it something like wonderGFP or GreenLantern (hey, it even has the Lan from the animal they developed it from).
(Update: see here for details on how to get your hands on this protein).
Shaner NC, Lambert GG, Chammas A, Ni Y, Cranfill PJ, Baird MA, Sell BR, Allen JR, Day RN, Israelsson M, Davidson MW, & Wang J (2013). A bright monomeric green fluorescent protein derived from Branchiostoma lanceolatum. Nature methods, 10, 407-409 PMID: 23524392
In part 1 I discussed the directed evolution of fast-folding GFPs. These were developed for specific purposes of improving the solubility, stability and folding of the protein. Now, I will discuss the maturation step and how it was measured for a variety of GFPs.
As mentioned in the first part, maturation is the final step in the transformation of GFP from a chain of amino acids to a fluorescent protein – the creation of the chromofore. The chromofore has a very long name (p-hydroxybenzylideneimidazolidone) which in wild type GFP is formed from amino acids S65, Y66 and G67. In this process, the amide nitrogen of G67 backbone performs a nucleophilic attack on the carbonyl carbon of S65. Oxidation with atmospheric O2 and dehydration reactions create the imidazolinone ring, which is conjugated to the side chain of Y66.
In most papers that measure GFP maturation, the authors do not distinguish folding from maturation. In many cases, the experiments involve complete denaturation of the protein in vitro (e.g. by urea or guanidinium chloride) and then measuring fluorescence following re-folding. In vivo, it would be difficult to assess folding kinetics (because you do not really have an exact time zero for when the protein is being translated). [In our lab, we are trying to develop a system to visualize translation in vivo in real time by imaging techniques. If it works, it would be useful for determining exact folding & maturation rates in vivo of a single fluorescent protein].
In any case, it is easier to measure the kinetics of the maturation step, both in vivo and in vitro, simply by removing and adding oxygen. I mentioned in part 1 that this is how the maturation rate of GFP-S65 was calculated in vivo. A new paper from 2011 from the lab of Funatsu analyzed the maturation rates of several GFPs in vitro. Their idea was simple – using a commercial kit, the performed in vitro transcription-translation reaction of the FPs at anaerobic conditions (they added catalase and glucose oxidase to get 0.1mg/l of oxygen). They then stopped the reaction and added oxygen, then followed fluorescence.
The proteins they assayed were: wild-type GFP, GFP-S65T, GFP-S65T/S147P, EGFP, sfGFP, Emerald, GFPm, GFPmut2, GFPmut3, sgGFP, “cycle 3”(a.k.a GFPuv), frGFP (a.k.a GFPuv3), GFPuv4 and 5 and two yellow FPs: EYFP and Venus..
Their results are interesting. As expected, the S65T mutation improved the maturation rate by 3.2 fold. However, EGFP matured only 20% faster than the S65T alone. The S65T/S147P is a variant that is stable under a range of temperatures. This mutant matured faster than EGFP, at a rate 40% faster than S65T alone. GFPmut2 and GFPmut3 (S65G/S72A) both showed a 7-fold faster maturation than WT GFP (>2-fold the S65T alone). sgGFP (SuperGloGFP, F64L/S65C/I167T, from Qbiogene) showed improved maturation rate compared to S65T (70% faster). Emerald, which contains 4 mutations on top of the EGFP mutations showed only a slight increase in maturation rate (similar to S65T/S147P). “cycle 3” was only slightly better than WT GFP, frGFP (which is cycle 3 +EGFP mutations combined) showed a maturation rate which was 10% less than EGFP. Addition of the I167T mutation to create GFPuv5 increased the maturation rate by ~70% (just like in sgGFP).
Most interesting is the super-folder GFP (sfGFP), which showed a maturation rate of only 2.3-fold over the WT (that is ~70% of that of S65T). Thus, though this protein may be more stable and may fold very fast compared to other variants, the important step of maturation is the slowest among all variants tested (except the WT). Since folding assays measure fluorescence as the output for a mature protein, it means that the folding step (prior to maturation) is much faster than previously appreciated.
GFPm is a weird case. GFPm, developed by David Tirrell, is a variant with mutations from cycle 3 and GFPmut3. However, Tirrell tried something unique – to replace the leucines in the protein with 5,5,5- tri-fluoroleucine (tfl). The fluoreinated form proved to be insoluble, and fluorescent of the cells (E. coli) was 500-fold less than the regular protein. They then went on with mutagenesis, developing new variants which were brighter (up to 650-fold over GFPm). Personally, I do not understand how this can be of much use, since using tfl will probably have major effects on every aspect of cell biology we are interested in. Anyway, the maturation rate of GFPm (not fluoreinated) is itself pretty high – almost 3-fold over S65T and is actually the fastest maturing GFP variant tested. Oddly, they do not discuss this result anywhere in the text. Perhaps there is a technical issue they wanted to avoid?
Last but not least, the two YFPs showed maturation rates which are similar to WT GFP (Venus) or ~2.3-fold better (EYFP).
So how does all this information help us?
First, if we know which mutations enhance maturation and which slow it down, we can design faster-maturing proteins.
Second, we can use this data to estimate translation or translocation rates in vivo. However, we should remember that the data obtained in vitro (at 37C) does not neccessarily agree with actual maturation rates in vivo in every cell type. for instance, yeast or fly grow in colder temperatures which may affect maturation. Oxygen levels in tissue culture dish are differnt than in entire animals, and also differnt in differnt organs. Also, if the GFP is fused to another protein, it may also affect folding as we learned in part 1, as well as maturation. Finally, the in vitro environment in the tube lacks many biomolecules (proteins other than used for translation, small molecules, ions, oxidizing molecules, antioxidants etc) which can affect oxygen availability in the immediate environment of the newly translated GFP.
Third, we can use such data to design cool experiments. So… stay tuned for part 3, which I will dedicate to biological timers.
Iizuka R, Yamagishi-Shirasaki M, & Funatsu T (2011). Kinetic study of de novo chromophore maturation of fluorescent proteins. Analytical biochemistry, 414 (2), 173-8 PMID: 21459075
Yoo TH, Link AJ, & Tirrell DA (2007). Evolution of a fluorinated green fluorescent protein. Proceedings of the National Academy of Sciences of the United States of America, 104 (35), 13887-90 PMID: 17717085
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: