Category Archives: timers

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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Folding and maturation (part 3) – fluorescent timers

In the previous two parts (1)(2), I described the directed evolution of fast folding fluorescent proteins. But why is it important? Why do we need fast folding GFP? Why do we need to know the maturation time?

For most applications, it usually doesn’t matter. If we express these proteins constitutively, then we should already have enough fluorescent protein in the cells when we get to the experiment. Even in induced systems, we rarely take into account the maturation time of the protein. We follow fluorescent as it appears; usually without taking into account that the immature protein may have been present in the cells minutes or hours before.

However, when studying cellular dynamics, timing is important. The spatiotemporal dynamics of a protein is not an easy task to determine. Fluorescent timers (FT) are fluorescent proteins which change color with time due to a chemical conversion of the chromophore. Early FTs were first developed on the basis of mCherry. These FTs change color from blue to red at rates ranging from 10 min (fast-FT) to 28hr (slow FT). How are FT used? Basically, the ratio between the two colors should indicate the passage of time. For instance, a ration of red/blue=0 will indicate that the FT is a nascent protein that has not converted yet. A ratio of red/blue=1 indicates that there are no new proteins – all have converted. The ratio between 0 and 1 will tell us how much new protein still exists at the specific cellular location.

However, because of their tendency to oligomerize, or because of low brightness, these FTs were not widely used. Another recently developed monomeric green to orange FT is called Kusabira green orange (mK-GO). However, the 10h color transition is not suitable to measure fast dynamics.

A new paper sent to me by one of the blog readers (thanks Andrius) has taken a unique approach to develop FTs which they call tandem FT (tFT).

FT – regular fluorescent timer. tFT – tandem FT. m1, m2 – maturation rates. If m1<<m2 than option 2 is negligable compared to option 1.

Instead of using directed evolution to develop a novel FT, they took two known fluorescent proteins which have significantly different maturation times (sfGFP and mCherry) and fused them together in tandem. The logic of this system is quite beautiful: the sfGFP will mature fast, and the green signal will remain consistent. The mCherry will mature slower (half-time of 40min). Thus, they get a two color timer. If the signal is only green, it means that the protein was recently synthesized. A larger ratio of red/green will indicate passage of time based on mCherry maturation rate. A ratio of 1 will indicate that no new protein is present at that particular location.

They then started playing with in in different scenarios in yeast. First, they followed the formation of the spindle pole body (SPB) (the yeast equivalent to centrosomes). It is already known that in yeast, the SPB duplicates during metaphase and the “old” SPB travels to the bud tip during anaphase. They tagged an SPB protein with their tFT. Indeed, they see that the ratio of red/green in the bud is greater than in the mother cell. A mutation known to affect this segregation similarly reduces the ratio in the bud compared to the mother. In contrast, proteins that are known to be retained at the mother, whereas newly synthesized a transported to the bud show the exact opposite – i.e. red/green signal higher in the mother vs. bud cell.

A very neat experiment used a bud-scar protein tagged with tFT which shows several bud scars forming on the same cell over time, with different color blends (the old scar is red, newer in “yellow” and newest in green. This presents a nice “clock” of for the relative age of each scar. It’s too bad that the authors didn’t add the real time line (in minutes) in their image and tried to correlate the actual time with the tFT “time”.

A protein localized to bud scars is fused to tFT. the older the scar, the more red it is. Source: Khmelinski et al. (2012) Nat Biotech 30:708.

They then started to explore other stuff. First, they looked at the segregation of nuclear pores during mitosis. Nuclear pores (NPC) are complex structures that allow transfer of proteins and RNA in and out of the nucleus in a regulated manner. The question they asked is whether “old” NPCs remain in the mother cell and the bud get newly synthesized proteins, or vice versa (or equal distribution, i.e. non discriminated transfer). They individually tagged each of the NPC proteins (plus some control) – a total of 36 proteins- with their tFT and measured whether we see older proteins at the mother or the bud. Surprisingly, the bud gets the older proteins. Their analysis suggests that there is active transport of the “old” NPCs into the bud. How this is achieved and why the bud should receive old and possibly damaged NPCs are good questions for further research.

NPC proteins tagged with tFT show preferential localization of “older” proteins at the bud compared to mother cell. source: Khmelinski (2012), Nat Biotech. 30:708.

Despite of their interesting results, the authors didn’t stop there and when to test another application. They show that a single readout of the red/green ratio can indicate the stability of the protein. Stabilization/de-stabilization by mutations or different inherent stability can easily be distinguished based on the red/green ratio. They further utilize this approach to screen for protein stability regulators, which was quite successful as they identified most known factors in the specific pathway they studied, as well as identified new factors in this pathway.

In conclusion – this paper shows how one can utilize “ordinary” fluorescent proteins as fluorescent timers. Moreover, this method is much easier and much more flexible than trying to evolve FTs by random or directed mutagenesis. In fact, using different pairs with different maturation times would easily enable us to create FTs for any time intervals we wish and in any color we wish.

They show here how to utilize this system to follow cell cycle events. Obviously, this system can be used to study other scheduled, localized events in the cell.

A straightforward application is to follow protein stability. However, this system can also be used to study mRNA stability, by combining these tFTs with the MS2 system.

I’m sure other applications will be developed in the future.
All in all, a good paper with a great idea! Khmelinskii A, Keller PJ, Bartosik A, Meurer M, Barry JD, Mardin BR, Kaufmann A, Trautmann S, Wachsmuth M, Pereira G, Huber W, Schiebel E, & Knop M (2012). Tandem fluorescent protein timers for in vivo analysis of protein dynamics. Nature biotechnology, 30 (7), 708-14 PMID: 22729030

All the colors of the rainbow

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).

Bacteria expressing differnt FPs were plated to create a nice picture (source: Roger Tsien lab)

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:

Stepanenko et. al. (2008) “Fluorescent proteins as biomarkers and Biosensors: Throwing color lights on molecular and cellular processes” Curr. Protein. Pept. Sci. 9(4):338.

Chudakov et. al. (2010) “Fluorescent proteins and their applications in imaging living cells and tissues”  Physiol. Rev. 90:1103.

Wu et. al. (2011) “Modern fluorescent proteins and imaging technologies to study gene expression, nuclear localization, and dynamics” Curr. Opin. Cell. Biol. 23:310.