Fluorescent protein quick guide

Living Colors AmCyan1 is a cyan fluorescent protein that was isolated from the coral reef organism Anemonia majano.  It is a very bright cyan, making it a good reporter and useful in multicolor analyses. It has been engineered for brighter fluorescence and higher expression in mammalian cells.

ZsGreen1 is our brightest green fluorescent protein, significantly brighter than EGFP. It is ideal for multicolor labeling applications.

AcGFP1 is a monomeric green fluorescent protein with 94% homology to EGFP at the amino acid level. It has been widely validated as a fusion tag with a wide variety of proteins with diverse functions and subcellular locations, as well as for multicolor labeling with DsRed-Monomer.

ZsYellow's true yellow emission makes it ideal for multicolor labeling applications.

mBanana is widely separated from our other yellow fluorescent protein, ZsYellow1. Its t_0.5 for maturation at 37°C is approximately 1 hour.

mOrange is an extremely bright orange fluorescent protein. Its high extinction coefficient and quantum yield make it an ideal FRET acceptor. Its t_0.5 for maturation at 37°C is approximately 2.5 hours. The original mOrange has been discontinued. mOrange2 is a variant of mOrange which has been modified for improved stability.

Red fluorescent proteins are often used for in vivo applications due to the relatively high penetration depth of red light and the lack of overlap with endogenous fluorophores which tend to be in the green wavelengths.

Monomeric fluorescent proteins such as DsRed-Monomer, mCherry, and mStrawberry are often ideal for fusions, as they tend to be the least likely to disrupt protein function. Fusion proteins containing mCherry have been reported in a variety of organisms and used for quantitative imaging techniques including FRET, FRAP, and FLIM. mCherry matures extremely quickly: its t_1/2 for maturation at 37°C is approximately 15 minutes, as compared to mStrawberry's maturation t_1/2 of 55 minutes.

DsRed-Express and DsRed-Express2 are ideally suited for flow cytometry, with an extremely low level of green autofluorescence. They can be used in combination with green or far red-shifted fluorescent proteins and have a maturation rate similar to that of EGFP.

DsRed2 is optimized for high solubility and low aggregation, making it highly amenable to fluorescence microscopy studies. DsRed2 has a high signal-to-noise ratio and distinct spectral properties, and is a good choice for multicolor labeling experiments.

tdTomato is our brightest red protein. Its t_1/2 for maturation at 37°C is approximately 1 hour. Although its tandem dimer structure means that it has a high molecular weight, it has been specifically developed to be non-aggregating.

Far red proteins such as HcRed1, mRaspberry, and mPlum are preferred for in vivo imaging since they avoid the natural green autofluorescence produced by plant and animal cells.

HcRed1 has a low tendency to form aggregates in living cells, and it can be detected just 16 hours after transfection via flow cytometry—a maturation rate comparable to that of EGFP.

mRaspberry's t_0.5 for maturation at 37°C is approximately 55 minutes, while mPlum's t_1/2 for maturation at 37°C is approximately 100 minutes.

E2-Crimson matures rapidly—its t_1/2 at 37°C is 26 minutes—and is very bright, photostable, and soluble. Its high solubility leads to low cytotoxicity. It is well-suited for in vivo applications involving sensitive cells such as primary cells and stem cells.

PAmCherry and Dendra2 are photoswitchable proteins. PAmCherry is nonfluorescent until activated by a short exposure to 350–400 nm light, while Dendra2 is a monomeric, green-to-red photoswitchable fluorescent protein. Once it is irreversibly switched to its red form, Dendra2 is highly photostable. Switch on these fluorescent proteins in a subset of cells, proteins, or organelles in order to track their movement.

Use the Timer fluorescent protein to follow the on and off phases of gene expression. Initially, the Timer fluorescent protein emits green fluorescence, but as time passes, the fluorophore undergoes additional changes that shift its fluorescence to longer wavelengths. When fully matured, the protein is bright red.

Monomeric fluorescent proteins are often ideal for fusions as they tend to be least disruptive to the function of the protein. In many cases, oligomers can also be effective choices. We offer N- and C-terminal vectors for creating fusion constructs.

mCherry fusion constructs

mCherry has been successfully fused to several proteins, including actin and tubulin (Figure 1). mCherry fusions have been reported in Arabidopsis (Song et al. 2007), zebrafish (Pisharath et al. 2007), E. coli (Pradel et al. 2007), HIV virions (Campbell et al. 2007), and yeast (Snaith, Samejima, and Sawin 2005). These fusions have also been used for quantitative imaging techniques including fluorescence resonance energy transfer (FRET) (Anderson et al. 2006), fluorescence recovery after photobleaching (FRAP) (Picard, Suslova, and Briand 2006), and fluorescence lifetime imaging microscopy (FLIM) (Tramier et al. 2006).

Figure 1. mCherry fusion constructs. HeLa cells were transiently transfected via a lipid-based method, with mammalian expression vectors encoding mCherry fused to either human cytoplasmic actin (Panel A) or tubulin (Panel B). Cells were fixed using 4% paraformaldehyde and imaged 36 hr posttransfection with a 40X objective on a Zeiss Axioskop microscope using the 575/50, 610, 640/50 filter set.

E2-Crimson

E2-Crimson was derived from DsRed-Express2 and retains DsRed-Express2's fast maturation, high photostability, high solubility, and low cytotoxicity (Strack et al. 2008). It is well-suited for in vivo applications involving sensitive cells such as primary cells and stem cells:

• Extremely bright
• Highly soluble
• Fastest maturing red or far-red protein—half-time of 26 minutes at 37°C
• Efficient excitation with standard far-red lasers
• Suitable for multicolor experiments

Figure 2. E2-Crimson is useful for confocal and STED (stimulated emission depletion) microscopy. The mammalian ER was imaged by conventional confocal microscopy (left) or by STED microscopy (right) with 635 nm excitation and a STED wavelength of 760 nm. The scale bar is 1 µm.

DsRed-Monomer

DsRed-Monomer is a true monomeric mutant of our red fluorescent protein from Discosoma sp. reef coral, which makes it the optimal choice for use as a red fluorescent fusion tag. DsRed-Monomer is well-tolerated by mammalian cells and has been successfully used to create stably transfected clonal cell lines. The DsRed-Monomer chromophore matures rapidly and is readily detected 12 hours after transfection and the fluorescent protein is extremely stable, allowing you to monitor fluorescence over extended periods of time.

An ideal fusion tag

Ideally, when you label a protein of interest, the fluorescent tag itself should not interfere with the biological function of the target protein. If the fluorescent protein has a strong tendency to form oligomers, it is more likely to alter or hinder the original function of the tagged protein. Because DsRed-Monomer is a true monomer, it is the optimal choice for use as a red fluorescent fusion tag. When expressed in mammalian cells, the protein is highly soluble and homogeneously distributed within the cytosol, with no detectable aggregation (Figure 3).

Figure 3. DsRed-Monomer is soluble and displays even, consistent, and homogeneous distribution in HeLa cells. DsRed-Monomer has been expressed as a fusion with a large panel of diverse proteins with diverse functions and subcellular locations. The localization of the resulting tagged protein was monitored and all the tested proteins localized properly. For example, the DsRed-Monomer-Actin fusion protein correctly incorporates into the actin filament system of the cytoskeletal network, ruffling edges, and filipodia.

A true monomer

The monomeric nature of the DsRed-Monomer protein (28 kDa, calculated molecular weight based on amino acid sequence) has been confirmed by two independent methods:

• FPLC gel filtration chromatography of recombinant DsRed-Monomer yields a single elution peak at a retention time consistent with a 28-kDa molecular weight. The elution profile does not display higher molecular weight species and provides strong evidence that DsRed-Monomer is a true monomer. In contrast, recombinant DsRed-Express protein elutes earlier than DsRed-Monomer because of its tetrameric structure.
• Pseudonative gel electrophoresis yields a fractionation profile that is consistent with a monomeric protein, and similar to the monomeric green fluorescent protein AcGFP1.
  
  

Figure 4. Living Colors DsRed-Monomer is a monomeric protein. Panel A. Recombinant DsRed-Express and DsRed-Monomer (100 µg) were analyzed by FPLC gel-filtration chromatography. Overall absorbance (A₂₈₀) and chromophore excitation (A₅₅₇) of the eluted material were monitored simultaneously. DsRed-Monomer elutes from the column at a retention time (39 min) corresponding to a molecular weight of 28 kDa. The calculated molecular weight of DsRed-Monomer is 26.8 kDa. DsRed-Express is a tetrameric protein that elutes at an earlier retention time (33 min) corresponding to a molecular weight of 89 kDa. Panel B. Pseudonative gel analysis of proteins. The oligomeric structure of proteins is preserved during SDS-PAGE analysis if samples are kept at 4°C and not boiled prior to loading on a gel. Boiled and unboiled recombinant proteins (7.5 µg) were separated by SDS-PAGE electrophoresis (12% acrylamide). In both the boiled (denatured) and unboiled (nondenatured) samples, DsRed-Monomer runs as a uniform band of ~30 kDa due to its monomeric structure. The unboiled (nondenatured) DsRed-Express runs at a much higher molecular weight than its boiled (denatured) counterpart due to its tetrameric structure.

Spectral properties

DsRed-Monomer preserves key spectral features of other DsRed variants, in particular, DsRed-Express. DsRed-Monomer has an excitation maximum of 556 nm and an emission maximum of 586 nm. Its spectral profile is virtually identical to our other DsRed fluorescent protein variants, allowing DsRed-Monomer to be detected using existing standard filter sets or with custom-tailored, optimized sets such as those available from Chroma Technology Corporation (see http://www.chroma.com/ for details). Although DsRed-Monomer is somewhat less bright than DsRed-Express, it is nevertheless an excellent choice for fluorescence microscopy imaging and flow cytometry. As with our other red fluorescent proteins, DsRed-Monomer performs well when multiplexed with compatible fluorescent proteins such as AcGFP1 (Figure 5).

Figure 5. Fluorescence excitation and emission spectra of DsRed-Monomer and AcGFP1.

References

Anderson, K. I., Sanderson, J., Gerwig, S. & Peychl, J. A new configuration of the Zeiss LSM 510 for simultaneous optical separation of green and red fluorescent protein pairs. Cytometry. A 69, 920–9 (2006).       

Campbell, E. M., Perez, O., Melar, M. & Hope, T. J. Labeling HIV-1 virions with two fluorescent proteins allows identification of virions that have productively entered the target cell. Virology 360, 286–93 (2007).

Picard, D., Suslova, E. & Briand, P.-A. 2-color photobleaching experiments reveal distinct intracellular dynamics of two components of the Hsp90 complex. Exp. Cell Res. 312, 3949–58 (2006).

Pisharath, H., Rhee, J. M., Swanson, M. A., Leach, S. D. & Parsons, M. J. Targeted ablation of beta cells in the embryonic zebrafish pancreas using E. coli nitroreductase. Mech. Dev. 124, 218–29 (2007).             

Pradel, N. et al. Polar positional information in Escherichia coli spherical cells. Biochem. Biophys. Res. Commun. 353, 493–500 (2007).

Snaith, H. A., Samejima, I. & Sawin, K. E. Multistep and multimode cortical anchoring of tea1p at cell tips in fission yeast. EMBO J. 24, 3690–9 (2005).             

Song, L., Han, M.-H., Lesicka, J. & Fedoroff, N. Arabidopsis primary microRNA processing proteins HYL1 and DCL1 define a nuclear body distinct from the Cajal body. Proc. Natl. Acad. Sci. U. S. A. 104, 5437–42 (2007).     

Strack, R. L. et al. A noncytotoxic DsRed variant for whole-cell labeling. Nat. Methods 5, 955–7 (2008).       

Tramier, M., Zahid, M., Mevel, J.-C., Masse, M.-J. & Coppey-Moisan, M. Sensitivity of CFP/YFP and GFP/mCherry pairs to donor photobleaching on FRET determination by fluorescence lifetime imaging microscopy in living cells. Microsc. Res. Tech. 69, 933–9 (2006).       

AcGFP1: A true GFP monomer

AcGFP1 is an engineered, fluorescent mutant of the wild-type protein derived from the jellyfish Aequorea coerulescens. It is a true monomer that does not form dimers as has been reported for AvGFP and EGFP (Jain et al. 2001; Chen et al. 2002). AcGFP1 is a superior alternative to Aequorea victoria GFPs (EGFP and AvGFP), especially if you are looking for a true monomer for fusion applications (Gurskaya et al. 2003). Its open reading frame has been human codon-optimized to increase the translational efficiency of the AcGFP1 mRNA, which results in higher expression in mammalian cells. The AcGFP1 protein is stable, allowing you to monitor fluorescence over extended periods of time. The chromophore matures rapidly and is readily detected 8–12 hours after transfection. As with all of our fluorescent proteins, AcGFP1 is well-tolerated by mammalian cells and has been successfully used to establish stably transfected clonal cell lines.

AcGFP1 has an excitation maximum of 475 nm and an emission maximum of 505 nm. Its brightness and spectral properties allow AcGFP1 to be detected using existing filter sets for FITC. This includes standard factory-installed microscope filters as well as custom-tailored, optimized sets such as those available from Chroma Technology Corporation (see www.chroma.com for details). AcGFP1 can be easily detected via both Western blot and immunoprecipitation applications with our wide array of Living Colors antibodies.

Ideal for visualizing & tracking your protein of interest with fusions

AcGFP1 has been widely validated as a fusion tag, for a wide variety of proteins with diverse functions and subcellular locations (Figure 1, Panel C). AcGFP1 is particularly suited for use in multicolor applications (e.g., to simultaneously visualize the subcellular localization of two proteins of interest). It also performs well in cell-based assays that monitor protein subcellular trafficking (Figure 1, Panels A and B). Cells expressing AcGFP1 are easily detected and sorted by flow cytometry.

Figure 1. Use of AcGFP1 for fusions and fluorescence microscopy applications. Panels A and B. Activation of Protein Kinase C alpha was monitored with Living Colors AcGFP1. Panel A. HEK 293 cells were stably transfected with a plasmid encoding AcGFP1 fused to PKC alpha. Panel B. Cells were induced with 1.5 µg/ml PMA for 3 min. The PKC alpha-AcGFP1 fusion moves from the cytosol to the plasma membrane, a result consistent with the known mobilization pattern of PKC alpha. Panel C. HeLa cells were transiently transfected with pAcGFP1-Actin and visualized by fluorescence microscopy.

A true monomer

The monomeric nature of the AcGFP1 protein (26.9 kDa, calculated molecular weight based on amino acid sequence) has been confirmed by three independent methods:

• FPLC gel-filtration chromatography of recombinant AcGFP1 yields a single elution peak (Figure 2, Panel A).
• Sucrose density gradient ultracentrifugation yields a fractionation profile consistent with a monomeric protein (Figure 2, Panel B).
• Pseudonative gel electrophoresis of recombinant AcGFP1 protein in comparison to an oligomeric fluorescent protein supports the same conclusion (Figure 2, Panel C).

Figure 2. AcGFP1 is a monomeric protein. Panel A. Recombinant AcGFP1 protein was analyzed by FPLC gel-filtration chromatography. Overall protein absorbance (A₂₈₀) and chromophore excitation (A₄₇₇) of the eluted material were monitored simultaneously. AcGFP1 elutes from the column at a retention time corresponding to a molecular weight of 24 kDa. The calculated molecular weight of AcGFP1 is 26.9 kDa. Panel B. Recombinant AcGFP1 protein was analyzed by sucrose density ultracentrifugation using a continuous gradient. Panel C. Pseudonative gel analysis of proteins. The oligomeric structure of proteins is preserved during SDS-PAGE analysis if samples are kept at 4°C and not boiled prior to loading on a gel. Boiled and unboiled recombinant proteins (7.5 µg) were separated by SDS-PAGE electrophoresis (12% acrylamide). In both the boiled (denatured) and unboiled (nondenatured) samples, AcGFP1 runs as a uniform band of ~30 kDa due to their monomeric structure. The unboiled (native) DsRed-Express runs at a much higher molecular weight than its denatured (boiled) counterpart due to its oligomeric structure.

References

Chen, Y., Müller, J. D., Ruan, Q. & Gratton, E. Molecular brightness characterization of EGFP in vivo by fluorescence fluctuation spectroscopy. Biophys. J. 82, 133–44 (2002).  

Gurskaya, N. G. et al. A colourless green fluorescent protein homologue from the non-fluorescent hydromedusa Aequorea coerulescens and its fluorescent mutants. Biochem. J. 373, 403–8 (2003).           

Jain, R. K., Joyce, P. B., Molinete, M., Halban, P. A. & Gorr, S. U. Oligomerization of green fluorescent protein in the secretory pathway of endocrine cells. Biochem. J. 360, 645–9 (2001).        

Detecting protein-protein interactions requires a good (high-efficiency) FRET pair: a donor with a high quantum yield (Q_D) and an acceptor with a high Förster radius (R₀).

The following four pairs have been reported in the literature to be suitable for FRET:

*mOrange has been discontinued. mOrange2 is a variant of mOrange which has been modified for improved stability.

Our subcellular localization vectors allow you to target a wide range of structures such as:
actin, autophagosome, cytoskeleton, endosome, ER, Golgi apparatus, mitochondria, nucleus, peroxisomes, plasma membrane, and tubulin.

The subcellular localization vectors encode fusions of fluorescent protein variants and localization signals or subcellular structural proteins, which target the fluorescent protein to a specific organelle or subcellular structure. The vectors are available in a variety of organelle- and cytoskeleton-targeted color variants. They are ideal for multiplex labeling experiments. 

On-demand protein stabilization using the Lenti-X ProteoTuner Systems to visualize rapid events 

Eukaryotic cells have the ability to quickly change their shapes, for example, while moving toward the site of an infection or during intracellular transport of macromolecules, membrane vesicles, and organelles. These events are organized via a subcellular system called the cytoskeleton, which consists of three different types of protein filaments: microtubules, intermediate filaments, and the actin filament network.

Actin reorganization

Actin is a 42-kDa protein that forms a filament system that spans the cell and is continuously self-assembling and disassembling (reviewed in Holmes et al. 1990; Ono 2007). The "plus" end of each filament has a much higher affinity for monomeric actin than the opposite or "minus" end of the filament (Figure 1).

Figure 1. Dynamic polymerization and depolymerization of actin. Self-assembly of actin filaments occurs at the plus end of an existing actin filament, as monomeric actin is incorporated. Conversely, disassembly occurs at the minus end where actin monomers depolymerize from the filament, causing a continuous rearrangement of the actin filament network.

To date, it has been difficult to estimate how quickly a cell's actin filament network rearranges. Existing visualization methods use markers such as fluorescently labeled actin, anti-actin antibodies, or fluorescently labeled phalloidin (a protein that specifically binds to actin filaments). But these techniques are limited to capturing the state of the actin filament network at the time of staining or labeling, and cannot monitor the dynamics of actin filament rearrangement.

In order to monitor actin rearrangement, microinjection of in vitro-labeled actin has been used to specifically identify actin filaments that are made or rearranged after the time of injection. The actin filament network has been reported to rearrange completely in one hour in PtK2 epithelial cells, based on the incorporation of the injected labeled actin into the complete intracellular actin network (Theriot and Mitchison 1991). Unfortunately, microinjection is very labor-intensive and can only be performed on a limited number of cells at one time. For this reason, we took advantage of Lenti-X lentiviral delivery and ProteoTuner technology to develop a simpler alternative.

Controlling actin stability with the ProteoTuner systems

Our ProteoTuner systems now make it possible to observe fast processes such as the cytoskeletal rearrangement of the actin filament network in a simpler and more reliable fashion. The ProteoTuner systems allow a protein of interest to be expressed as a fusion protein containing an extremely labile destabilization domain (DD). Proteins fused to the DD are quickly degraded by the proteasomes. However, if cells expressing the DD-tagged fusion protein are cultured in media containing the stabilizing ligand Shield1, the protein is protected from degradation and can accumulate inside the cell. The fusion protein rapidly accumulates to detectable levels, in as few as 15–20 minutes after adding Shield1 to the culture media (Banaszynski et al. 2006).

Thus, we took advantage of the fast kinetics of the ProteoTuner systems in order to investigate the dynamic turnover of actin filaments inside cells (Figure 2). Specifically, we used the Lenti-X ProteoTuner system for this experiment in order to deliver and express DD-tagged actin molecules in neural progenitor cells.

Figure 2. Experimental design. In the absence of Shield1, only mCherry-Actin is present to be incorporated into newly forming actin filaments. When Shield1 is added, DDAcGFP1-Actin is stabilized and is therefore incorporated together with mCherry-Actin into newly formed actin filaments.

We designed two lentiviral constructs: One encoded human a-actin fused at its N-terminus to mCherry, which constitutively labeled all actin filaments in the transduced cells. The other encoded human a-actin fused at its N-terminus to DD-tagged AcGFP1, to create DD-AcGFP1-Actin. This DD-AcGPF1-Actin fusion was only stable in the presence of Shield1, which protected it from degradation. In the absence of Shield1, DD-AcGFP1-Actin was immediately targeted to the proteasomes, without being incorporated into the cellular actin filament network. HeLa cells and neural progenitor cells were each coinfected with the mCherryActin and DD-AcGFP1-Actin lentiviral constructs and cultured either with or without Shield1. The addition of Shield1 caused very quick stabilization of DD-AcGFP1-Actin, allowing us to monitor its incorporation into the actin filaments.

Results

As predicted, in the absence of Shield1 we observed very strong red cytoskeletal labeling (from mCherry-Actin) without any obvious signal in the green channel (Figure 3). This suggests that DD-AcGFP1-Actin was degraded rapidly, before integrating into the dynamically changing actin filament network (Figure 3, Panels A–B and E–F). One hour after adding Shield1 to the HeLa cells, DD-AcGFP1 was stabilized and integrated into the actin filament network together with mCherry-Actin (Figure 3, Panels C–D). In fact, after only 15 minutes of Shield1 treatment, DD-AcGFP1-Actin could be observed colocalized with mCherryActin in the actin filaments (data not shown). Experiments with infected neural progenitor cells (Figure 3, Panels E–H) allowed us to conclude that actin filament turnover occurs in a rapid fashion (three hours or less) in these cells as well.

Figure 3. Controlling actin stability with the ProteoTuner system. In the absence of Shield1, DD-AcGFP1-Actin is not present (Panels B & F) despite a normal, mCherry-labeled actin filament network (Panels A & E). In the presence of Shield1, DD-AcGFP-Actin is stabilized, and present in the actin network (Panels D & H) in addition to mCherry-labeled actin (Panels C & G). Cells were fixed using 4% paraformaldehyde and imaged with a 40X objective on a Zeiss Axioskop^M microscope using the green Chroma filter set HQ460/40, Q490LP, HQ515/30 and the red Chroma filter set HQ 540/40X, 570DCPL, D600/50M.

This experiment illustrated that the actin filament network was completely remodeled in less than an hour in HeLa cells, using both mCherry-Actin and Shield1-stabilized DD-AcGFP1-Actin. These results are in agreement with the rapid dynamics reported in the complicated microinjection experiment described above (Theriot and Mitchison 1991) but were obtained using the far simpler ProteoTuner-based method.

In addition to observing the fast restructuring of the actin network, this experiment demonstrated that the DD did not interfere with the proper incorporation of the fusion protein monomers into actin filaments. The same filament pattern was observed with DD-AcGFP1-Actin as with mCherryActin (Figure 3, Panels C–D and G–H). Furthermore, no toxicity was observed in the presence of Shield1.

ProteoTuner systems make it possible to change the amount of a protein of interest rapidly, allowing us to observe highly dynamic processes such as cytoskeletal rearrangement by simply expressing the protein of interest as a DD-fusion and controlling its stability by adding or removing Shield1.

References

Banaszynski, L. A., Chen, L.-C., Maynard-Smith, L. A., Ooi, A. G. L. & Wandless, T. J. A rapid, reversible, and tunable method to regulate protein function in living cells using synthetic small molecules. Cell 126, 995–1004 (2006). 

Holmes, K. C., Popp, D., Gebhard, W. & Kabsch, W. Atomic model of the actin filament. Nature 347, 44–9 (1990).

Ono, S. Mechanism of depolymerization and severing of actin filaments and its significance in cytoskeletal dynamics. Int. Rev. Cytol. 258, 1–82 (2007).

Theriot, J. A. & Mitchison, T. J. Actin microfilament dynamics in locomoting cells. Nature 352, 126-31 (1991). 

Destabilized fluorescent protein vectors

In contrast to standard Living Colors fluorescent proteins, which are extremely stable, the destabilized fluorescent protein vectors (pFP-DR) display rapid turnover rates. This shorter half-life makes destabilized pFP-DR variants ideal for:

• Studies that require rapid reporter turnover
• Quantitative reporter assays and kinetic studies
• Accurately measuring the kinetics of transient mRNA transcription from regulated promoters
• Monitoring gene expression during development
• Characterizing cis-regulatory elements
• Analyzing the activity of a promoter or promoter/enhancer element inserted into the MCS located upstream of the reporter gene
• Developing stably transformed cell lines (because destabilized proteins can be expressed without excess buildup of the fluorescent protein)

The Destabilized Fluorescent Protein Vectors are promoterless vectors that encode rapidly degraded forms of ZsGreen1, DsRed-Express, and HcRed1. They include a portion of the mouse ornithine decarboxylase (MODC) gene, fused to the downstream end of the gene for the fluorescent protein. This region of MODC contains a PEST domain that targets the pFP-DR fusion protein for degradation, giving it a substantially shorter half-life (Li et al. 1998). These destabilized variants can be used in cells from any organism that employs PEST sequence-mediated degradation pathways.

As with other fluorescent proteins, the pFP-DR vectors are human codon-optimized and retain the same spectral properties as our standard fluorescent proteins. Destabilized fluorescent proteins can be easily and immediately visualized by fluorescence microscopy or analyzed by flow cytometry.

References

Li, X. et al. Generation of destabilized green fluorescent protein as a transcription reporter. J. Biol. Chem. 273, 34970–5 (1998).

Bright colors that can be multiplexed (i. e., have very different excitation and emission maxima) are ideal for cell labeling and imaging. We offer the widest spectral range—so you can choose based on your color, filter, or multiplexing needs.

• DsRed-Express2
• mCherry
• tdTomato
• mOrange*
• ZsGreen1
• AmCyan1

*mOrange has been discontinued. mOrange2 is a variant of mOrange which has been modified for improved stability.

Red fluorescent proteins are ideal for in vivo imaging due to reduced autofluorescence. Like all fluorescent proteins, they can be detected in cells without adding cofactors or substrates, making them valuable, noninvasive tools for investigating biological events in living cells (Matz et al. 1999).

mCherry

mCherry is one of the Fruit fluorescent proteins, which were developed in Dr. Roger Tsien's lab by directed mutagenesis of mRFP1, a monomeric mutant of DsRed (Campbell et al. 2002; Shaner et al. 2004; Wang et al. 2004; Shu et al. 2006).

mCherry fusion constructs

mCherry has been successfully fused to several proteins, including actin and tubulin (Figure 1). mCherry fusions have been reported in Arabidopsis (Song et al. 2007), zebrafish (Pisharath et al. 2007), E. coli (Pradel et al. 2007), HIV virions (Campbell et al. 2007), and yeast (Snaith, Samejima, and Sawin 2005). These fusions have also been used for quantitative imaging techniques including fluorescence resonance energy transfer (FRET; Anderson et al. 2006), fluorescence recovery after photobleaching (FRAP; Picard, Suslova, and Briand 2006), and fluorescence lifetime imaging microscopy (FLIM; Tramier et al. 2006).

Figure 1. mCherry fusion constructs. HeLa cells were transiently transfected via a lipid-based method, with mammalian expression vectors encoding mCherry fused to either human cytoplasmic actin (Panel A) or tubulin (Panel B). Cells were fixed using 4% paraformaldehyde and imaged 36 hr posttransfection with a 40X objective on a Zeiss Axioskop microscope using the 575/50, 610, 640/50 filter set.

Stable cell lines

We have established three stably transfected HEK 293 cell lines with different levels of mCherry expression (as measured by flow cytometry). Transfected cells were observed to grow at a rate similar to nontransfected control cells, without increased cell death, as determined by visual inspection.

tdTomato

tdTomato is another Fruit fluorescent protein (Campbell et al. 2002; Shaner et al. 2004; Wang et al. 2004; Shu et al. 2006). It is a genetic fusion of two copies of the dTomato gene (Shu et al. 2006) which was specifically designed for low aggregation. Its tandem dimer structure plays an important role in the exceptional brightness of tdTomato. Because tdTomato forms an intramolecular dimer, it behaves like a monomer.

Outstanding in vivo imaging with tdTomato

tdTomato's emission wavelength (581 nm) and brightness make it ideal for live animal imaging studies. In one xenograft mouse model of metastatic breast cancer, tdTomato was easily detected as deep as 1 cm below the surface, and extremely small lesions could be detected and tracked over time (Figure 2; Winnard, Kluth, and Raman 2006). A second model used tdTomato to quantify breast cancer tumor growth in response to target gene activation (Johnstone et al. 2008). Transgenic mouse models have also been developed, including one where tdTomato was used as a reporter for Cre recombination. This model was also useful as a tool for cell-lineage tracing, transplantation studies, and analysis of cell morphology in vivo (Muzumdar et al. 2007). tdTomato has also been used very effectively in fusion protein applications (Bjørkøy et al. 2005) and as a promoter reporter (Alandete-Saez, Ron, and McCormick 2008).

Figure 2. tdTomato, but not GFP, can be detected in the SCID mouse cadaver phantom model. False-color overlay images (regions of interest encircled) indicate that the imaging system could detect tdTomato fluorescence in the cadaver model, but not GFP fluorescence. Panel A. Implanted tube with 100 x 10⁶ MDA-MB-231-tdTomato-expressing cells, imaged with the DsRed filter set. Exposure time: 1 sec. Panel B. Implanted tube with 100 x 10⁶ MDA-MB-231-GFP-expressing cells, imaged with the GFP filter set. Exposure time: 1 sec.

Green fluorescent proteins in plant research

Plants and plant tissues can be successfully imaged with green fluorescent proteins. Although many plants contain endogenous compounds such as phenolics that fluoresce in the green to yellow range with low excitation wavelengths (Stewart 2006; Lin, Irani, and Grotewold 2003), the use of proper controls eliminates the problems associated with this autofluorescence.

Figure 1. Expression of ZsGreen in soybean under white light (Panel A) and fluorescent light (Panel B). Bar = 1 cm.

Figure 2. AcGFP1 expression in maize tissues and plants. Panels A and D. Maize callus under white light and fluorescent light. Panels B and E. Transformed maize leaf and root under white light and fluorescent light. Panels C and F. Kernels from a segregating transformed line under white light and fluorescent light. Imaging used a Zeiss Sv-11 dissecting scope, HB100 light source, and a GFP (plant) cube; exciter HQ470/40, dichroic 495LP, emission D525/50 BP.

ZsGreen labeling of breast cancer cells to visualize metastasis

The brightness and enhanced ability of ZsGreen1 to withstand fixation makes it an ideal fluorescent protein for in vivo metastasis studies. ZsGreen1 was used to label human estrogen receptor-positive breast cancer cells, and fluorescent whole-body imaging on living, unanesthetized, immunocompromised nude mice identified lymph node metastases that arose from estrogen-dependent tumors grown orthotopically in mammary glands. Metastatic cancer cells injected directly into the circulation were detected in bone, brain, adrenal gland, and lung by fluorescence imaging (Figures 3–4; Harrell et al. 2006).

Figure 3. Fluorescent whole-body image of a mouse with bilateral solid tumors derived from estrogen receptor-positive MCF7 human breast cancer cells. Note the presence of ZsGreen1-positive cells in tumors, lymphatic vessels (LV), and inguinal and axillary lymph nodes (LN).

Figure 4. Metastatic lung tumor expressing ZsGreen1. Mouse lung bearing an MCF7+ ZsGreen tumor was fixed, embedded, and sectioned. Sections were stained with hematoxylin and eosin (left; H & E) or deparaffinized, coverslipped, and imaged via fluorescence microscopy (right) [480 nm excitation; 505 nm emission; 20X objective]. ZsGreen1 fluorescence was maintained even after histological processing.

DsRed-Express & DsRed-Express2

DsRed-Express is a rapidly maturing variant of Discosoma sp. red fluorescent protein (DsRed) with mutations that enhance its solubility, reduce its green emission, and accelerate its maturation. Although DsRed-Express most likely forms the same tetrameric structure as wild-type DsRed, DsRed-Express displays a reduced tendency to aggregate (Bevis and Glick 2002). DsRed-Express2 has even higher solubility and was designed to be better suited for cell and stem cell applications (Strack et al. 2008).

Figure 3. Robust expression of DsRed-Express2 in mouse bone marrow hematopoietic stem and progenitor cells. Mononuclear bone marrow cells were transduced with retroviral vectors encoding DsRed-Express (Panel A), DsRed-Express2 (Panel B), or EGFP (Panel C); and fluorescent cells were sorted 87 hr later. Red and green fluorescence signals were detected using the PE and FITC filter sets, respectively. The lines represent gates defined by analyzing untransduced cells.

DsRed2

DsRed2 is a variant of our original red fluorescent protein (DsRed), modified with six point mutations to improve solubility and decrease the time from transfection to detection. DsRed2 retains the benefits typical of red fluorescent proteins, such as a high signal-to-noise ratio and distinct spectral properties for use in multicolor labeling experiments.

mStrawberry

mStrawberry is another Fruit fluorescent protein (Campbell et al. 2002; Shaner et al. 2004; Wang et al. 2004). Its excitation and emission maxima are 574 nm and 596 nm, respectively.

AsRed2

AsRed2 is a novel fluorescent protein that has been adapted from the corresponding full-length cDNA for higher solubility, brighter emission, and rapid chromophore maturation (8–12 hours). It has also been human codon-optimized to enhance translation efficiency in mammalian cells. It is a very good reporter and is useful in two-color analyses with AmCyan1 or ZsGreen1.

References

Alandete-Saez, M., Ron, M. & McCormick, S. GEX3, expressed in the male gametophyte and in the egg cell of Arabidopsis thaliana, is essential for micropylar pollen tube guidance and plays a role during early embryogenesis. Mol. Plant 1, 586–98 (2008).

Anderson, K. I., Sanderson, J., Gerwig, S. & Peychl, J. A new configuration of the Zeiss LSM 510 for simultaneous optical separation of green and red fluorescent protein pairs. Cytometry. A 69, 920–9 (2006).

Bevis, B. J. & Glick, B. S. Rapidly maturing variants of the Discosoma red fluorescent protein (DsRed). Nat. Biotechnol. 20, 83–7 (2002).

Bjørkøy, G. et al. p62/SQSTM1 forms protein aggregates degraded by autophagy and has a protective effect on huntingtin-induced cell death. J. Cell Biol. 171, 603–14 (2005).      

Campbell, E. M., Perez, O., Melar, M. & Hope, T. J. Labeling HIV-1 virions with two fluorescent proteins allows identification of virions that have productively entered the target cell. Virology 360, 286–93 (2007).           

Campbell, R. E. et al. A monomeric red fluorescent protein. Proc. Natl. Acad. Sci. U. S. A. 99, 7877–82 (2002).        

Harrell, J. C. et al. Estrogen receptor positive breast cancer metastasis: altered hormonal sensitivity and tumor aggressiveness in lymphatic vessels and lymph nodes. Cancer Res. 66, 9308–15 (2006).

Johnstone, C. N. et al. Parvin-beta inhibits breast cancer tumorigenicity and promotes CDK9-mediated peroxisome proliferator-activated receptor gamma 1 phosphorylation. Mol. Cell. Biol. 28, 687–704 (2008).      

Lin, Y., Irani, N. G. & Grotewold, E. Sub-cellular trafficking of phytochemicals explored using auto-fluorescent compounds in maize cells. BMC Plant Biol. 3, 10 (2003).

Matz, M. V et al. Fluorescent proteins from nonbioluminescent Anthozoa species. Nat. Biotechnol. 17, 969–73 (1999). 

Muzumdar, M. D., Tasic, B., Miyamichi, K., Li, L. & Luo, L. A global double-fluorescent Cre reporter mouse. Genesis 45, 593–605 (2007).

Picard, D., Suslova, E. & Briand, P.-A. 2-color photobleaching experiments reveal distinct intracellular dynamics of two components of the Hsp90 complex. Exp. Cell Res. 312, 3949–58 (2006).    

Pisharath, H., Rhee, J. M., Swanson, M. A., Leach, S. D. & Parsons, M. J. Targeted ablation of beta cells in the embryonic zebrafish pancreas using E. coli nitroreductase. Mech. Dev. 124, 218–29 (2007).             

Pradel, N. et al. Polar positional information in Escherichia coli spherical cells. Biochem. Biophys. Res. Commun. 353, 493–500 (2007).

Shaner, N. C. et al. Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein. Nat. Biotechnol. 22, 1567–72 (2004).

Shu, X., Shaner, N. C., Yarbrough, C. A., Tsien, R. Y. & Remington, S. J. Novel chromophores and buried charges control color in mFruits. Biochemistry 45, 9639–9647 (2006).        

Snaith, H. A., Samejima, I. & Sawin, K. E. Multistep and multimode cortical anchoring of tea1p at cell tips in fission yeast. EMBO J. 24, 3690–9 (2005).             

Song, L., Han, M.-H., Lesicka, J. & Fedoroff, N. Arabidopsis primary microRNA processing proteins HYL1 and DCL1 define a nuclear body distinct from the Cajal body. Proc. Natl. Acad. Sci. U. S. A. 104, 5437–42 (2007).

Stewart, C. N. Go with the glow: fluorescent proteins to light transgenic organisms. Trends Biotechnol. 24, 155–62 (2006).

Strack, R. L. et al. A noncytotoxic DsRed variant for whole-cell labeling. Nat. Methods 5, 955–7 (2008).     

Tramier, M., Zahid, M., Mevel, J.-C., Masse, M.-J. & Coppey-Moisan, M. Sensitivity of CFP/YFP and GFP/mCherry pairs to donor photobleaching on FRET determination by fluorescence lifetime imaging microscopy in living cells. Microsc. Res. Tech. 69, 933–9 (2006).       

Wang, L., Jackson, W. C., Steinbach, P. a & Tsien, R. Y. Evolution of new nonantibody proteins via iterative somatic hypermutation. Proc. Natl. Acad. Sci. U. S. A. 101, 16745–16749 (2004).   

Winnard, P. T., Kluth, J. B. & Raman, V. Noninvasive optical tracking of red fluorescent protein-expressing cancer cells in a model of metastatic breast cancer. Neoplasia 8, 796–806 (2006).

Bright fluorescent proteins make excellent reporters. Monitor the activation of different promoters or promoter/enhancer combinations using traditional promoterless reporter constructs or next-generation on-demand reporters, which use ProteoTuner technology to provide extremely low background and bright signals.

On-Demand (DD) ZsGreen1 reporter system

No more tradeoffs—perform reporter studies with both a low background and a broad dynamic range. The DD-ZsGreen1 Reporter System combines a bright fluorescent protein reporter (ZsGreen1) for high signal intensity, with ligand-dependent ProteoTuner protein stabilization/destabilization technology to eliminate background. ZsGreen1 is expressed as a fusion with a ligand-dependent destabilization domain (DD). The DD rapidly targets the reporter protein for proteasomal degradation, guaranteeing a low reporter-background signal at the start of your experiment. However, when the small, membrane-permeant ligand Shield1 is added to the sample, it binds to the DD and protects the reporter from degradation so that it can accumulate. The DD-ZsGreen1 Reporter System is available in plasmid and lentiviral formats.

Viral vectors allow you to deliver and express your gene of interest and a fluorescent protein in a wide variety of cell types. Depending on which vector you choose, the fluorescent protein and protein of interest can be fused together for subcellular localization or tag studies, or simultaneously coexpressed from the same transcript but as separate proteins (from IRES vectors).

• Use adenoviral vectors to transduce dividing and non-dividing cells. Obtain very high titers, high expression, and a broad host range. Cloning is even simpler than standard plasmid cloning.
• Use lentiviral vectors to transduce a wide range of hard-to-transfect cell types, including dividing and nondividing cells, stem cells, terminally differentiated cells, and neuronal cells. Optimized for high expression and high titers.
• Use retroviral vectors to transduce hard-to-transfect dividing cells. Easily create stable cell lines. Read about our bicistronic and fusion retroviral vectors.

Choose from our lentiviral, retroviral or adenoviral vectors.

Fluorescent timer vectors

Study promoter activity using the Living Colors Fluorescent Timer, a fluorescent protein that shifts color from green to red over time (Terskikh et al. 2000). This color change provides a way to visualize the timeframe of promoter activity, indicating where in an organism the promoter is active and also when it becomes inactive. Easily detect the red and green emissions indicating promoter activity with fluorescence microscopy or flow cytometry.

Easily characterize promoter activity

The Fluorescent Timer protein is a mutant form of the DsRed fluorescent reporter, containing two amino acid substitutions which increase its fluorescence intensity and endow it with a distinct spectral property: as the Fluorescent Timer matures, it changes color—in a matter of hours, depending on the expression system used. Shortly after its synthesis, the Fluorescent Timer begins emitting green fluorescence but as time passes, the fluorophore undergoes additional changes that shift its fluorescence to longer wavelengths. When fully matured, the protein is bright red. The protein's color shift can be used to follow the on and off phases of gene expression (e.g., during embryogenesis and cell differentiation).

Fluorescent Timer under the control of the heat shock promoter hsp16-41 in a transgenic C. elegans embryo. The embryo was heat-shocked in a 33°C water bath. Promoter activity was studied during the heat-shock recovery period. Green fluorescence was observed in the embryo as early as two hours into the recovery period. By 50 hr after heat shock, promoter activity had ceased, as indicated by the lack of green color.

Timer vectors

pTimer (left) is primarily intended to serve as a convenient source of the Fluorescent Timer cDNA. Use pTimer-1 (right) to monitor transcription from different promoters and promoter/ enhancer combinations inserted into the MCS located upstream of the Fluorescent Timer coding sequence. Without the addition of a functional promoter, this vector will not express the Fluorescent Timer.

Detecting Timer fluorescent protein

You can detect the Fluorescent Timer with the Living Colors DsRed Polyclonal Antibody.

References

Terskikh, A. et al. "Fluorescent timer": protein that changes color with time. Science 290, 1585–88 (2000).

GFPuv (a variant of Aequorea victoria green fluorescent protein) is optimized for maximal fluorescence when excited by UV light [360–400 nm]. GFPuv expression is driven by the lac promoter.

We also offer a variety of bacterial expression vectors for other fluorescent proteins (also driven by P_lac), see table below.

Fucci cell cycle reporter vectors deliver fluorescent, ubiquitination-based, cell-cycle indicators that allow you to identify cells in various phases of the cell cycle. Fucci vectors contain Cdt1 or Geminin proteins, whose levels fluctuate differentially throughout the cell cycle, plus a red or cyan fluorescent tag.

Movie of cell cycle progression visualized with Fucci probes. No color: cells transitioning between phases M and G1. Red: cells in G1 phase. Light pink/violet: cells transitioning between phases G1 and S. Cyan: the nuclei of cells in phases S through M.

Our Proteasome Sensor Vector is ideal for image-based assays for compounds with proteasome-inhibiting or activating properties.

Fluorescently labeled stem cells can be used to analyze features and behaviors, and to monitor events including interactions with adjacent cells with precise spatial and temporal resolution. EF-1 alpha promoter vectors are useful in embryonic stem cells, where the CMV promoter has diminished activity in these cell types.

• AcGFP1
• DsRed-Monomer
• mCherry
• tdTomato
• DsRed-Express2
• E2-Crimson
• EF-1 alpha promoter vectors