The higher molecular weight bands observed by immunoblots Figs 1 and 2 , Supplementary Fig. Comparisons of fluorescence intensity measurements revealed a decrease of fluorescence signal in cells transiently expressing the non-optimized FP variant. This measurable decrease in fluorescence signal was specifically a consequence of the oxidizing environment of the ER. When the two BFPs were localized to the cytoplasm, in the absence of disulphide formation, the BFPs exhibited nearly identical mean fluorescence intensities Fig.
Error bars signify s. Both freely diffusing luminal and membrane-localized non-optimized FPs within the secretory pathway are prone to inappropriate disulphide bond formation.
We localized an optimized and a non-optimized FP to the GC with the 1,4-galactosyltransferase, GalT, signal anchor transmembrane domain Fig. To investigate where the misfolded dark secretory pathway protein fusions localize, we compared where optimized and non-optimized GalT—FP reporters localize in cells.
These data suggest that, in GalT—mGFP, the interchain disulphide bonds that we observed via nonreducing immunoblots Fig. These examples establish the importance of using the inert oxFPs for studies of secretory compartments in cells. MCherry does not contain cysteine residues, is reportedly monomeric 13 , and it has been successfully expressed in oxidizing environments, including bacterial periplasm Therefore, mCherry would be expected to be a suitable FP option for use in secretory protein fusions.
However, for GC-localized membrane fusion proteins, we have previously found mCherry to be a potentially problematic choice, as mCherry fusions often localize to numerous punctate structures distinct from GFP family members fused to GalT 6. Others have reported that mCherry protein fusions can aggregate 41 , impair growth in yeast and development of tissues in Xenopus laevis 27 , These issues motivated us to test the utility of our oxFPs for dual-color labelling schemes, especially for labelling the GC.
In contrast, GalT—mCherry fusions localized in a dispersed pattern of puncta throughout the cell. However, at lower expression levels, especially in stably transfected cells, a mixed puncta and GC distribution can be observed for GalT—mCherry Supplementary Fig. If so, oligomers or aggregates could disrupt proper localization in the GC. We examined mCherry oligomerization using the live cell assay that we recently developed for assessing FP monomericity in cells.
In addition, we also tested whether other FPs were sufficiently monomeric to avoid FP oligomerization artefacts 21 , Therefore, we subsequently monomerized all of our constructs to create monomeric oxidizing environment-optimized mox FPs.
On the basis of our results, the tendency of GFP family members to dimerize does not appear to impact GC structure. Therefore, we consider oligomerization unlikely to explain the punctate structures observed in GalT—mCherry expressing cells.
We note that the reported monomericity of mRuby2 was assessed for the parent protein, mRuby, by passing a solution of purified protein over a molecular sizing column 25 , Using our new moxFPs, we tested their suitability for use in GalT reporters and for dual-colour imaging pairs.
Thus, the ox- and monomerized ox- FPs can be exploited for at least two different two-colour pairs for multicolour imaging. Of these red FPs, only mCherry is cysteine-free.
Thus, many RFPs appear to be suboptimal for use in the secretory pathway and even mCherry appears to be problematic when used with a GC-localized membrane protein.
However, given the need for red FPs in multicolour imaging, the punctate distribution of GalT—mCherry merited further investigation before rejecting use of mCherry in the secretory pathway.
Importantly, upon closer examination of the GC distribution of the other GalT—moxFP fusions, we encountered a surprising result. As moxCerulean3 Table 2 and Supplementary Fig. RFPs are essential reagents for cell biologists and they have many desirable properties, including brightness and illumination with longer less phototoxic wavelengths that are less prone to exciting autofluorescent substrates in cells.
Several of the red FPs were evolved from inherently dimeric or tetrameric proteins. For fusion proteins, it is critical that the FP of choice not oligomerize. We Supplementary Fig. We observed that mCherry only weakly oligomerizes Supplementary Fig. Against this background, we sought to understand the behaviour of GalT—mCherry.
Both markers robustly co-localize with the compact perinuclear GC structures labelled by the GalT fusions, but neither labels the mCherry puncta throughout the cell Fig. Several red puncta co-localize with the LAMP1 positive structures. Next, we revisited our data with the GalT-moxCer3 reporter, which also localized in a punctate distribution Fig. Aside from spectra, a major difference between Cerulean3 moxCer3 and other GFP family members is the exceptionally low pKa 3.
The pKa is the pH at which a fluorophore will be quenched in brightness by one half. FPs often fully quench at only slightly lower pH values. In addition, mCherry is resistant to proteolysis in lysosomes, while GFP family members are highly susceptible Katayama et al. We considered the possibility that a similar phenomenon occurs for mCherry fused to membrane proteins in the secretory pathway. The pH of late GC compartments and the endosomal—lysosomal system decreases from 6.
To test whether GalT—moxGFP traffics similarly could accumulate in puncta, the pH gradient of the secretory pathway was disrupted.
Together, these observations have important implications for imaging studies in the secretory pathway. The choice of FP clearly influences the localization pattern. Our data suggest that GalT eventually traffics to the lysosome; though we cannot currently rule out the possibility that mCherry contains a cryptic lysosome-targeting sequence.
In addition, mCherry reportedly matures more slowly than the GFP family member 51 and this property could decrease the visible pool of GalT—mCherry in the GC at a steady state. Huang et al. Whether the properties of a GFP family member or mCherry are advantageous or disadvantageous will depend on the goal of the experiment, though awareness of these issues will enable investigators to exploit these different properties.
Therefore, when using FP fusions in experiments, it is vital to validate the behaviour of the fusion protein relative to the native protein.
Finally, to confirm the utility of the moxFPs, we sought to confirm the functionality of the FPs in the cytoplasm. We found that all of the GFP family member moxFPs functioned comparably with the parental proteins when fused into standard actin and tubulin constructs and gave expected distributions of actin stress fibres and microtubules Supplementary Fig.
We had different results with mNeonGreen and moxNeonGreen. Both versions formed actin stress fibres Supplementary Fig. However, neither version was significantly incorporated into microtubules Supplementary Fig. In the original report 38 , it was unclear how long the linker was, but we tried both lengths and had the same result Supplementary Fig.
We recommend caution when making fusions with mNeonGreen and its variants. Despite the resulting undesirable disulphide -bonded misfolded oligomers, the higher molecular weight oligomers represent a curious protein-folding phenomenon that suggests potentially important insights into the process of FP folding in cells.
As nascent FPs enter the crowded ER lumen, they will transiently contact numerous diffusing secretory proteins. We have no evidence that FPs form inappropriate disulphide bonds arbitrarily with random luminal ER proteins. We would have expected to see smears on nonreducing immunoblots, instead of discrete bands at specific molecular weights that correspond to FP interchain disulphide-bonded FP species.
Instead, FPs likely form persistent at least for seconds and maybe longer one or more intermediate species FP int that can oligomerize via an interface distinct from the hydrophobic dimerizing interface on the outer surface of GFP. These species are probably also recognized by resident ER chaperones. It is unclear whether the quasi-species are reversible and can eventually form two separately folded and fluorescent FPs in the cytoplasm or ER.
Within the ER, cysteine residues in the FP int are exposed and could preferentially form a disulphide bond with an interacting FP int species. This scenario is especially plausible if the FP int has a high affinity for homo-oligomerization, increasing the likelihood of disulphide bonds forming between FPs, probably in cooperation with a PDI family member. To circumvent the challenges that subcellular organelles pose to FPs, we sought to optimize FP technologies. We have created a set of bright oxFPs Table 2 that can be used for quantitative multicolour labelling strategies.
Furthermore, the moxFPs versions are monomeric and encode no apparent sequence-specific sites accessible for N-linked glycosylation or disulphide bond formation.
The moxFPs are currently the most practical solution to reliably engineer FP fusions with soluble and membrane cellular proteins of interest. Our results emphasize the urgent need to develop better cysteine-less and robustly monomeric red FPs for use in the secretory pathway.
We have extensively studied the impact of the chemically distinct and reactive environments of the eukaryotic secretory compartments on the functionality of FPs. As we explored popular FP applications, it became apparent that moxFP modifications are essential.
Findings with GalT—mGFP further confirm significant loss and mislocalization of functional fluorescent molecules due to the formation of non-native disulphide bonds Fig.
We predict that the accumulation of misfolded non-fluorescent FP molecules within the ER could lead to underappreciated off-target effects, including changes to the ER environment through titration of ER chaperones and crowding effects of aggregated misfolded proteins leading to unintended side effects on secretory resident proteins and protein trafficking.
Equally importantly, the non-FP component of a dark pool of fusion proteins may remain functional and could lead to gross underestimations of the location and activity levels of fusion proteins. In such cases, the concentration of oxFP fusions expressed in cells will be most accurately defined by biochemical analysis instead of imaging analysis.
For example, the observed FP-fusion fluorescence would correlate with the levels of the correctly folded monomeric species detected by immunoblot.
As the optimized oxFPs do not accumulate misfolded, non-fluorescent species, total levels of fluorescence will quantitatively reflect FP-fusion levels.
This last point is especially pertinent to the current thrust towards quantitative single-cell imaging for modern cell biologists. CMV promoter-driven transiently transfected plasmids have been useful for studying proteins in cells, but it remains unclear how much the behaviour of grossly overexpressed proteins accurately reflects physiologic cell and protein behaviours.
New engineering technologies, especially CRISPR, have made it possible to chromosomally tag endogenous genes with FPs to enable truly physiologic expression and regulation of levels of FP-fusion proteins In the absence of CMV promoter-driven expression levels, it will be vital for each FP fusion to correctly fold and fluoresce to quantitatively visualize FP-fusions using fluorescence microscopy.
Quantitative experiments to investigate cell-to-cell variation, proteomics and to help develop quantitative cellular models will require accurate censuses of cellular protein populations. FP fusions absolutely must not distort or perturb cellular compartments. The bright multicolour moxFPs developed here were motivated by these needs and represent essential tools for cell biologists studying diverse intracellular environments.
Optimized oxCerulean, oxCerulean3, moxNeonGreen and oxVenus were synthesized by GenScript Piscataway, NJ and included cysteine to serine mutations, superfolder mutations, and were human codon optimized. To create cytoplasmic and ER-localized reporters, FP-encoding sequences were amplified using primers listed in Supplementary Table 1. To investigate specific amino-acid substitution consequences on FP fluorescence, reverting mutations were re-introduced into FPs using designated primers Supplementary Table 1.
All the constructs were confirmed by sequencing. FP sequences were amplified using primers listed in Supplementary Table 1. Live cell images were acquired using an Axiovert wide-field fluorescence microscope Carl Zeiss Microimaging Inc. Images were acquired with QCapture software.
Cells were fixed with freshly diluted 3. Images were acquired using identical imaging parameters. At least five fields of view were captured for each condition on three separate days.
Images were analysed with ImageJ software. To measure a cell MFI, first the cell nucleus was excluded and detectable fluorescence signal was selected using ImageJ threshold. Images of the anti-Alexa Fluor distributions were captured using identical imaging conditions and then analysed using ImageJ. Purified protein samples were measured in PBS at room temperature. To determine extinction coefficients and quantum yields, absorbance and fluorescence of the new fluorescent variants were compared with that of parental proteins.
To assess photobleaching sensitivity under comparable imaging parameters, oxFPs and parental FPs were transiently expressed in U-2 OS cells. Cells were imaged using standard live cell imaging conditions. Images were acquired at five frames per second for frames. How to cite this article: Costantini, L. A palette of fluorescent proteins optimized for diverse cellular environments. Prasher, D. Primary structure of the Aequorea victoria green-fluorescent protein.
Gene , — Rizzo, M. Fluorescent protein tracking and detection: fluorescent protein structure and color variants. Cold Spring Harb. PubMed Google Scholar. Shaner, N. Advances in fluorescent protein technology. Cell Sci. Miyawaki, A. Red fluorescent proteins: chromophore formation and cellular applications.
Shcherbakova, D. Red fluorescent proteins: advanced imaging applications and future design. CAS Google Scholar. Costantini, L. Fluorescent proteins in cellular organelles: serious pitfalls and some solutions. DNA Cell Biol. In many microscopes the filters are not narrow enough to distinguish between closely related colors.
Furthermore, most FPs have a broad range of emission which will be detected by longer-wavelength filters e. GFP also emits yellow light. CFP excites the fluorescence of another FP e. Indeed, FRET is often used to determine if two proteins interact.
Oligomerization: The first generations of FPs were prone to oligomerize. This may affect the biological function of the FP-fusion protein. Therefore, these FPs cannot be used in oxygen deprived environment. Recently, a new GFP isolated from the Unagi eel was shown to mature independently of oxygen, making suitble for use in anaerobic conditions.
Maturation Time: Maturation time is the time it takes the FP to correctly fold and create the chromophore. This can be from a few minutes after it is translated to a few hours. Temperature: FPs maturation times and fluorescent intensity can be affected by the temperature.
Brightness: Brightness is a measure of how bright is the emission. Brightness is calculated as the product of extinction coefficient and quantum yield of the protein, divided by In many cases the brightness is compared to that of EGFP which is set as 1.
Some proteins are very dim e. TagRFP, which has a brightness of 0. Photostability can be affected by experimental parameters e. Photostability: Fluorescent molecules gets bleached i.
However, for most FPs it is a few seconds to a few minutes. Thank You to Our Guest Blogger! Chudakov et. In a footnote to his paper describing the isolation of the bioluminescent protein aequorin from Aequorea , Shimomura wrote, "A protein giving solutions that look slightly greenish in sunlight though only yellowish under tungsten lights, and exhibiting a very bright, greenish fluorescence in the ultraviolet of a Mineralite [a handheld ultraviolet lamp], has also been isolated from squeezates" Shimomura et al.
Soon after, Shimomura reported the fluorescence emission spectrum of this protein and suggested that energy transfer from aequorin to this green fluorescent protein could explain why the in vivo luminescence of Aequorea is greenish and not blue like the luminescence of purified aequorin Johnson et al.
In Morin and Hastings described very similar green fluorescent proteins isolated from Obelia phylum Cnidaria, class Hydrozoa and the sea pansy Renilla reniformis phylum Cnidaria, class Anthozoa. The nature of the chromophore itself remained a mystery until when Shimomura correctly determined the chromophore to be a 4- p -hydroxybenzylidene imidazolidinone moiety covalently linked within the polypeptide chain Shimomura Achieving this breakthrough necessitated not only the laborious harvest of mg of the naturally occurring green fluorescent protein from Aequorea , but also remarkable chemical intuition on the part of Shimomura.
The complete primary sequence of the amino acids of Aequorea green fluorescent protein accession P was not revealed until the cloning and sequencing of its cDNA by Prasher in Prasher et al. Just two years later came the first dramatic demonstrations that the gene was self-sufficient to undergo the post-translational modifications necessary for chromophore formation. Specifically, Chalfie reported the gene encoding Aequorea green fluorescent protein could be functionally expressed in the sensory neurons of the worm Caenorhabditis elegans Chalfie et al.
The biological research community, armed with the arsenal of powerful molecular biology techniques developed during the preceding two decades, was quick to recognize the unique utility of a genetically-encoded fluorophore as a marker of gene expression and protein localization.
Accordingly, by November there were at least 36 additional examples of applications of recombinant Aequorea green fluorescent protein Cubitt et al. In hindsight, the cloning of the gene and the first demonstrations of recombinant expression in non-jellyfish organisms marks a clearly discernible turning point in the history of fluorescent protein research.
Shimomura's assignment of the chromophore structure was unambiguously confirmed by a detailed reinvestigation of a chromophore-containing peptide fragment in Cody et al. The exact order and mechanism of these steps is a matter of ongoing investigation Zhang et al. An early, and still generally accepted, proposed mechanism is shown in Figure 3B Heim et al.
In this mechanism, chromophore formation starts with the nucleophilic glycine 67 amide nitrogen attacking the electrophilic serine 65 carbonyl carbon to form a 5-membered ring in the main chain of the protein. The resulting tetrahedral hemiaminal intermediate undergoes an elimination of water to form a second intermediate. The installation of this double bond simultaneously converts the 5-membered ring into an aromatic system and puts it into conjugation with the aromatic phenol ring of the tyrosine side chain.
Excitation of the neutral phenol species results in the fast tens of picoseconds excited state proton transfer ESPT Chattoraj et al.
Variants of Aequorea green fluorescent protein with the ground state equilibrium shifted to either the phenol Tsien ; Zapata-Hommer et al. The growing popularity of Aequorea green fluorescent protein, and the demand for additional variants that fluoresced at wavelengths other than green, prompted researchers to begin the search for homologues in other marine organisms.
This effort came to fruition in late when at team of researchers from the Russian Academy of Science reported that reef Anthozoa contain fluorescent proteins with hues ranging from cyan to red Figure 4 Matz et al. The inspiration behind this breakthrough discovery is credited to the evolutionary biologist Yulii A. Labas who had prompted Mikhail V. Matz, then a graduate student in the lab of Sergey A.
Lukyanov, to attempt to clone Aequorea green fluorescent protein homologues from the brightly colored tentacle tips of a sea anemone and several other Anthozoan organisms. An amusing recountal of this episode can be found here. A red fluorescent protein commonly known as DsRed from Discosoma sp. It is now generally understood that many of the bright and varied colors of reef coral are due to fluorescent proteins and their nonfluorescent homologues Dove et al.
Indeed, 4-years prior to the groundbreaking paper from Lukyanov, the pink pigment of the Anthozoan coral Pocillopora damicornis had been isolated and shown to be a proteinaceous pigment, as opposed to a non-protein pigment, which the authors dubbed pocilloporin Dove et al. The authors did not, at that time, recognize that the protein was indeed a member of the GFP-like superfamily. This revelation was reported sometime after the Lukyanov paper, when the gene encoding pocilloporin was sequenced and shown to encode a protein with It is now apparent that, in terms of extinction coefficient and quantum yield, naturally occurring fluorescent proteins lie on a broad continuum.
Pocilloporin often referred to as a nonfluorescent chromoprotein can be conceptualized as a fluorescent protein that has a quantum yield of effectively zero and thus lies at one extreme of the continuum.
In recent years fluorescent proteins have been identified in several non-Cnidarian animals. Specifically, fluorescent proteins have been identified in 3 species of lancelet phylum Chordata, subphylum Cephalochordata Deheyn et al.
Although a peer-reviewed report has not appeared, Axxam Milan, Italy has filed a patent on a fluorescent protein gene derived from a comb jelly, possibly adding the phylum Ctenophora to this list WIPO Pub. The archetypical Aequorea green fluorescent protein chromophore is remarkably tolerant of chemical modifications that change its spectral properties. The limits of this tolerance have been explored by both natural protein evolution undoubtedly and unnatural laboratory engineering see below.
In terms of the range of natural variations of fluorescent protein chromophores, researchers have now discovered at least 5 chromophores with chemically distinct conjugated systems Figure 7. Each of these chemical structures is associated with a range of fluorescence hues, following the general trend that more extended conjugation produces longer wavelength fluorescence. For example, the structures shown in A , C , and B of Figure 7 , are associated with greenish, yellowish, and reddish hues, respectively.
The exact excitation and emission wavelengths for a particular chromophore structure also depends strongly on the protein microenvironment that surrounds the chromophore. For example, naturally occurring cyan fluorescent proteins from Anemonia majano Henderson and Remington , Discosoma striata Malo et al. Another type of variation in chromophore structure is the stereochemistry about the exocyclic double bond of the imidazolidinone moiety Figure 7F. The two possible stereoisomers are often referred to as cis and trans , but the Z and E designation is the more appropriate nomenclature.
For example, the Discosoma red fluorescent protein chromophore is the Z stereoisomer shown in Figure 7B , while a nonfluorescent chromoprotein from Montipora efflorescens Prescott et al. Photoinduced isomerizations between Z and E isomers in the same protein have also been shown to occur in some cases Henderson et al.
A photoinduced isomerization of Z and E stereoisomers of the chromophore shown in Figure 7D is also the likely mechanism by which the Anemonia sulcata "kindling" nonfluorescent chromoprotein converts into a red fluorescent protein upon illumination Chudakov et al. In addition to inducing interconversion of stereoisomers, illumination can also result in photochemical reactions that change the covalent structure of the chromophore. For example Trachyphyllia geoffroyi "Kaede" red fluorescent protein Ando et al.
Proteins that undergo effectively identical reactions have also been cloned from Lobophyllia hemprichii Wiedenmann et al.
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