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What is the difference between GFP and rhodopsin?

What is the difference between GFP and rhodopsin?



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GFP and Rhodopsin are both fluorescent proteins. What is the main difference between GFP and Rhodopsin in terms of how they work? I know that their excitation and emission wavelengths are different, but in terms of the biological mechanism, how are they different?


Transgenic expression of a GFP-rhodopsin COOH-terminal fusion protein in zebrafish rod photoreceptors

To facilitate the identification and characterization of mutations affecting the retina and photoreceptors in the zebrafish, a transgene expressing green fluorescent protein (GFP) fused to the C-terminal 44 amino acids of Xenopus rhodopsin (Tam et al., 2000) under the control of the 1.3-kb proximal Xenopus opsin promoter was inserted into the zebrafish genome. GFP expression was easily observed in a ventral patch of retinal cells at 4 days postfertilization (dpf). Between 45-50% of the progeny from the F1, F2, and F3 generations expressed the transgene, consistent with a single integration event following microinjection. Immunohistochemical analysis demonstrated that GFP is expressed exclusively in rod photoreceptors and not in the UV, blue, or red/green double cones. Furthermore, GFP is localized to the rod outer segments with little to no fluorescence in the rod inner segments, rod cell bodies, or rod synapse regions, indicating proper targeting and transport of the GFP fusion protein. Application of exogenous retinoic acid (RA) increased the number of GFP-expressing cells throughout the retina, and possibly the level of expressed rhodopsin. When bred to a zebrafish rod degeneration mutant, fewer GFP-expressing rods were seen in living mutants as compared to wild-type siblings. This transgenic line will facilitate the search for recessive and dominant mutations affecting rod photoreceptor development and survival as well as proper rhodopsin expression, targeting, and transport.


New GFP techniques

For a long time, neuroscientists were unable to activate/stimulate single neurons given that they could only stimulate the brain cells with electrodes. However, through optogenetics, it is now possible to stimulate individual neurons instantly. This is achieved by using an algae protein (attached to the neuron of interest) as well as light.

Here, the fluorescent protein is used to indicate which of the neurons has been manipulated to become the on and off switch.


2 Materials and methods

2.1 Materials

The design and chemical synthesis of the gene for rhodopsin used in these studies has been published [9] . pEGFP-C2 and pEGFP-N1, sources of the gene for EGFP, were from Clontech. n-Dodecyl-β- D -maltoside (DDM) was from Calbiochem. The anti-rhodopsin monoclonal antibody 1D4, which specifically recognizes the C-terminal eight amino acids of rhodopsin (Glu-Thr-Ser-Gln-Val-Ala-Pro-Ala) [10, 11] , was purified from hybridoma growth medium and coupled to Sepharose 4B [12] for use in immunoaffinity purification of the fusion proteins, as previously described for rhodopsin [13] . Peptide I, with sequence Asp-Glu-Ala-Ser-Thr-Thr-Val-Ser-Lys-Thr-Glu-Thr-Ser-Gln-Val-Ala-Pro-Ala corresponding to the C-terminal 18 amino acids of rhodopsin, was from American Peptide. 11-cis-Retinal was synthesized according to published procedures [14] .

2.2 Genes for fusion proteins rho/EGFP and rho/EGFPi

We prepared genes for two fusion proteins in this work using standard methods. Both genes were cloned between the EcoRI and NotI cloning sites of the mammalian cell expression vector pMT3 [15] . Rho/EGFP (Fig. 1A ) is composed of N-terminal rhodopsin followed by the entire coding sequence of EGFP followed by the C-terminal eight codons of rhodopsin (epitope for the 1D4 antibody). Thus, there are two copies of the C-terminal eight amino acids of rhodopsin in this construct. This was necessary to ensure proper targeting of the protein in rod photoreceptor cells [7] and to ensure that the fusion protein would be recognized by the 1D4 antibody which requires a C-terminal epitope [16] .

Rho/EGFPi (Fig. 1B) is composed of full-length rhodopsin with the coding sequence for EGFP inserted between amino acids Ala333 and Ser334 in the C-terminal tail. Rho/EGFPi was designed to mimic the Moritz et al. construct [1] , and while not identical it does display a similar phenotype with respect to in vitro assays for transducin activation and phosphorylation by RK, as is shown below.

2.3 Expression in COS cells and purification of the proteins

Rho/EGFP and rho/EGFPi were expressed transiently in COS cells transfected using DEAE-dextran as described previously for rhodopsin [13] . Fluorescence microscopy employed an Olympus IX50/IX70 microscope equipped with a U-MWB (exciter filter BP450–480 nm) cube to follow expression of the fusion proteins in intact cells (photographs presented in Fig. 1C,D were taken 40 h post transfection).

Cells were harvested 72 h after initial exposure to DNA and treated with 20 μM 11-cis-retinal in 10 mM sodium phosphate buffer, pH 7.0, containing 150 mM NaCl (PBS) for 1 h to reconstitute the pigments (all procedures beginning with and following initial treatment of the sample with retinal were performed in the dark under illumination from a 15 W incandescent bulb filtered through a Kodak Safelight #2 filter). Cell membranes were solubilized in PBS containing 1% (w/v) DDM and 0.1 mg/ml phenylmethylsulfonyl fluoride, and the post-nuclear supernatant fraction applied to a 1D4-Sepharose 4B matrix for purification of the fusion proteins by immunoaffinity chromatography, as has been described previously for rhodopsin [13] . Bound protein was eluted with 50 μM peptide I in PBS containing 0.1% (w/v) DDM.

2.4 Absorption spectroscopy

UV/visible absorption spectra were obtained on a Hitachi model U-3210 spectrophotometer modified by the manufacturer for dark room use. All spectra were recorded on samples of 1.0 cm path length and analyzed using Kaleidagraph (Version 3.08d). Fusion protein spectra were analyzed by simulation using a linear combination of constituent spectra recorded from purified samples of wild-type rhodopsin and EGFP after normalization on the basis of extinction coefficient (ϵ at 500 nm=42 700 cm −1 M −1 for rhodopsin [17] ϵ at 488 nm=55 000 cm −1 M −1 for EGFP [Living Colors user manual, Clontech]). The concentration of the protein for use in activity assays was then determined from the rhodopsin component. An implicit assumption in this analysis is that the spectral properties (absorption) of rhodopsin and EGFP are unaffected by being brought together in the fusion protein. This assumption was judged to be reasonable on the basis of the close fit of simulated to experimental spectra.

2.5 Miscellaneous procedures for in vitro characterization of the fusion proteins

Western blot analysis [14] was performed using 2.8 pmol protein per lane with 1D4 as the primary antibody and horseradish peroxidase-conjugated anti-mouse IgG (Santa Cruz Biotechnology) as the secondary. Assays for the activation of transducin were performed with 5 nM purified pigment by following the binding of [ 35 S]guanosine 5′-O-(3-thiotriphosphate) as previously described [18] . Assays for phosphorylation by RK were performed as described [19] using RK from a post-nuclear supernatant fraction of transiently transfected COS cells [20] . Reactions were carried out in mixtures containing 125 nM rhodopsin or fusion protein, 6 μl RK extract, 75 mM [1,3-bis[tris(hydroxymethyl)methylamino]propane] buffer, pH 6.7, 10 mM Mg(OAc)2, 5 mM dithiothreitol, 0.01% DDM, and 100 μM [γ- 32 P]ATP (2000 cpm/pmol). Reactions were performed either in the dark or under light from a 300 W tungsten bulb filtered through a 490 nm cut-on filter. Samples were incubated for 30 min at 30°C and then assayed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis and autoradiography.

2.6 Transgenic X. laevis

Transgenic experiments were performed as previously described [21] . Briefly, transgenic X. laevis were generated by the method of Kroll and Amaya using restriction enzyme-mediated integration of DNA into sperm chromosomes prior to transplantation of the sperm nuclei into unfertilized oocytes [3] . Cell-specific expression in the primary rod photoreceptor cells of the Xenopus retina was achieved with a vector (pXOP(−508/+41)Rho/EGFP) that is similar to those previously described [1, 5, 6] in which a fragment (nucleotides −508 to +41) from the X. laevis opsin promoter, XOP(−508/+41) [5, 22] , is used to drive expression of rho/EGFP. The XOP promoter was isolated by using polymerase chain reaction amplification of X. laevis genomic DNA with the primers ATAAACTGCAGCCCCTAGGCCA and CCAAAGGATCCCTAGAAGCCTG for the sense and antisense strands, respectively, followed by digestion with PstI and BamHI to obtain the −508/+41 fragment [4] .

2.7 Histochemistry and confocal microscopy

Histochemistry was performed essentially as previously described [23] . Briefly, transgenic tadpoles (1 month post injection, ∼stage 51) were anesthetized in 0.25% (w/v) tricane solution, fixed with 4% (w/v) paraformaldehyde in 0.1 M sodium phosphate buffer (PB), pH 7.2, at 4°C overnight, incubated first with 5% and then 15% (w/v) sucrose in PB at room temperature, and finally embedded in TBS (Triangle Biomedical Sciences) and frozen on dry ice. Cryosections (16 μm thickness) were prepared with a Jung CM3000 cryostat (Leica) and then melt-mounted onto glass slides. Samples were allowed to thaw in 0.1 M sodium phosphate buffer, pH 7.2, containing 150 mM NaCl (PBS*) at 37°C. The tissue was then incubated in 5% (v/v) goat serum (Sigma) in PBS* containing 0.3% (v/v) Triton X-100 (PBS*-Tx) for 0.5 h. 1D4 antibody (1 μg/ml diluted in PBS*-Tx) was added and incubation continued for 3–4 h at room temperature, the sample washed four times with PBS*-Tx, and incubated with a 1:100 dilution of tetramethylrhodamine isothiocyanate-conjugated goat anti-mouse IgG (Sigma) for 1 h. The samples were covered with 50% (v/v) glycerol in PB and coverslips and then used to obtain images from a Leica TCS SP2 confocal scanning microscope (Leica Microsystems). Fluorescence images were obtained using excitation wavelengths of 488 nm for EGFP and 543 nm for rhodamine. The Nomarski DIC images were obtained from a Zeiss Axioplan-2 optical (Carl Zeiss) microscope equipped with Nomarski optic and GFP filter (FITC LP #41012, Chroma Technology).


Conclusions

The results presented here demonstrate that knock-ins of human rhodopsin-GFP fusion genes produce expression patterns that are consistently identical to the expression patterns of normal mouse rhodopsin. Moreover, expression can be tuned by manipulation of the 5′ untranslated region of the gene to produce amounts ranging from 16% to 80% of endogenous levels. In the homozygous state, these alleles make models available for studying recessive retinal degeneration and the role of protein levels in disease progression. In the heterozygous state, at which they do not perturb rod-cell structure, these alleles provide an exquisitely sensitive and specific signal for rod-cell status and for the structure and function of the gene itself. Thus, they provide a sensitive way to monitor the progress of retinal degeneration and the efficacy of gene-based therapies in dissected retinas and through noninvasive techniques in the retinas of living mice.


Discussion

The localization signals of many epithelial and neuronal proteins are currently being sought as well as the cellular components that interact with these signals. In the retinal degenerative disease retinitis pigmentosa, it has been hypothesized that a group of naturally occurring mutations clustered at the COOH terminus of rhodopsin might potentiate the disease by disrupting targeting of rhodopsin. Many in vitro studies support this hypothesis but because of the lack of a suitable expression system, the essential premise of the theory, that a sequence within the COOH terminus is a ROS targeting signal, has remained untested in vivo. Studies have been done in transgenic mice, rats, and pigs but have not excluded contributions from the upstream domains of rhodopsin.

We have now identified a ROS localization signal within the cytoplasmic tail of rhodopsin. By both loss and gain of function, we have shown that the last eight amino acids of rhodopsin are necessary and sufficient for ROS localization when the peptide sequence is membrane bound. Truncation of the distal five amino acids or mutation of the penultimate proline of rhodopsin resulted in partial mislocalization of the fusion proteins to RIS membranes, whereas the fusion protein containing the entire COOH-terminal tail targeted exclusively to the ROS. Furthermore, the COOH-terminal octapeptide of rhodopsin contains sufficient information to direct a predominantly RIS-synapse–localized protein (GFP-AAR) to the ROS (GFP-AAR[CC]rho8). This is the first demonstration of a gain of ROS sorting function of a rhodopsin domain. Although the mechanisms governing polarized targeting of neuronal proteins are widely studied, only a few amino acid–based axonal or dendritic targeting motifs have been identified. There were no obvious sequence homologies among the last eight amino acids of rhodopsin and the targeting signals of these other proteins.

Rhodopsin localization in rods may involve several steps, not all of which are necessarily relevant in epithelial cells, and vice versa. In a study of rhodopsin localization in MDCK cells, rhodopsin exhibited polarized apical distribution. However, truncation of the COOH-terminal 32, but not 22, residues resulted in partial delocalization to the basolateral surface (Chuang and Sung 1998). This result suggests that the apical targeting signal resides in the 10 amino acids between the two truncations, a region which includes the two palmitoylated cysteines. In rods, however, the targeting of GFP-AAR(CC)rho8 to the ROS and the delocalized distribution of GFP-CT44del25 demonstrate that the distal COOH-terminal region is important for ROS localization and not the region immediately surrounding the palmitoylated cysteines. Unpalmitoylated rhodopsin localized to the apical surface of MDCK cells, indicating that palmitoylation was not required for apical targeting (Chuang and Sung 1998). These results may be a consequence of a peptide-based apical targeting signal within rhodopsin's COOH-terminal 22 amino acids that functions in epithelial cells, but that in its absence, palmitoylation may be a second distinct signal for apical localization. We have shown, however, that palmitoylation cannot act by itself as a ROS localization signal in rod photoreceptors. The issues raised here highlight some of the problems with interpreting the targeting of neuronal proteins in polarized epithelial cells as a predictor of targeting in neuronal cells in vivo.

Two potential roles for rhodopsin's ROS localization signal are sorting of rhodopsin into the correct post-Golgi vesicular compartment and transporting rhodopsin-containing membranes from the TGN to the base of the connecting cilium. The abnormal accumulation, in the RIS plasma membrane, Golgi, and synapse, of the GFP fusion proteins lacking the intact ROS localization signal (Fig. 4, A–C, and Fig. 6A and Fig. B) can most easily be explained by their inability to sort to the proper post-Golgi vesicles. Without the proper ROS signal, individual GFP fusion proteins might randomly associate with any or all vesicles exiting the Golgi including those destined for the ROS, lateral plasma membrane, and synapse. In broken retinal cell preparations, COOH-terminal rhodopsin peptides and antibodies to this region inhibit the production of post-Golgi vesicles, also suggesting that this signal is required for sorting (Deretic et al. 1996, Deretic et al. 1998). The ability of wild-type, but not mutant, rhodopsin COOH terminus to interact directly with Tctex-1 (a cytoplasmic dynein light chain) was interpreted to indicate involvement of the signal in vectorial transport of rhodopsin-containing membranes (Tai et al. 1999). However, the rate of rhodopsin production precludes the possibility that Tctex-1 could transport individual rhodopsin molecules. Given that post-Golgi vesicles containing rhodopsin have a mean diameter of 300 nm (Deretic and Papermaster 1991), the density of rhodopsin on disk membranes is 20,000 μm 2 (Chen and Hubbel 1973), and the density of molecules on a periciliary vesicle is ∼40% that found in the disks (Besharse and Pfenninger 1980), a post-Golgi vesicle would contain ∼2,000 rhodopsin molecules. This estimate coupled with the low ratio of transgene product to endogenous rhodopsin (<1:2) suggests that there should be abundant rhodopsin COOH termini present in each vesicle to supply the required targeting signal and to drive transport of both rhodopsin and fusion protein to the ROS. We do in fact see this bulk flow phenomenon to a certain extent (see discussion below). However, we see that even transgene products lacking an intact targeting signal expressed at very low levels consistently delocalized to RIS membranes. The probability of a vesicle containing only the mutant rhodopsin COOH-terminal fusions, and therefore entirely unable to interact with Tctex-1, due to random sorting is exceedingly small. Even in human retinitis pigmentosa disease states where 50% of the rhodopsin molecules are mutated (i.e., 1:1 ratio), the vast majority of vesicles should contain abundant targeting information. Therefore, a sorting event, involving the COOH terminus, must exist that separates individual rhodopsin molecules from proteins destined for other parts of the cell and Tctex-1, given its specificity, may even be a part of that event. Expression of our GFP fusion proteins did not disrupt general transport mechanisms since endogenous rhodopsin did not delocalize. Furthermore, Green et al. 2000 showed that peripherin and the cGMP-gated channel localization were, likewise, unaffected in transgenic rats by the presence of a rhodopsin COOH-terminal truncation mutant.

A paraciliary transport pathway has been proposed in which rhodopsin-bearing vesicles bud from the apex of the RIS and fuse with nascent disks of the ROS (Besharse and Wetzel 1995). In transgenic mice, mutant rhodopsin (P347S) is released in vesicles into the interphotoreceptor space (Li et al. 1996). This might support a paraciliary pathway if the COOH terminus of rhodopsin were required for fusion of the extracellular rhodopsin-bearing vesicles with nascent disks. However, our study and other studies of transgenic animals expressing COOH-terminal rhodopsin mutants did not reveal the same vesicular accumulation (Sung et al. 1994 Li et al. 1998 Green et al. 2000). Alternatively, rhodopsin may be transported to the ROS via the cilium after docking at the periciliary ridge complex (Peters et al. 1983 Papermaster et al. 1985 Wolfram and Schmitt 2000). Additional sorting steps may take place at these locations.

Finally, the distal amino acids of rhodopsin may form an ROS retention signal as well as, or rather than, a targeting signal. Because rhodopsin exhibits a high degree of rotational and lateral mobility in disk membranes (Brown 1972 Poo and Cone 1974), a mechanism exists to limit reentry from the ROS to the RIS plasma membrane. However, a defect in retention alone cannot explain the presence of delocalized fusion proteins in the Golgi since loss of retention would result primarily in mislocalization to the RIS plasma membrane. It is possible, however, that rhodopsin in the lateral membrane might be endocytosed and recycled to the Golgi. Kinetic labeling studies would be required to determine if this is the case.

In our experimental system, we used posttranslational lipid modifications to confer membrane association properties to the fusion proteins. Membrane targeting via palmitoylation and/or myristoylation is well documented (Pellman et al. 1985 Gonzalo and Linder 1998 McCabe and Berthiaume 1999 Resh 1999). Since the overwhelming majority of both membrane lipids and proteins synthesized by photoreceptors are destined for the ROS, membrane proteins that lack a ROS localization signal may be passively cotransported with the rhodopsin-bearing vesicles (i.e., bulk flow). This would explain the high levels of GFP-CT44del5, GFP-CT44del25, GFP-CT44P353S, GFP-CT44P353L, GFP-AAR(CC)rho6, and mGFP in the ROS even though these molecules lack the ROS targeting signal. In contrast, GFP-AAR appeared to be almost excluded from the ROS. The COOH-terminal tail of AAR may contain a synaptic sorting/retention signal recognized by photoreceptors which is partially disrupted by the addition of a second cysteine in GFP-AAR(CC) or eliminated by removal of its last eight residues in GFP-AAR(CC)rho8. Expression of a COOH-terminal mutant rhodopsin in a rhodopsin knockout mouse could unambiguously address the issue of bulk flow in photoreceptors.

Although the role of COOH-terminal palmitoylation has been elucidated in other heptahelical G protein–coupled receptors (Moffett et al. 1996 Tanaka et al. 1998), the function of rhodopsin palmitoylation is unclear. Its role on rhodopsin phosphorylation, transducin activation, and rhodopsin regeneration remains controversial (Morrison et al. 1991 Karnik et al. 1993). Recently, Sachs et al. 2000 proposed that the palmitoyl moieties form a second retinal binding pocket. A potential role for the palmitoylation of rhodopsin, suggested by our results, may be to anchor the extreme COOH terminus in an orientation and proximity to the membrane that is optimal for interaction with its cognate sorting/transport components. It is important to note, however, that many cone opsins are not palmitoylated but achieve polarized outer segment localization in their respective cells (Ostrer et al. 1998).

Although we showed that the last eight amino acids of rhodopsin can direct ROS localization, both mGFP-CT9 and mGFP-CT25 delocalized, to different extents, to RIS membranes. It is possible that complete ROS localization of these two fusion proteins was inhibited through steric hindrance of the rhodopsin peptide by GFP with the cell's sorting/transport components. However, mGFP-CT44C322/323S also partially delocalized in contrast to the palmitoylated version of the same protein (mGFP-CT44) which targeted only to ROS, even though the lengths of these two rhodopsin peptides were identical. Although palmitoylation has been described as a reversible process, the relative absence of GFP-CT44 in the cytoplasm suggests that it was predominantly attached to membranes, and thus at least one of the cysteines was palmitoylated at any given time.

Colocalization of palmitoylated GFP fusion proteins with TR-WGA–labeled intracellular membranes indicates that they were present in Golgi/post-Golgi membranes and therefore may have used the same trafficking pathway as rhodopsin. It is intriguing that previous studies of rhodopsin missorting in transgenic animals have not documented accumulation in the Golgi. This may be due to the fact that detection of the intrinsic fluorescence of our fusion proteins may be more sensitive than indirect antibody labeling. Although we used only the COOH terminus of rhodopsin and not the entire protein, a rhodopsin–GFP fusion protein which lacked the final 14 amino acids of rhodopsin was also detected in the Golgi as well as the RIS plasma membrane and synapse (our unpublished results). The palmitoylation consensus sequence might act as an ER recruiting signal since palmitoylation of rhodopsin occurs in the ER before entering the Golgi (St. Jules and O'Brien 1986 St. Jules et al. 1990). Once attached to ER membranes, the palmitoylated fusion proteins can follow normal membrane protein transport pathways. In contrast, myristoylation is a cotranslational process (Wilcox et al. 1987). Once released from the ribosomes, the myristoylated fusion proteins distributed throughout all RIS membranes, including mitochondrial membranes and the plasma membrane, where they would not be accessible to the post-Golgi sorting/transport pathway. This provides an alternate explanation for why the myristoylated, but not palmitoylated, GFP fusion proteins (e.g., mGFP-CT9) were not fully localized to the ROS.

The mislocalization of mutant rhodopsins may be involved in the pathogenesis of rod photoreceptors in retinitis pigmentosa and macular degeneration. It is therefore necessary to determine how rhodopsin achieves its specific localization in order to understand how mutations disrupt the normal process. Using transgenic frog photoreceptors as an expression system has aided us in approaching and answering some of these issues. In the future it will also be important to determine how other ROS proteins achieve their localization.


Why GFP Is Important

No one actually knows the function of bioluminescence or fluorescence in the crystal jelly. Roger Tsien, the American biochemist who shared the 2008 Nobel Prize in Chemistry, speculated the jellyfish might be able to change the color of its bioluminescence from the pressure change of changing its depth. However, the jellyfish population in Friday Harbor, Washington, suffered a collapse, making it difficult to study the animal in its natural habitat.

While the importance of fluorescence to the jellyfish is unclear, the effect the protein has had on scientific research is staggering. Small fluorescent molecules tend to be toxic to living cells and negatively affected by water, limiting their use. GFP, on the other hand, can be used to see and track proteins in living cells. This is done by joining the gene for GFP to the gene of a protein. When the protein is made in a cell, the fluorescent marker is attached to it. Shining a light at the cell makes the protein glow. Fluorescence microscopy is used to observe, photograph, and film living cells or intracellular processes without interfering with them. The technique works to track a virus or bacteria as it infects a cell or to label and track cancer cells. In a nutshell, the cloning and refining of GFP have made it possible for scientists to examine the microscopic living world.

Improvements in GFP have made it useful as a biosensor. The modified proteins as act molecular machines that react to changes in pH or ion concentration or signal when proteins bind to each other. The protein can signal off/on by whether or not it fluoresces or can emit certain colors depending on the conditions.


What is the difference between GFP and rhodopsin? - Biology

Figure 1: Illustration of some of the palette of fluorescent proteins that has revolutionized cell biology. (A) Fluorescent proteins spanning a range of excitation and emission wavelengths. (B) Illustration of a petri dish with bacteria harboring eight different colors of fluorescent protein and used to “paint” an idyllic beach scene. (Adapted from: R. Y. Tsien, Nobel lecture, Integr. Biol.,2, 77-93, (2010).)

Fluorescent proteins have become a dominant tool for the exploration of the dynamics and localization of the macromolecular contents of living cells. Given how pervasive the palette of different fluorescent proteins shown in Figure 1 with their many colors, and properties has become, it is incredible that we have really only seen a decade of concerted effort with these revolutionary tools. Indeed, it is difficult to imagine any part of biology that has not been touched in some way or another (and often deeply) by the use of fluorescent reporter proteins.

Figure 2: Schematic diagram of the chromophore formation in maturing enhanced green fluorescent protein (EGFP). (A) The prematuration EGFP fluorophore tripeptide amino acid sequence (Thr65-Tyr66-Gly67) stretched into a linear configuration. The first step in maturation is a series of torsional adjustments (B) and (C). These torsional adjustments allow a nucleophilic attack that results in formation of a ring system (the cyclization step). (D) Fluorescence occurs following oxidation of the tyrosine by molecular oxygen. The final conjugated and fluorescent core atoms are shaded. (Adapted from: The Fluorescent Protein Color Palette, Scott G. Olenych, Nathan S. Claxton, Gregory K. Ottenberg, Michael W. Davidson, 2007).

Table 1: Common fluorescent proteins maturation times. Because different approaches and conditions still give quite different values one should be very careful in studies where the maturation time can affect the conclusions. In mCherry there are indications of two time scales, the first leading to fluorescence at a different wavelength regime (Khmelinskii et al., 2012). Values are rounded to one significant digit. Comprehensive table can be found at Lizuka et al, 2011. For definitions of fluorophores via mutations relative to WT see Table S2 of Shaner et al, 2005.

However, as a tool for exploring the many facets of cellular dynamics, fluorescent proteins have both advantages and disadvantages. Once a fluorescent protein is expressed it has to go through several stages until it becomes functional as shown in Figure 2. These processes are together termed maturation. Until completion of the maturation process, the protein, even though already synthesized, is not fluorescent. To study dynamics, it is most useful if there is a separation of time scales between the reporter maturation process (which preferably should take place on “fast” time scales) and the dynamics of the process of real interest (that should be much slower than the maturation time). The first stage in the maturation process (not depicted) is the most intuitive and refers to the protein folding itself, which is relatively fast and should take less than a minute, assuming there is no aggregation. The next stage is a torsional rearrangement (Figure 2B, C) of what can be thought of as the active site of the fluorophore, the amino acids where the conjugated electrons that will fluoresce are located. The next step, known as cyclization (where a ring is formed between two amino acids, Figure 2 C, D), is longer but still fast in comparison to the final and rate-limiting step of oxidation. In this final oxidation step, molecular oxygen grabs electrons from the fluorophore, creating the final system of conjugated bonds. All these steps are a prerequisite to making the active site fluoresce.

There are only a limited number of reliable measurements of the maturation time that we tried to summarize in Table 1, and the values are still far from being completely agreed upon. One approach to measure fluorophore maturation is by moving from anaerobic growth where the fluorophore protein is expressed but cannot perform the slowest step of oxidation to aerobic conditions and watching the rate of fluorescent signal formation. More commonly, inducible promoters or cycloheximide induced translation arrest are used. Nagai et al. (BNID 103780) measure a time scale of less than 5 minutes for the maturation of YFP and 7 minutes for the corresponding maturation of GFP in E. coli. By way of contrast, Gordon et al. (BNID 102974) report a time scale of ≈40 minutes for the maturation of YFP and a very slow ≈50 minutes for the maturation of CFP though part of the difference can be explained by the fact that in this case the measurements were carried out in yeast at 25 o C. The measurements were done by inducing expression and after 30 minutes inhibiting translation using cycloheximide. The dynamics of continued fluorophore accumulation even after no new proteins are synthesized was used to infer the maturation time scale. Note that for many of the processes that occur during a cell cycle such as expression of genes in response to environmental cues, the maturation time can be a substantial fraction of the time scale of the process of interest. If a marathon runner stops for a drink in the middle of a race, this will hardly affect the overall time of the racer’s performance. On the other hand, if the runner stops to have a massage, this will materially affect the time scale at which the racer completes the race. By analogy with the runner stopping at a restaurant, the maturation time can seriously plague our ability to accurately monitor the dynamics of a variety of cellular processes.

Chromophore maturation effectively follows first-order kinetics in most studies performed. As a result, this implies that we will find a small fraction of functional flurophores much earlier than the maturation time. Still, to have the majority of the population active, the characteristic time scale we need to wait is roughly the maturation time itself. This effect results in a built-in delay in the reporting system and should be heeded when estimating response times based on fluorescent reporters. Similarly, if translation is being stopped (say by the use of a ribosome inhibitor such as cyclohexamide) one would still have a period of time where some proteins that were translated before the inhibition are coming “online” and add to the signal. This again should be taken into account when estimating degradation times.

Another dynamical feature of these proteins that can make them tricky for precisely characterizing cellular dynamics is the existence of photobleaching. This process has a characteristic time scale of tens of seconds using standard levels of illumination and magnification. This value means that after a continuous exposure to illumination for several tens of seconds, the fluorescent intensity will have decayed to 1/e of its original value. Though sometimes a nuisance, recently this apparent disadvantage has been used as a trick both in the context of fluorescence recovery after photobleaching (FRAP) that allows inference about diffusion rates and in superresolution microscopy techniques where the bleaching of individual fluorophores makes it possible to localize these proteins with nanometer scale resolution.

Differences in maturation times of different fluorophores were recently turned into a way to measure rates of degradation and translocation without the need for time course measurements (A. Khmelinskii et al., Nat. Biotech., 30:708, 2012). The protein of interest was fused not to one, but to two fluorescent tags, a fast maturing GFP, so called superfolder, and a slower maturing mCherry. The ratio of intensities was measured and this can serve as a built-in timer. If the protein of interest is short lived, the slowly maturing tag would often not have time enough to fluoresce before the protein is degraded, and its intensity ratio to the quickly maturing tag would be low. At the other extreme, if the protein is long lived, there is ample time for the more slowly maturing tag to fluoresce, and its ratio to the fast dividing tag would be high. The ratio of intensities thus serves as a timer which was used for example to show that daughter cells tend to get the old copies of some protein complexes such as spindle pole bodies and nuclear pore complexes, while the mothers retain the newly formed copies.


OTHER METHODS OF COVALENTLY LABELING A PROTEIN

As well as using intrinsically fluorescent proteins related in structure or sequence to GFP, there are three other methods of covalently labeling a protein (Crivat and Taraska, 2012 ). First, self-labeling enzymes (Halo tags and SNAP/CLIP tags) can be used. These are used to overcome the limitations of fluorescent proteins, such as the fact that the photophysical properties of fluorescent proteins are not as good as organic dyes. For example, (1) fluorescent proteins can “blink” or emit light intermittently, (2) fluorescent proteins are not very bright, (3) some are not photostable, and (4) fluorescent proteins have limited colors and chemistry. This class of protein-based fusion tags can catalyze the covalent autoattachment of an organic fluorophore inside living cells. The enzymes are fused to a protein, and the pair is labeled by introduction of a fluorescent ligand, which covalently reacts with the fusion tag. These tags are similar in size to a fluorescent protein. The proteins are not innately fluorescent when expressed but become fluorescent when the cells are exposed to a fluorescent ligand.

Second, an enzyme can be used to covalently attach a fluorophore ligand to another protein or peptide. An example of this is the small 9-kDa acyl carrier protein (ACP) tag, which can be covalently labeled with CoA derivatives by the enzyme phosphopantetheine transferase (PPTase) (George et al., 2004 ). The ACP fusion protein is incubated with the CoA derivatives, which are labeled with fluorescence. However, since substrates for this system are not membrane-permeable, use of ACP is restricted to extracellular proteins.

Third, small cell-permeable biarsenical dyes have been developed. The smallest and most successful genetically encoded tag for covalent small-fluorophore labeling is the tetracysteine-biarsenical dyes, FLAsH and ReAsH. A short peptide sequence is genetically introduced into the sequence of a target protein. This sequence can specifically react with a membrane-permeable biarsenical dye. There are two advantages of biarsenical dyes. First, the targeting motifs are small. Therefore, there is less opportunity for introduced sequences to disrupt the overall fold and function of the labeled protein. The second advantage is that pulse-labeling procedures are easy. There are three problems when using biarsenical dyes: the level of background fluorescence, cellular toxicity of the ligands, and the effect of the tag on protein function or localization.

Covalent labeling of cellular proteins can be achieved with aminoreactive molecules, such as the succinimide ester of carboxyfluorescein (CFSE) (Wallace and Muirhead, 2007 ). One of the most common uses of CFSE labeling has been in combination with antibodies against phenotypic and functional markers (e.g., lymphocyte subsets, intracellular cytokines, and hematopoietic lineage) to allow monitoring of proliferative responses in complex populations in vitro. CFSE has several advantages for in vitro or ex vivo tagging of immune cells. This method can be applied to most any cell type as long as toxicity due to overlabeling is avoided. Very high-intensity labeling can be achieved because a large fraction of cellular proteins becomes labeled. A limitation is that the specificity, affinity, and/or function of labeled proteins may be compromised if critical residues involved in receptor/coreceptor binding or antigen recognition sites are modified. This is because covalent protein labeling is random (any succinimide reactive residue that is accessible has the potential to become labeled). Another limitation is that, unlike stable genetic tags that have the same intensity in all daughter cells, CFSE intensity is halved with each round of cell division, which limits the length of time that daughter cells can be distinguished from unlabeled cells. Additionally, fixation, or the presence of necrotic or apoptotic cells, can lose labeled protein as fragmentation or shedding of apoptotic vesicles occurs. This can considerably distort CFSE fluorescence intensity distributions and complicate proliferation analysis. Reducing the concentration of CFSE used for labeling can limit some of the above problems, but as a result, the number of cell divisions that can be distinguished is reduced.


Discussion

The localization signals of many epithelial and neuronal proteins are currently being sought as well as the cellular components that interact with these signals. In the retinal degenerative disease retinitis pigmentosa, it has been hypothesized that a group of naturally occurring mutations clustered at the COOH terminus of rhodopsin might potentiate the disease by disrupting targeting of rhodopsin. Many in vitro studies support this hypothesis but because of the lack of a suitable expression system, the essential premise of the theory, that a sequence within the COOH terminus is a ROS targeting signal, has remained untested in vivo. Studies have been done in transgenic mice, rats, and pigs but have not excluded contributions from the upstream domains of rhodopsin.

We have now identified a ROS localization signal within the cytoplasmic tail of rhodopsin. By both loss and gain of function, we have shown that the last eight amino acids of rhodopsin are necessary and sufficient for ROS localization when the peptide sequence is membrane bound. Truncation of the distal five amino acids or mutation of the penultimate proline of rhodopsin resulted in partial mislocalization of the fusion proteins to RIS membranes, whereas the fusion protein containing the entire COOH-terminal tail targeted exclusively to the ROS. Furthermore, the COOH-terminal octapeptide of rhodopsin contains sufficient information to direct a predominantly RIS-synapse–localized protein (GFP-AAR) to the ROS (GFP-AAR[CC]rho8). This is the first demonstration of a gain of ROS sorting function of a rhodopsin domain. Although the mechanisms governing polarized targeting of neuronal proteins are widely studied, only a few amino acid–based axonal or dendritic targeting motifs have been identified. There were no obvious sequence homologies among the last eight amino acids of rhodopsin and the targeting signals of these other proteins.

Rhodopsin localization in rods may involve several steps, not all of which are necessarily relevant in epithelial cells, and vice versa. In a study of rhodopsin localization in MDCK cells, rhodopsin exhibited polarized apical distribution. However, truncation of the COOH-terminal 32, but not 22, residues resulted in partial delocalization to the basolateral surface (Chuang and Sung 1998). This result suggests that the apical targeting signal resides in the 10 amino acids between the two truncations, a region which includes the two palmitoylated cysteines. In rods, however, the targeting of GFP-AAR(CC)rho8 to the ROS and the delocalized distribution of GFP-CT44del25 demonstrate that the distal COOH-terminal region is important for ROS localization and not the region immediately surrounding the palmitoylated cysteines. Unpalmitoylated rhodopsin localized to the apical surface of MDCK cells, indicating that palmitoylation was not required for apical targeting (Chuang and Sung 1998). These results may be a consequence of a peptide-based apical targeting signal within rhodopsin's COOH-terminal 22 amino acids that functions in epithelial cells, but that in its absence, palmitoylation may be a second distinct signal for apical localization. We have shown, however, that palmitoylation cannot act by itself as a ROS localization signal in rod photoreceptors. The issues raised here highlight some of the problems with interpreting the targeting of neuronal proteins in polarized epithelial cells as a predictor of targeting in neuronal cells in vivo.

Two potential roles for rhodopsin's ROS localization signal are sorting of rhodopsin into the correct post-Golgi vesicular compartment and transporting rhodopsin-containing membranes from the TGN to the base of the connecting cilium. The abnormal accumulation, in the RIS plasma membrane, Golgi, and synapse, of the GFP fusion proteins lacking the intact ROS localization signal (Fig. 4, A–C, and Fig. 6A and Fig. B) can most easily be explained by their inability to sort to the proper post-Golgi vesicles. Without the proper ROS signal, individual GFP fusion proteins might randomly associate with any or all vesicles exiting the Golgi including those destined for the ROS, lateral plasma membrane, and synapse. In broken retinal cell preparations, COOH-terminal rhodopsin peptides and antibodies to this region inhibit the production of post-Golgi vesicles, also suggesting that this signal is required for sorting (Deretic et al. 1996, Deretic et al. 1998). The ability of wild-type, but not mutant, rhodopsin COOH terminus to interact directly with Tctex-1 (a cytoplasmic dynein light chain) was interpreted to indicate involvement of the signal in vectorial transport of rhodopsin-containing membranes (Tai et al. 1999). However, the rate of rhodopsin production precludes the possibility that Tctex-1 could transport individual rhodopsin molecules. Given that post-Golgi vesicles containing rhodopsin have a mean diameter of 300 nm (Deretic and Papermaster 1991), the density of rhodopsin on disk membranes is 20,000 μm 2 (Chen and Hubbel 1973), and the density of molecules on a periciliary vesicle is ∼40% that found in the disks (Besharse and Pfenninger 1980), a post-Golgi vesicle would contain ∼2,000 rhodopsin molecules. This estimate coupled with the low ratio of transgene product to endogenous rhodopsin (<1:2) suggests that there should be abundant rhodopsin COOH termini present in each vesicle to supply the required targeting signal and to drive transport of both rhodopsin and fusion protein to the ROS. We do in fact see this bulk flow phenomenon to a certain extent (see discussion below). However, we see that even transgene products lacking an intact targeting signal expressed at very low levels consistently delocalized to RIS membranes. The probability of a vesicle containing only the mutant rhodopsin COOH-terminal fusions, and therefore entirely unable to interact with Tctex-1, due to random sorting is exceedingly small. Even in human retinitis pigmentosa disease states where 50% of the rhodopsin molecules are mutated (i.e., 1:1 ratio), the vast majority of vesicles should contain abundant targeting information. Therefore, a sorting event, involving the COOH terminus, must exist that separates individual rhodopsin molecules from proteins destined for other parts of the cell and Tctex-1, given its specificity, may even be a part of that event. Expression of our GFP fusion proteins did not disrupt general transport mechanisms since endogenous rhodopsin did not delocalize. Furthermore, Green et al. 2000 showed that peripherin and the cGMP-gated channel localization were, likewise, unaffected in transgenic rats by the presence of a rhodopsin COOH-terminal truncation mutant.

A paraciliary transport pathway has been proposed in which rhodopsin-bearing vesicles bud from the apex of the RIS and fuse with nascent disks of the ROS (Besharse and Wetzel 1995). In transgenic mice, mutant rhodopsin (P347S) is released in vesicles into the interphotoreceptor space (Li et al. 1996). This might support a paraciliary pathway if the COOH terminus of rhodopsin were required for fusion of the extracellular rhodopsin-bearing vesicles with nascent disks. However, our study and other studies of transgenic animals expressing COOH-terminal rhodopsin mutants did not reveal the same vesicular accumulation (Sung et al. 1994 Li et al. 1998 Green et al. 2000). Alternatively, rhodopsin may be transported to the ROS via the cilium after docking at the periciliary ridge complex (Peters et al. 1983 Papermaster et al. 1985 Wolfram and Schmitt 2000). Additional sorting steps may take place at these locations.

Finally, the distal amino acids of rhodopsin may form an ROS retention signal as well as, or rather than, a targeting signal. Because rhodopsin exhibits a high degree of rotational and lateral mobility in disk membranes (Brown 1972 Poo and Cone 1974), a mechanism exists to limit reentry from the ROS to the RIS plasma membrane. However, a defect in retention alone cannot explain the presence of delocalized fusion proteins in the Golgi since loss of retention would result primarily in mislocalization to the RIS plasma membrane. It is possible, however, that rhodopsin in the lateral membrane might be endocytosed and recycled to the Golgi. Kinetic labeling studies would be required to determine if this is the case.

In our experimental system, we used posttranslational lipid modifications to confer membrane association properties to the fusion proteins. Membrane targeting via palmitoylation and/or myristoylation is well documented (Pellman et al. 1985 Gonzalo and Linder 1998 McCabe and Berthiaume 1999 Resh 1999). Since the overwhelming majority of both membrane lipids and proteins synthesized by photoreceptors are destined for the ROS, membrane proteins that lack a ROS localization signal may be passively cotransported with the rhodopsin-bearing vesicles (i.e., bulk flow). This would explain the high levels of GFP-CT44del5, GFP-CT44del25, GFP-CT44P353S, GFP-CT44P353L, GFP-AAR(CC)rho6, and mGFP in the ROS even though these molecules lack the ROS targeting signal. In contrast, GFP-AAR appeared to be almost excluded from the ROS. The COOH-terminal tail of AAR may contain a synaptic sorting/retention signal recognized by photoreceptors which is partially disrupted by the addition of a second cysteine in GFP-AAR(CC) or eliminated by removal of its last eight residues in GFP-AAR(CC)rho8. Expression of a COOH-terminal mutant rhodopsin in a rhodopsin knockout mouse could unambiguously address the issue of bulk flow in photoreceptors.

Although the role of COOH-terminal palmitoylation has been elucidated in other heptahelical G protein–coupled receptors (Moffett et al. 1996 Tanaka et al. 1998), the function of rhodopsin palmitoylation is unclear. Its role on rhodopsin phosphorylation, transducin activation, and rhodopsin regeneration remains controversial (Morrison et al. 1991 Karnik et al. 1993). Recently, Sachs et al. 2000 proposed that the palmitoyl moieties form a second retinal binding pocket. A potential role for the palmitoylation of rhodopsin, suggested by our results, may be to anchor the extreme COOH terminus in an orientation and proximity to the membrane that is optimal for interaction with its cognate sorting/transport components. It is important to note, however, that many cone opsins are not palmitoylated but achieve polarized outer segment localization in their respective cells (Ostrer et al. 1998).

Although we showed that the last eight amino acids of rhodopsin can direct ROS localization, both mGFP-CT9 and mGFP-CT25 delocalized, to different extents, to RIS membranes. It is possible that complete ROS localization of these two fusion proteins was inhibited through steric hindrance of the rhodopsin peptide by GFP with the cell's sorting/transport components. However, mGFP-CT44C322/323S also partially delocalized in contrast to the palmitoylated version of the same protein (mGFP-CT44) which targeted only to ROS, even though the lengths of these two rhodopsin peptides were identical. Although palmitoylation has been described as a reversible process, the relative absence of GFP-CT44 in the cytoplasm suggests that it was predominantly attached to membranes, and thus at least one of the cysteines was palmitoylated at any given time.

Colocalization of palmitoylated GFP fusion proteins with TR-WGA–labeled intracellular membranes indicates that they were present in Golgi/post-Golgi membranes and therefore may have used the same trafficking pathway as rhodopsin. It is intriguing that previous studies of rhodopsin missorting in transgenic animals have not documented accumulation in the Golgi. This may be due to the fact that detection of the intrinsic fluorescence of our fusion proteins may be more sensitive than indirect antibody labeling. Although we used only the COOH terminus of rhodopsin and not the entire protein, a rhodopsin–GFP fusion protein which lacked the final 14 amino acids of rhodopsin was also detected in the Golgi as well as the RIS plasma membrane and synapse (our unpublished results). The palmitoylation consensus sequence might act as an ER recruiting signal since palmitoylation of rhodopsin occurs in the ER before entering the Golgi (St. Jules and O'Brien 1986 St. Jules et al. 1990). Once attached to ER membranes, the palmitoylated fusion proteins can follow normal membrane protein transport pathways. In contrast, myristoylation is a cotranslational process (Wilcox et al. 1987). Once released from the ribosomes, the myristoylated fusion proteins distributed throughout all RIS membranes, including mitochondrial membranes and the plasma membrane, where they would not be accessible to the post-Golgi sorting/transport pathway. This provides an alternate explanation for why the myristoylated, but not palmitoylated, GFP fusion proteins (e.g., mGFP-CT9) were not fully localized to the ROS.

The mislocalization of mutant rhodopsins may be involved in the pathogenesis of rod photoreceptors in retinitis pigmentosa and macular degeneration. It is therefore necessary to determine how rhodopsin achieves its specific localization in order to understand how mutations disrupt the normal process. Using transgenic frog photoreceptors as an expression system has aided us in approaching and answering some of these issues. In the future it will also be important to determine how other ROS proteins achieve their localization.


Watch the video: Green Fluorescent Protein GFP: GFP Tagging and USe (August 2022).