Memorizing spatiotemporal patternsAtsushi Miyawaki & Satoshi Karasawa

Live samples are intrinsically highly dynamic, yet techniques to monitor these complex environments usually reflect snapshots, thus making time-lapse imaging necessary to explore temporal progression of biological functions. Recent results indicate that exploiting some basic features of fluorescent protein maturation, such as green-to-red maturation of engineered proteins, should allow probing of temporally regulated information.

Real-time fluorescence imaging of live cells is useful for the identification and establishment of the hierarchy of many molecules involved in signal transduction. Signal transduction diagrams, in which arrows link molecules to indicate enzyme reactions and intermolecu-lar interactions, have become commonplace. However, to gain a better understanding of any signal transduction pathway, such diagrams must contain a fourth dimension in addition to the three spatial axes: time. Since the com-plementary DNA cloning of green fluorescent protein (GFP) in 1992 from bioluminescent jellyfish1 (Aequorea victoria) and later from its relatives2,3, researchers have attempted to devise tools that would enable direct visual-ization of biological functions. Microscopy using fluorescence resonance energy transfer (FRET)4,5 with GFP derivatives has offered a first glimpse of the spatiotemporal aspects of biological activities. Similarly, dual-color fluorescence cross-correlation spectroscopy (FCCS) is a promising technique for quanti-fying protein-protein interactions6. Although

FCCS does not provide direct images, it gives more quantitative information of interactions than FRET. Recently, a simple but efficient FCCS system has been developed that uses a pair of fluorescent proteins that can be excited at the same wavelength but that emit fluores-cence in different color regions7. Finally, bimo-lecular fluorescence complementation (BiFC) has been used to reliably analyze the occur-rence and subcellular localization of protein- protein interactions in live cells8–10. Although this technique has one limitation—the irrevers-ible association of the fluorescent protein frag-ments8—it can provide temporally integrated information about protein-protein interactions with high sensitivity. We will discuss how BiFC combined with ‘fluorescent timer’ proteins11 can expand the applications of this technique to monitor cellular processes in defined spatial and temporal measurements.

Chromophore formation: from green to redFluorescent proteins typically share the Xxx-Tyr-Gly motif, which serves as the chro-mophore-forming sequence. The known exceptions are the blue- and cyan-emitting variants of Aequorea GFP (BFP and CFP), which contain histidine and tryptophan instead of tyrosine in the tripeptide, respec-tively. The wild-type Aequorea GFP contains the tripeptide Ser65-Tyr66-Gly67. The process of formation of the basic green-emitting chro-mophore is illustrated in Figure 1a (ref. 12). Concomitant with (or following) the folding of the β-barrel is the formation of an imidazo-linone by a nucleophilic attack of the amide of Gly67 on the carbonyl of Ser65, followed by dehydration. Molecular oxygen then dehydro-genates the α-β bond of Tyr66 to conjugate its aromatic group with the imidazolinone. The

resulting chromophore is a 4-(p-hydroxyben-zylidene)-5-imidazolinone that emits green fluorescence. Many green-emitting fluorescent proteins retain this chromophore throughout their lifetime.

Of those proteins that continue to mature, red-emitting fluorescent proteins (RFPs) initially create the basic green-emitting chromophore 4-(p-hydroxybenzylidene)-5-imidazolinone, but then an additional reac-tion occurs to expand the π-conjugated system (Fig. 1a). This conversion occurs differently in Kaede13 and EosFP (ref. 14), two coral fluo-rescent proteins that contain the tripeptide His62-Tyr63-Gly64, a green chromophore that is photoconverted to red. UV irradiation promotes an unconventional cleavage within the protein between the amide nitrogen and the Cα at His62 via a formal β-elimination reaction (Fig. 1b). The subsequent forma-tion of a double bond between His62 Cα and His62 Cβ �extends the π-conjugation to the imidazole ring of His62, thereby creating a new red-emitting chromophore: 2-[(1E)-2-(5-imidazolyl)ethenyl]-4-(p-hydroxybenzyli-dene)-5-imidazolinone15,16.

In common RFPs, an additional oxidation reaction occurs autocatalytically to modify the green-emitting chromophore. In certain preco-cious RFPs, such as DsRed, this oxidation reac-tion is so efficient that the green intermediate exists only transiently (Fig. 1b, top)17,18. Slow and incomplete chromophore modification gives rise to residual green fluorescence, thus prohibiting the combined use of this molecule with green-emitting fluorescent proteins in dual-color labeling experiments. Recently, a new generation of RFPs and engineered vari-ants of DsRed featuring fast and complete green-to-red maturation has enabled research-ers to overcome these problems19.

Atsushi Miyawaki is in the Laboratory for Cell Function and Dynamics, Advanced Technology Development Group, Brain Science Institute, RIKEN, 2-1 Hirosawa, Wako-city, Saitama 351-0198, Japan, and the Life Function and Dynamics Project, Exploratory Research for Advanced Technology, JST, 2-1 Hirosawa, Wako-city, Saitama 351-0198, Japan. Satoshi Karasawa is in the Laboratory for Cell Function and Dynamics, Advanced Technology Development Group, Brain Science Institute, RIKEN, 2-1 Hirosawa, Wako-city, Saitama 351-0198, Japan and Amalgaam Co., Ltd., 2-9-3 Itabashi, Itabashi-ku, Tokyo 173-0004, Japan. e-mail: matsushi@brain.riken.jp

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Mutagenesis studies have also generated several DsRed mutants that mature completely but slowly. Comparative analysis of the DsRed mutants’ crystal structures suggested that the free space around the chromophore is critical for fast maturation20. In certain cases, slow maturation with a long-lived green state can be advantageous, especially for the analysis of the duration of gene expression in cells (Fig. 1b, bottom). E5, a mutant of DsRed, is particularly useful because it changes color from green to red with a predictable time course11. This fea-ture allows for the ratiometric analysis of the green-to-red emission as an estimate of the time elapsed after initiation of reporter gene expression. Therefore, E5 functions as a ‘fluo-rescent timer’ that yields temporal and spatial information regarding protein translation and target promoter activity. For instance, E5 was used to trace the expression of the homeobox gene Otx-2, which is involved in the pattern-ing of anterior structures in Xenopus laevis; green fluorescence indicated the onset of gene expression, whereas red fluorescence indicated that the expression was suppressed11.

This approach of using E5 has also been used to examine the functional segregation of secre-tory vesicles in neuroendocrine cells. By tagging atrial natriuretic factor (ANF) with E5, Duncan et al. analyzed the age-dependent distribution of large dense-core vesicles in bovine adrenal chromaffin cells21. They observed that newly assembled vesicles are immobile and docked at the plasma membrane shortly after biogenesis, whereas older vesicles are mobile and located at

greater distances from the plasma membrane. A similar approach has also been used to moni-tor insulin synthesis in vivo with a transgenic mouse that expresses proinsulin II tagged with E5 (ref. 22).

Like DsRed, E5 forms an obligate tetramer. Although oligomerization does not prevent the use of E5 for reporting gene expression, it may preclude its use in certain fusion protein applications. A monomeric version of fluores-cent timer proteins would be most suitable for analyzing age-dependent distribution of func-tional proteins.

Bimolecular fluorescence complementationDistinct proteins containing the N- and C-ter-minal halves of a fluorescent protein do not efficiently assemble a functional fluorescent protein if cotransfected into cells. The BiFC assay, then, is based on the reconstitution of a fluorescent protein from two nonfluores-cent fragments8–10. If these two fragments are brought into close proximity by a physical interaction between proteins fused to each frag-ment (Fig. 2), the β-barrel structure will be eas-ily reconstituted, and fluorescence will emerge. Despite a tendency for intrinsic association of two fragments23, which produces background signal, the BiFC has been used practically to analyze the occurrence and subcellular localiza-tion of protein-protein interactions in live cells. Aequorea GFP variants are usually separated between positions 154 and 155 or between positions 172 and 173, because these positions

are located in loop regions. Hu et al. initially used EYFP residues 1–154 (YN) and residues 155–238 (YC) as the two fragments for study-ing BiFC (ref. 24). By generating fusion pro-teins in which the basic leucine zipper (bZIP) and members of the Rel transcription factor families are fused to YN and YC, they identi-fied the intracellular locations where associa-tion between these proteins occurs. By using different Aequorea GFP variants, such as GFP, CFP and BFP, Hu et al. further extended this technique to multicolor BiFC, which permits direct visualization of multiple protein inter-actions within the same cell and allows for comparison of complex formation efficien-cies with different interacting partners25. The color range available for BiFC has expanded to red with the development of an improved monomeric red fluorescent protein (mRFP1-Q66T), which is amenable to fragmentation26. Furthermore, the efficiency of BiFC has been greatly improved27 by using newly identified fluorescent protein fragments derived from Venus28 and Cerulean29, the Aequorea GFP variants that show fast maturation.

Association of fluorescent protein fragments in BiFC has been proposed to be irreversible8,10. Therefore, a potential drawback of this assay is that the dimerization of the fused interacting proteins may also be irreversible. This could interfere with the cellular physiological state of the system and impede the investigation of dynamic protein interactions. However, irre-versible binding makes BiFC a highly sensitive assay, which may allow for the visualization of

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Figure 1 A comparison of schemes for the formation and maturation of chromophores in fluorescent proteins. The β-can structure represents the native conformation of the protein, while the denatured form is depicted as an irregular chain. π-conjugation for visible-light absorption is indicated in green or red. Neighboring amino acids (single-letter code) have been added. (a) Scheme for the formation of 4-(p-hydroxybenzylidene)-5-imidazolinone as the basic chromophore in green-emitting fluorescent proteins, including the wild-type Aequorea GFP. (b) Scheme for the formation of 4-(p-hydroxybenzylidene)-5-imidazolinone and its autocatalytic modification in DsRed (top) and E5 (bottom).

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transient or low-affinity interactions, such as those between enzymes and substrates30–35.

A unique BiFC system has also been devel-oped. Demidov et al. determined that the N-terminal EGFP fragment (residues 1–158) is large enough to develop a compact folded structure necessary for the synthesis of the chromophore36. Owing to this unique protein environment, however, the chromophore is not fluorescent; this preformation of the chromo-phore within the N-terminal fragment offers an advantage because complementation with the C-terminal fragment via DNA hybridiza-tion rapidly restores fluorescence. Importantly, the BiFC system seems to be reversible; the restored protein fluorescence was quenched by dissociating the DNA duplex.

Mapping protein-protein interactions temporally deep within tissuesThe power of this sensitivity of BiFC, primar-ily caused by the irreversible binding, may also be harnessed to obtain integrated fluo-rescence signals for protein-protein interac-tions that occur before imaging (Fig. 3). This feature may be particularly useful for studying biological events transpiring in deep regions

such as tissue in the brain. Because some in vivo imaging technologies use visible light, the depth of penetration into samples is limited by scattering, thus rendering fluorescent imag-ing of deep brain tissues challenging. Though the development of long-wavelength dyes and multiphoton excitation microscopy armed with modern nonlinear optics has increased the depth of penetration into brain tissue37, rela-tively more reliable signals can still be obtained from sections of fixed brain samples given that spatial information should be independent of depth from the surface. Using transgenic mice that coexpress two components for BiFC under

an inducible promoter, such as the Tet-on or Tet-off expression system38, we may be able to examine the biological events that occur dur-ing a certain time interval within any given section. This approach may reveal alterations in the spatial pattern of fluorescence, which may reflect changes in neural function, such as those in response to a behavioral task.

The next step is to extract temporal informa-tion from such imaging of brain sections. If a fluorescent timer protein can be compatible with fragmentation and reconstitution, BiFC using the protein will permit studies that aim to examine the temporal aspect of protein-protein interactions. The emission ratio of green to red will reveal their age. Green would indicate young, whereas red would indicate old. As mentioned previously, this approach can be further extended to analyzing activity-dependent functions in fixed tissues. Because the temporal resolution is rather poor, well-designed timing of the expression of the two components will be necessary. Of course, expanding the range of t1/2 for green-to-red maturation of the BiFC technique would also be a useful improvement.

Sequential labeling with differently colored organic dyesRecently, various techniques have been devel-oped for site-specific labeling of proteins in live cells with small-molecule probes39. One of the promising techniques is biarsenic fluo-rophore labeling40, which labels proteins that have been genetically altered to contain a tetra-cysteine motif (CCXXCC). The small-molecule probes include FlAsH and ReAsH. FlAsH is created by introducing two arsenoxide groups into fluorescein. ReAsH is a red analog of FlAsH and has been synthesized using the red

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Figure 2 Principle of the bimolecular fluorescence complementation assay. The assay is based on the formation of a fluorescent complex (β-barrel) by fragments of a fluorescent protein brought together by the association of two interacting partners (A and B) fused to the fragments.

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Figure 3 Schematic of a BiFC technique combined with a fluorescent timer protein.

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fluorophore resorufin. Their label is initially membrane-permeable and nonfluorescent, and it will acquire fluorescence only upon binding to the CCXXCC motif. Using the combination of FlAsH and ReAsH, Gaietta et al. have deter-mined the mechanism by which connexin 43 (Cx43), a subunit of gap junction channels, is added to and removed from gap junction plaques41. By introducing the CCXXCC motif into the Cx43 protein and then alternately labeling cells with FlAsH or ReAsH, different pools of the protein could be followed over time. Such partitioning between red and green fluorescence has revealed novel characteristics of Cx43 transport, assembly into channels and turnover. This study demonstrates the benefit of the FlAsH/ReAsH technique for analyzing protein aging over any chosen time frame, rather than the fixed time frame characteristic of the green-to-red shifting fluorescent timer protein.

ConclusionsTime is an important parameter in biological systems, with signaling cascades, protein syn-thesis, and cell cycles showing distinct activities during different temporal phases. The devel-opment of fluorescent techniques to record information along the dimension of time has already revealed important insights into bio-logical assembly and gene expression; further technological improvements will undoubtedly advance our understanding of cellular behav-ior. Indeed, future studies using these and other

‘timer’ fluorescent proteins will help reveal spatiotemporal information encoded within biological systems.

ACKNOWLEDGMENTSThe authors thank H. Mizuno and T. Kogure for preparation of figures and D. Mou for critical reading of the manuscript.

COMPETING INTERESTS STATEMENTThe authors declare no competing financial interests.

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