Perspective pubs.acs.org/biochemistry
Chemistry Is Dead. Long Live Chemistry! Luke D. Lavis* Howard Hughes Medical Institute, Janelia Research Campus, 19700 Helix Drive, Ashburn, Virginia 20147, United States ABSTRACT: Chemistry, once king of fluorescence microscopy, was usurped by the field of fluorescent proteins. The increased demands of modern microscopy techniques on the “photon budget” require better and brighter fluorophores, causing a renewed interest in synthetic dyes. Here, we review the recent advances in biochemistry, protein engineering, and organic synthesis that have allowed a triumphant return of chemical fluorophores to modern biological imaging.
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oligonucleotide in cells. To improve the performance of small molecule labels, chemists introduced different functionality, such as sulfonates to increase solubility and rigidification motifs to improve brightness and photostability. This resulted in a panel of excellent and still widely used fluorophores, such as the “Alexa Fluor” dyes (8−11) developed by Molecular Probes6 and “CyDyes” (e.g., Cy5, 12) developed by Alan Waggoner7 (Figure 1C). In addition to these improved biomolecule labels, other synthetic fluorophore reagents were being developed for live-cell microscopy, including stains for specific organelles, fluorescent ion indicators, and photoactivatable dyes for advanced imaging experiments.3,4 Chemistry was king... ...and then everything changed. The discovery of green fluorescent protein (GFP) sparked a revolution in biological imaging.8 Cell biologists were no longer beholden to chemists and their relatively expensive synthetic fluorophores. Easily replicated DNA plasmids replaced vials of exhaustible dyes, and cells proved to be perfectly capable of synthesizing fluorophore fusions on their own. The general utility of fluorescent protein fusions superseded many specific organelle markers, and these labels could be multiplexed with established and emerging small molecule lipid and nucleic acid stains. Subsequent advances in fluorescent proteins have replicated many of the properties once exclusive to small molecules, such as red-shifted spectra, ion sensitivity, and photoactivation.2,8,9 These important advances lead to an obvious question: In this age of GFP and its ilk, is the field of fluorophore chemistry dead? Although fluorescent proteins will remain an essential part of the fluorescence microscopy tool kit, small molecule dyes cannot be ignored. There has been a resurgence in chemical fluorophores for imaging in the past decade, driven largely by advances in fluorescence microscopy techniques, which place increased demands on the “photon budget”. The photon budget equals the number of fluorophores in a given sample
he ability to visualize the structure and dynamics of molecules inside cells is an essential part of unraveling fundamental biological processes. Fluorescence microscopy is uniquely suited for such efforts, given the high sensitivity of fluorescence measurements and its compatibility with live cells, tissues, and animals. The applications of fluorescence microscopy are broad, ranging from tracking of individual protein molecules in single cells to measuring neuronal activity over vast swaths of tissue in vivo.1,2 All of these experiments rely on fluorescent dyes, and a wide variety of fluorophores have emerged as useful labels for microscopy: organic fluorophores, fluorescent proteins, lanthanide chelates, and quantum dots. Here, we review the relatively old field of small molecule fluorescent dyes, which is undergoing a renaissance to meet the needs of 21st century biochemistry and biology.3,4 The story of fluorescence and fluorescence microscopy is one of chemistry. Fluorescence was first observed and elucidated using the fluorescent natural product quinine [1 (Figure 1A)]. Quinine was also the target of William Perkin, who instead accidentally made the first synthetic dye, mauvine (2), in 1856. This discovery of synthetic colored compounds set off a flurry of activity, and the majority of the classic fluorophores were synthesized in the subsequent decades, including coumarins (e.g., umbelliferone, 3, ca. 1884), fluorescein (4, ca. 1871), rhodamines (e.g., tetramethylrhodamine, TMR, 5, ca. 1887), phenoxazines (e.g., resorufin, 6, ca. 1903), and cyanines (e.g., hexamethylindocyanine, 7, ca. 1924) (Figure 1B).4 These classic dyes played an important role in the development of fluorescence-based technologies such as fluorescence microscopy. For example, fluorescein (4) sparked the field of immunofluorescence, where the amine-reactive fluorescein isothiocyanate (FITC) derivative was developed specifically for preparing fluorescent antibody bioconjugates.5 The growth of fluorescence microscopy and the development of other labeling strategies [e.g., fluorescence in situ hybridization (FISH)] demanded more sophisticated biomolecule labels. Classic dyes such as 3−7 have several problems: poor photostability, low brightness, and relatively high hydrophobicity. Attachment of these dyes to biomolecules often had detrimental effects on the binding of an antibody or an © 2017 American Chemical Society
Special Issue: Seeing Into Cells Received: June 1, 2017 Revised: July 10, 2017 Published: July 13, 2017 5165
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Figure 1. Chemical structures of early and optimized fluorophores. (A) The first small molecule fluorophore, quinine (1), and the first synthetic dye, mauvine (2). (B) Classic fluorophores: umbelliferone (3), fluorescein (4), tetramethylrhodamine (5), resorufin (6), and hexamethylindocyanine (7). (C) Examples of optimized Alexa Fluor (8−11) and CyDye (12) fluorophores.
ligand (e.g., FlAsH, 13) and a short genetically encoded tetracysteine (Cys4) peptide tag could be used to label proteins in cells (Figure 2A).13 This idea sparked other strategies for labeling biomolecules in cells, including self-labeling tags such as the SNAP-tag14 and HaloTag15 proteins. These widely used systems consist of a genetically encoded enzyme variant tag that reacts specifically and irreversibly with a small substrate ligand motif attached to a fluorophore such as TMR-HaloTag ligand 14 (Figure 2B). Another approach involves the use of engineered ligases, such as lipoic acid ligase, that catalyze the covalent attachment of a fluorophore ligand such as resorufin derivative 15 to a small peptide tag (Figure 2C).16 Non-natural amino acids such as coumarin 16 can be incorporated into a protein structure using exogenous tRNA and tRNA synthetases to place fluorophores at specific sites (Figure 2D).11 Engineered binding motifs such as the antibody-based fluorogen activating proteins (FAPs) bind and enhance small molecule fluorogens such as Malachite Green derivative 17 (Figure 2E).17 Finally, new cellular stains can be created by attachment of a dye to a molecular species with high affinity for an endogenous molecular target such as paclitaxel−fluorescein conjugate 18 (Figure 2F).18,19 All of these labeling strategies have trade-offs among the size of the genetically encoded tag, the speed and selectivity of the fluorophore attachment, the brightness of the resulting conjugate, and the complexity of the system. The bisarsenical approach utilizes a small tag, but ligands such as FlAsH (13) label slowly and exhibit relatively high background staining in cells, presumably because of reaction with other endogenous cysteine residues. Self-labeling tags exhibit rapid and selective labeling with cell-permeable ligands such as 14 but are typically as large as or larger than GFP, which can alter the biological activity of the fusion protein. Ligases combine a small tag with enzyme-mediated kinetics; however, the system requires separate expression of an exogenous ligase, and the scope of compatible fluorophores is limited to relatively small dyes such as resorufin 15. Non-natural amino acids are perhaps the smallest possible perturbation but require exogenous expression of two biochemical machines and the delivery of the non-
multiplied by the number of photons emitted by each fluorophore before photobleaching. The amount of information one can extract from a biological sample is wholly dependent on the photon budget; pushing the frontiers of fluorescence microscopy often requires more photons. For example, superresolution localization microscopy places a large burden on the photon budget, as the number of photons emitted by each fluorophore determines how precisely individual molecules can be localized.10 Likewise, moving from transient overexpression of protein fusions to gene-edited cells with endogenous expression levels typically decreases the number of fluorophores, also compromising the photon budget. The importance of this parameter in imaging has led to the development of new microscope geometries expressly designed to capture more photons.3 In addition to optical physics methods, chemistry can be used to improve the photon budget. Chemical fluorophores can be substantially brighter and more photostable than fluorescent proteins, providing a straightforward way to increase the number of photons emitted by a sample. Of course, “regressing” to small molecule dyes seems to be unpleasant at best; classic methods for introducing otherwise cell-impermanent fluorescent conjugates in live cells involve tedious microinjection, electroporation, or bead loading. Fortunately, over the past 20 years, clever chemists and biochemists have developed techniques for making the labeling chemistry easier and more functional in complex biological environments such as live cells and tissues (Figure 2). These flexible strategies allow the excellent photophysics of chemical dyes to be combined with the genetic specificity of fluorescent proteins. The majority of in-cell labeling strategies have two parts: (i) a genetically encoded “tag” expressed as a fusion with your favorite protein and (ii) a synthetic fluorophore-containing “ligand” that binds to the tag.11,12 Instead of introducing an entire fluorescent bioconjugate into cells, one is faced with only getting a small molecule across the lipid bilayer. As with most useful ideas in biological imaging, the initial breakthrough for in-cell labeling techniques was provided by Roger Tsien, who showed that the selective interaction between a bisarsenical dye 5166
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Figure 2. Cellular labeling strategies using small molecule fluorophores. (A) Labeling of tetracysteine-containing peptides with bisarsenical 13. (B) Attachment of HaloTag ligand 14 to a self-labeling HaloTag fusion protein. (C) Amidation of the lipoic acid ligase tag with ligand 15 catalyzed by engineered lipoic acid ligase variant. (D) Use of non-natural amino acid technology to incorporate coumarin amino acid 16 into proteins. (E) Antibody-based fluorogen activating proteins (FAPs) bind Malachite Green derivatives such as 17. (F) Cellluar stains such as paclitaxel 18 bind endogenous proteins via a “GMO-free” strategy. Light gray denotes the protein of interest and dark gray the exogenous tag.
biorthogonal labeling by a larger fluorophore moiety.11,12 In the FAP system, the binding of the ligands is fluorogenic and reversible; this can improve photobleaching through fluorophore exchange but also decreases the overall brightness.17 Finally, development of stains is attractive as this approach
natural amino acid. In addition, only relatively small fluorophores such as coumarin 16 are compatible with the ribosome, which further limits the utility of this approach. One strategy for circumventing this problem is to incorporate a small “click chemistry” handle into the protein for subsequent 5167
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Figure 3. Modern chemistry yields new dye derivatives. (A) Addition of metalated aryl species to xanthone analogues yields a variety of rhodamine derivatives. (B) The “open−closed” equilibrium of carborhodamines and Si-rhodamines is shifted toward the closed lactone form. (C) Reagents based on carborhodamines and Si-rhodamines: SiR700-SNAP-tag ligand (21), spontaneous blinking HM-SiR (22), and fluorogenic carborhodamine HaloTag ligand 23. (D) Use of Pd-catalyzed cross-coupling to synthesize known (TMR, 5) and novel (Janelia Fluor 549, 25) rhodamines from a simple fluorescein derivative (24). (E) Replacing N,N-dialkyl substituents in classic dyes yields a panel of bright, photostable “Janelia Fluor” dyes (25−31) and allows fine-tuning of spectral and chemical properties.
and oligonucleotide labels for fixed-cell imaging, not for live-cell applications. The polar functionality and relatively high molecular weight of these dyes (Figure 1C) complicate their use with these new labeling strategies due to poor cell membrane permeability and/or incompatibility with enzymemediated labeling. For this reason, many of the existing small molecule labeling techniques have focused on classic, small, and net-neutral dyes that date back a century, as demonstrated in example labels 13−18 (Figure 2). The development of these new labeling strategies and their use in advanced microcopy experiments have necessitated revisiting and reinventing fluorophore chemistry.4 Although the focus on synthetic dyes in the 19th century yielded useful and general synthetic approaches, these methods utilized reagents and techniques from the earliest era of chemistry, predating canonical chemistry concepts such as organometallic reagents and transition metal catalysis. To expand the scope of dye synthesis, chemists have been applying modern methods to classic fluorophores. For example, instead of the century-old acid-catalyzed chemistry to synthesize rhodamines, the addition of aryl-Grignard or aryl-lithium species to xanthone-type derivatives is an alternative method for preparing rhodamines and red-shifted analogues where the xanthene oxygen is replaced with a gem-dimethyl carbon,21,22 gem-dimethyl silicon,23,24 phosphinate,25 or sulfone26 group (Figure 3A). In particular, the far-red Si-substituted rhodamine scaffold has emerged as a bright and photostable framework that can substantially increase the size of the photon budget in imaging experiments. The synthesis of these analogues would be
dispenses with the requirement of a genetically encoded tag and can label endogenous proteins. Nevertheless, this system is not as generalizable as other methods, requiring a unique small molecule binding element for each target protein.19,20 Currently, self-labeling tag systems are perhaps the best method for live-cell labeling and the easiest switch from fluorescent proteins given the relative simplicity of the system, the generality of the labeling reaction with diverse chemical functionality, and the availability of fluorescent and fluorogenic ligands. Of these, the HaloTag system reigns supreme, based on its rapid labeling kinetics (k2 ≈ 107 M−1 s−1) and the high cell permeability of the ligand moiety, which allows labeling of live cells with low concentrations of the fluorophore (∼100 nM) and facilitates use in tissue and in vivo. Nevertheless, the larger size of the HaloTag (∼33 kDa) can be problematic with some protein fusions. The SNAP-tag is smaller (∼20 kDa) but labels slower (k2 ≈ 104 M−1 s−1), and the benzyl guanine ligand motif is less cell-permeable than the chloroalkane HaloTag ligand. Thus, the SNAP-tag system requires higher ligand concentrations (∼1 μM), which necessitates more rigorous washing protocols. Continued optimization and benchmarking of these systems and exploration of other protein-based self-labeling tags (e.g., dihydrofolate reductase, β-lactamase, and photoactive yellow protein) are needed to enable multicolor imaging in live cells using synthetic fluorophores. Given the advances in both dye and protein engineering, it would seem obvious to combine the commercial panels of advanced fluorophores (Figure 1C) with these improved livecell labeling technologies (Figure 2). However, the majority of optimized commercial fluorophores were designed for antibody 5168
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resolution in fluorescence microcopy demands increasingly large photon budgets. Innovative labeling strategies and improved fluorophores are making chemical dyes progressively attractive and accessible to cell biologists, particularly for imaging in the far-red region or for super-resolution microscopy. The continued development of fluorescent and fluorogenic stains (Figure 2F) circumvents the need for transfection, allowing “GMO-free” imaging in live cells. This triumphant return of chemical dyes is driven by chemistry; the application of modern synthetic techniques will allow further structural refinements to yield dyes with better brightness, higher fluorogenicity, improved photoactivation, and enhanced bioavailability. We look forward to the next era of imaging in which the frontiers of microscopy expand under the reign of both fluorescent proteins and synthetic dyes.
difficult or impossible using the conventional xanthene dye syntheses that rely on acid-catalyzed condensation. In addition to eliciting a bathochromic shift, these substitutions also affect chemical properties such as the equilibrium between the “open”, fluorescent, zwitterionic form and the “closed”, nonfluorescent, lactone form. Both the carbon-containing analogues (e.g., “carboTMR”, 19) and the silicon congeners (e.g., SiTMR or “SiR”, 20) are substantially shifted toward the closed form (Figure 3B). For SiTMR (20), the altered open−closed equilibrium ensures the free dye primarily adopts the colorless lactone form in solution, but the equilibrium can shift to the open form upon an environmental change such as binding to a polar protein surface. Discovered by Johnsson and co-workers, this property of SiTMR has allowed the synthesis of fluorogenic ligands for live-cell labeling, including self-labeling tags,27 stains for endogenous cytoskeletal proteins,19 and a fluorogenic DNA binder.20 This property appears to be general to Si-rhodamines and can be exploited to make fluorogenic reagents with farther red-shifted spectra [e.g., SiR700-SNAP-tag ligand 21; λmax ≈ 700 nm (Figure 3C)]28 or “spontaneous blinking” dyes such as hydroxymethyl-Si-rhodamine (HM-SiR, 22) that allow super-resolution localization microscopy without photoactivation;29 these are particularly useful for imaging intracellular membranes.30 Finally, carborhodamines can also be made into fluorogenic ligands by shifting the open−closed equilibrium through subtle structural modifications. A HaloTag ligand based on 2′,7′-difluorocarboTMR (23) shows a 9-fold increase in fluorescence upon binding to the HaloTag protein.31 Another modern chemical reaction that can be applied to fluorophores is Pd-catalyzed cross-coupling chemistry. Instead of the synthesis of TMR (5) using the traditional acid-catalyzed condensation chemistry, the dye can be prepared by Buchwald−Hartwig cross-coupling of dimethylamine with fluorescein bistriflate (24) (Figure 3D). This application of contemporary chemistry not only allows easier access to known structures with established biological utility, such as 5, but also enables exploration of new molecules with improved properties. The use of Pd-catalyzed cross-coupling chemistry allowed our laboratory to introduce a new auxochrome group, the fourmembered azetidine ring, which would not survive the classic acid-catalyzed condensation conditions (Figure 3D). This simple modification elicited substantial improvements in the brightness and photostability of TMR (5) with the azetidinyl analogue (25, “Janelia Fluor 549”) emitting 4-fold the number of photons under the same imaging conditions in live cells, thereby substantially increasing the size of the photon budget of microscopy experiments such as single-particle tracking.24 This strategy is general, allowing the design and synthesis of Janelia Fluor (JF) dyes that span the visible spectrum [25−31 (Figure 3E)] and exhibit properties superior to those of the N,Ndimethylamino-substituted parent dyes.24,32 The excellent brightness of these fluorophores makes them useful scaffolds for self-labeling tags or stains for cellular imaging and can be extended to more complex biological environments such as intact tissue or animals.32,33
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. ORCID
Luke D. Lavis: 0000-0002-0789-6343 Notes
The author declares the following competing financial interest(s): L.D.L. has filed patent applications on the Janelia Fluor dyes, whose value might be affected by this publication.
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ACKNOWLEDGMENTS An earlier version of this piece was published as a blog post on addgene.org. I thank Tyler Ford (addgene), Brett Mensh (Janelia), and Jonathan Grimm (Janelia) for helpful comments. Related work in our laboratory is supported by the Howard Hughes Medical Institute.
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REFERENCES
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CONCLUSION Biologists now have a wealth of options for visualizing and tracking molecules inside cells. The ease of use and continuing optimization of fluorescent proteins make these labels the go-to choice for the majority of fluorescence microscopy experiments. Nevertheless, the quest for higher spatial and temporal 5169
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