Small-Molecule Fluorescent Probes for Live-Cell Super-Resolution

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Small-Molecule Fluorescent Probes for Live-Cell Super-Resolution Microscopy Lu Wang, Michelle S. Frei, Aleksandar Salim, and Kai Johnsson J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.8b11134 • Publication Date (Web): 14 Dec 2018 Downloaded from http://pubs.acs.org on December 15, 2018

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Journal of the American Chemical Society

Small-Molecule Fluorescent Probes for Live-Cell Super-Resolution Microscopy Lu Wang1, Michelle S. Frei1-2, Aleksandar Salim1-2, Kai Johnsson1-2* 1

Department of Chemical Biology, Max Planck Institute for Medical Research, Jahnstrasse 29, 69120 Heidelberg, Germany

Institute of Chemical Sciences and Engineering (ISIC), École Polytechnique Fédérale de Lausanne (EPFL), 1015 Lausanne, Switzerland 2

KEYWORDS: Super-resolution microscopy, Fluorogenic probes, Live-cell imaging, Protein labeling, Bioorthogonal chemistry

ABSTRACT: Super-resolution fluorescence microscopy is a powerful tool to visualize biomolecules and cellular structures at the nanometer scale. Employing these techniques in living cells has opened up the possibility to study dynamic processes with unprecedented spatial and temporal resolution. Different physical approaches to super-resolution microscopy have been introduced over the last years. A bottleneck to apply these approaches for live-cell imaging has become the availability of appropriate fluorescent probes that can be specifically attached to biomolecules. In this perspective, we discuss the role of small-molecule fluorescent probes for live-cell super-resolution microscopy and the challenges that need to be overcome for their generation. Recent trends in the development of labeling strategies are reviewed together with the required chemical and spectroscopic properties of the probes. Finally, selected examples of the use of small-molecule fluorescent probes in live-cell super-resolution microscopy are given.

Introduction Characterizing cellular structures and biomolecules with high temporal resolution in living cells is a prerequisite for a molecular understanding of biology. So far, fluorescence microscopy has been the method of choice for this task as it offers good sensitivity and temporal resolution. However, until the introduction of super-resolution microscopy (SRM), the spatial resolution of conventional fluorescence microscopy of about 200 nm was not sufficient to resolve most biological structures. Different SRM techniques have emerged in the past two decades as unique tools to visualize cellular structures with nanoscale resolution.1-3 The common feature of the different approaches is to switch fluorophores between a fluorescent ‘on’ and a dark ‘off’ state, allowing to collect light from only a subset of precisely localized ‘on’ fluorophores.4 Different strategies to achieve the ‘on’-‘off’ switching exist. Switching can be induced at defined coordinates with sub-diffraction resolution as it is done in Stimulated Emission Depletion (STED) microscopy.5 In Single Molecule Localization Microscopy (SMLM) switching is a stochastic process and the emission of fluorophores within a diffraction-limited area are separated in time.6-9 These methods have been demonstrated in many cellular contexts and have led to important insights into the cellular organization at the nanoscale.1-4, 10-13 However, up to now many of these studies were only performed in fixed cells, which do not allow for observation of cellular processes in real time. Furthermore, fixation procedures come with the risk of introducing artefacts and destroying the ultrastructure of samples, which becomes exceedingly relevant at higher

resolutions.14 Live-cell SRM, in contrast, avoids the pitfalls of fixation procedures and offers insights into the dynamics of biological systems at the nanoscale. The main bottleneck in live-cell SRM has become a chemical problem: the specific attachment of a suitable fluorophore to the biomolecule of interest.15 Many types of fluorophores, including small-molecule fluorophores,16-17 fluorescent proteins,18-19 and nanoparticles,20 have been developed for SRM. However, stringent requirements apply for their use in live-cell SRM. These include photophysical criteria such as high brightness, photostability and switching kinetics. In addition, they need to be targetable and, in the case of synthetic fluorophores, be membrane-permeable in order to specifically stain cellular structures such as proteins or organelles. Small-molecule, synthetic fluorophores possess several features that make them ideally suited for live-cell SRM. First of all, they are smaller in size than both fluorescent proteins and nanoparticles, a feature that will become more and more relevant as the resolution of SRM is pushed towards a molecular level. Second, synthetic fluorophores can possess outstanding brightness and photostability, such that the resulting photon budget can be one order of magnitude higher than that of fluorescent proteins.15, 21 Often these properties can be rationally tuned using synthetic chemistry. Thirdly, their specific attachment to intracellular proteins is a more tractable problem than for nanoparticles.15-17 Examples of smallmolecule fluorophores used in live-cell SRM are shown in Figure 1.

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Figure 1. Examples of small-molecule fluorophores used in live-cell SRM. Absorption and emission maxima are given underneath the structures (abs/em). In parentheses the application in which these fluorophores were used is listed. R depicts the position where the fluorophores are substituted for labeling purposes. Reference: 515R (1)22, TMR (2)23-24, 610CP (3)22, 25, GeR (4)26-27, 680SiR (5)25, SiR700 (6)28, PicoGreen (7)29, PA-JF549 (8)30 LysoTracker Red (9)31 SiR (10)32 HMSiR (11)33, ATTO655 (12)34.

In recent years, various reviews have been published on different aspects of SRM such as different super-resolution techniques, biological applications or specific aspects of probe design.1, 4, 10-11, 15, 18, 21, 35 Thus, in this perspective we will mainly focus on probe design and labeling strategies for live-cell SRM. As stated, the development of fluorescent probes for live-cell imaging requires the optimization of very different parameters such as membrane permeability, solubility, emission wavelengths, brightness, photostability and switching kinetics. These properties cannot be optimized independently but influence each other, making probe development an iterative process. Therefore, we will first review how fluorescent probes can be specifically attached to biomolecules in living cells before discussing how the chemical and photophysical properties of small-molecule fluorophores can be optimized for live-cell SRM.

Labeling Specific labeling of biomolecules, organelles or membranes with small-molecule fluorophores in living cells represents a formidable challenge. Whereas fixed cells are conventionally stained by antibody conjugates and fluorescent proteins can be genetically encoded in engineered cells, more elaborate strategies have to be used for live-cell labeling with synthetic fluorescent probes (Figure 2). In the following, we will limit our discussion to those labeling approaches that have been shown to be applicable for labeling with fluorophores suitable for SRM and that do not require invasive methods such as microinjection

or electroporation. The most commonly used approach for livecell protein labeling is based on the expression of the protein of interest as a fusion protein with an additional polypeptide or protein (a so-called tag) that undergoes a specific, bioorthogonal reaction with a synthetic fluorescent probe.2, 17, 36-37 The first tag that possessed the required specificity, speed of labeling and could be labeled with a large variety of different fluorescent probes in different cellular compartments was SNAP-tag (Figure 2A, E).38 SNAP-tag undergoes an irreversible reaction with benzylguanine derivatives 13 in which the benzyl group, together with a label, such as a fluorophore, is transferred to an active-site cysteine residue. Since then a number of other protein tags for covalent labeling have been introduced, most notably HaloTag39 and CLIP-tag.40 In addition, tags which bind non-covalently to fluorescent ligands or which are modified through the action of specific ligases can be employed.41-42 Thus, fluorophores conjugated to the substrates of these tags can be used to label and observe proteins in live-cell SRM (Figure 2A, E). One inherent disadvantage of the approach is that it is based on fusing a tag to the protein of interest. SNAP-tag (20 kD) and HaloTag (33 kD) have diameters of about 3-4 nm and can therefore affect the native function of proteins. Care must be taken to ensure proper localization and ideally, the fusion protein should be able to rescue the defect of loss of endogenous protein. Moreover, changes in expression levels due to overexpression might result in artefacts such as mislocalization.43 A solution to the latter


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Figure 2. Overview of approaches suitable for live-cell labeling of biomolecules with synthetic fluorophores. (A) Fusion of proteins with self-labeling protein tags such as SNAP-tag, Clip-tag or HaloTag. (B) Exploiting click chemistry to site-selectively label proteins or lipids. (C) Labeling through ligands that specifically bind to biomolecules. (D) Labeling of organelles such as mitochondria. (E) Structures of ligands used in the labeling strategies (A-D). 13 SNAP-tag ligand benzylguanine. 14 strained alkyne and tetrazine 15 used in click chemistry. 16 F-actin-binding ligand jasplakinolide. 17 triphenylphosphine used to stain the mitochondrial membrane. R in 14 depicts the ligand needed for metabolic incorporation or an unnatural amino acid used.

problem is the use of genome editing to generate heterozygous or homozygous knock-in cell lines in which fusion proteins are expressed under their endogenous promotor. The development of the CRISPR/Cas9 system, a genome editing tool, permits to generate knock-in cell lines in a faster, cheaper and more accurate way than before. This provides a solution to overexpression artefact but still does not address the question of functionality of fusion proteins.44 Regardless, the approach always requires genetic engineering of the target cells or organism. Another limitation is that the position of the tags, and therefore the fluorophore, is generally limited to the N- or Ctermini of proteins. Inserting self-labeling protein tags into loops is possible but requires identification of a suitable insertion site. The use of unnatural amino acid incorporation into proteins is an elegant solution to this problem.45-46 Here, an unnatural amino acid encoding a biorthogonal handle such as a strained alkyne 14 is incorporated at the desired position in the protein of interest using amber codon suppression technology. Subsequently, the protein carrying the unnatural amino acid is fluorescence labeled with a suitable reaction partner such as a tetrazine derivative 15 (Figure 2B, E). The approach has the advantage that it gives freedom over the position at which the fluorophore is introduced and that no tag is added to the protein. However, the process of unnatural amino acid incorporation and the subsequent fluorescent labeling are technically challenging. In addition, recognition of the amber stop codon by release factor 1 can result in decreased levels of labeled protein.45 Until now, the approach has not yet become a routine method for live-cell imaging.45-46

The ideal labeling method for live-cell SRM would be direct labeling of the protein of interest with a fluorescent probe. This can be achieved by conjugating fluorophores to ligands that bind to proteins with high affinity and specificity (Figure 2C), provided that such ligands are available. A good example is the natural product jasplakinolide 16, which binds with high specificity to F-actin and has been used to generate powerful probes for live-cell imaging (Figure 2E).47-48 Ligands can be natural products such as jasplakinolide but also synthetic drugs, provided that they can be derivatized without losing the affinity for their target. Numerous small-molecule fluorescent probes based on natural products or synthetic drugs have been developed in the last years.48 The main limitation of this approach is that binding to their protein target usually affects the function of the protein; for example, jasplakinolide stabilizes F-actin. This is a potential pitfall that has to be kept in mind when performing live-cell SRM with such probes.48 The majority of live-cell SRM experiments up to date have been performed with labeled proteins. This can be explained by the key role that proteins play in almost all biological processes but also by the fact that live-cell labeling of proteins is a more tractable problem than that of other biomolecules as reactive handles can be genetically encoded into proteins. Labeling of lipids and nucleic acids for live-cell SRM can be achieved by metabolic incorporation of biorthogonal reactive groups and subsequent labeling through biorthogonal chemistry. Applying the ligand and the fluorophore successively in a two-step labeling process can solve cell-permeability issues of some fluorophore-ligand conjugates (see below).49 Furthermore, few



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Figure 3. Uptake of fluorophores into cells. The three main pathways of how small-molecule fluorophores can enter cells are depicted. In addition, the spirolactone equilibrium of rhodamines is depicted, using a SiR-carboxyl-based probe as a representative example. Note that the uncharged spirolactone 18 enters the cell through passive transport and that binding of the probe to its target shifts the equilibrium to the fluorescent zwitterion 10.

small-molecule fluorophores can stain organelles such as lysosomes due to their unique chemical properties.31, 50 Finally, several chemical groups have been developed to lead to accumulation of fluorophores in specific organelles or to target them to specific organellar membranes and DNA (Figure 2D, E).29, 31, 51-53

Permeability and solubility A common bottleneck of all labeling approaches discussed above is the difficulty in getting the fluorescent probe into cells, in particular when excluding more invasive techniques such as permeabilization,23 bead loading,54 microinjection55 and cellsqueezing.56 Passive diffusion along the concentration gradient is thought to be the main mechanism of cell permeation (Figure 3).57 While active transport may act simultaneously, efflux pumps can also work against the desired outcome. A third way of entering the cell is through endocytotic pathways. However, to reach the intracellular space efficient endosomal release is required.57 Passive diffusion is also the main mechanism of uptake of drugs targeting intracellular targets and the rules established by medicinal chemists for drug design over the last decades also apply to probes. Lipinski’s Rule of five summarizes several important aspects of drug design. Ideal drug candidates usually show no more than five hydrogen bond donors and no more than ten hydrogen bond acceptors. They have a molecular mass lower than 500 and the octanol-water partition coefficient (logP) is smaller than 5.58 Similarly, to obtain membrane permeable probes, polarity (hydrogen bond donors and acceptors) and lipophilicity (logP) have to be balanced. However, many fluorophores have been optimized only for brightness, photostability and quantum yield and consequently only a subset of these are also cell-permeable. These include coumarins, BODIPYS as well as the classical

rhodamines and their carbo, germanium and silicon rhodamine derivatives (Figure 1).2 The key to the relative good permeability of rhodamines is their capacity to reversibly form a spirolactone, which keeps the fluorophore in a dynamic equilibrium between a fluorescent, zwitterionic and a nonfluorescent, uncharged form (Figure 3). It is the latter form, which can more readily cross membranes.32, 59 Similarly, chemical fixation of the spirolactone form in fluoresceins and rhodamines was used to create cell-permeable fluorescent probes, provided that the chemical fixation could be reversed in cells.60-61 The importance of the reversible spirolactone formation of rhodamines for probe design became apparent with the introduction of SiR-carboxyl (Figure 3), which shows an increased propensity to form the spirolactone. Furthermore, by controlling the equilibrium of spirolactonization of rhodamines through careful tuning of the electron density of the xanthene ring, the approach was extended to a large palette of different fluorophores.62 Together, this has permitted the generation of numerous probes for live-cell SRM, all based on the rhodamine class of fluorophores.28, 30, 32, 48, 53, 63-64 The dynamic equilibrium between a zwitterion and an uncharged, non-fluorescent spirolactone also aides in reducing unspecific staining. Generally, probes that are too hydrophobic can cause solubility issues and can lead to unspecific staining of cellular organelles, thereby increasing the background in SRM.11 If the spirolactone form of a probe is responsible for such unspecific binding or aggregation, this will not result in increased fluorescence background signal. The flip side of the existence of a dynamic equilibrium between a fluorescent and non-fluorescent form is that it risks to reduce the overall brightness of a fluorescent probe. How this apparent disadvantage can be exploited to dramatically increase the


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performance of fluorescent probes in live-cell SRM is discussed in the next paragraph.



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Figure 4. (A) Chemical structures and (B) photophysical properties of selected small-molecule fluorophores. R groups show where the fluorophores are substituted for labeling purposes. a 10 mM HEPES buffer, pH 7.3. b 10 mM PBS pH 7.4. c Ethanol containing 0.1% (v/v) trifluoroacetic acid. d 10% methanol/PBS (10 mM, pH 7.4). e 10% acetonitrile/H2O.

Fluorogenicity Fluorogenicity in the context of imaging refers to the ability of a fluorophore to increase fluorescence upon interaction with its target. The importance for live-cell SRM is that fluorogenicity can dramatically reduce background fluorescence resulting from unspecific localization of probes. Even though various design strategies for the generation of fluorogenic probes exist,65 up to now mainly two approaches have been used to generate fluorogenic probes for SRM. In the first, target binding shifts the dynamic equilibrium between a fluorescent and a nonfluorescent form of small-molecule fluorophores to the fluorescent form. In the second, a quencher is released upon target binding. The first concept can be illustrated with the already introduced SiR-carboxyl. As discussed, SiR-carboxylbased probes exist in an equilibrium between a fluorescent 10 and non-fluorescent form 18 (Figure 3). The equilibrium is such that the non-fluorescent form 18 is predominant in aqueous environment. In addition, certain SiR-probes are also thought to form non-fluorescent aggregates, which further decreases the background signal.27 Upon binding of probe to its protein target the equilibrium is shifted to the zwitterionic fluorescent form 10. The magnitude of the fluorescence increase can be larger than 100-fold, as shown for the F-actin probe SiR-actin, which was obtained by fusing SiR-carboxyl to jasplakinolide 16. This degree of fluorogenicity even allows imaging experiments without removing excess probes through washing steps, which greatly facilitates live-cell SRM.48 Favoring of the zwitterionic form 10 over the spirolactone 18 upon protein binding of the probe is believed to be due to interactions of the probe with the polar surface of proteins and breaking up of aggregates.32 While the underlying mechanism of fluorogenicity will depend on different factors and their relative contributions will also differ for different probes, the approach has been applicable for a large number of probes for live-cell SRM, including probes for SNAP-tag and HaloTag,32 microtubules,48 F-actin,48 DNA,53 lysosomes,28 plasma membrane,63 endoplasmic reticulum

(ER),63 Golgi,63 and BACE1.64 Furthermore, the effect was exploited in probes based on rhodamines, carbopyronines, germanorhodamines and rhodols.22, 26, 62, 66 It should be noted that binding of a probe to its target can also shift the equilibrium to the non-fluorescent spirolactone, for example when the fluorescent probe is targeted to membranes. This apparent disadvantage has been ingeniously exploited in the creation of so-called high-density environmentally sensitive (HIDE) probes. HIDE probes accumulate predominantly in the dark state in membranes and can serve as a dark reservoir for-long term live-cell SRM.63 Another approach for the generation of fluorogenic probes is to conjugate quenchers, such as tetrazine 15, to cell-permeable fluorophores. Besides the ability to bio-orthogonally react with strained alkenes and alkynes, tetrazines can quench fluorophores via either long-range dipole−dipole interaction or through-bond energy transfer.67-68 Click reaction with the corresponding partners then removes the quenching effect of tetrazines. This approach has so far mostly been employed in fixed cell SRM67-68 or the quenching effect was not investigated in more depth.49, 69 Considering the potential of this approach for live-cell SRM, in particular in conjunction with unnatural amino acid technology, more research on the development of fluorogenic probes based on biorthogonal chemistry is warranted.

Absorption and Emission Wavelength When it comes to choosing the spectroscopic properties of fluorophores for live-cell imaging, absorption and emission wavelengths in the far-red to near infrared window (650 - 1350 nm) are advantageous due to reduced absorbance and autofluorescence of cells at this wavelength, and because of decreased phototoxicity of light at higher wavelengths.70 Nevertheless, the availability of fluorophores in different spectral ranges is needed to allow for multicolor imaging.23, 71 Tuning of the absorption and emission wavelengths of smallmolecule fluorophores through structural modification is aided


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by the large body of literature on this topic.61 Considering the predominance of rhodamine derivatives in live-cell SRM, we will limit our discussions on how absorption and emission wavelengths can be tuned in this class of fluorophores. The colors of other fluorophore classes such as cyanines can be rationally tuned as well, but often these molecules are not cellpermeable.72 In rhodamines, replacing the oxygen atom in the xanthene ring with other atoms significantly affects the spectral properties: Going from oxygen (TMR 19a, 548/572),24 to carbon (610CP 19b, 606/626),22, 66 to germanium (GeR 19c, 634/655)26 and finally silicon (SiR 19d, 643/662)32 shows how a single substitution can lead to a significant bathochromic shift (Figure 4A, B).70 Smaller modifications of the aromatic core or the nitrogen substituents are used to further fine-tune the absorption and emission wavelength (Figure 1).22, 28, 62 Moreover, small-molecule fluorophores with large stokes shifts are useful in multicolor STED microscopy as they allow the use of the same wavelength for depletion for several fluorophores while maintaining two or three different excitation windows to distinguish them.52 Several fluorophores have been modified to obtain large stokes shifts.73 For example, 9iminoanthrone 20 bearing different groups in the 10-position (X = CMe2, GeMe2, SiMe2, and SO2), (Figure 4A, B) display stokes shifts up to 165 nm and were successfully used in live-cell multicolor STED microscopy. For instance 20a was used in combination with 19b and 19c.52

concentrations (μM) through their anti-oxidative properties most TSQs have to be applied in rather high millimolar concentrations (mM) to induce the desired increase in photostability.75 This limits the utility of the approach for livecell imaging. To solve this problem, fluorophores were covalently conjugated to triplet state quenchers, e.g. pnitrobenzyl alcohols (NBA) 23, cyclooctatetraene (COT) 24, or trolox 25.76-79 It was demonstrated that intramolecular triplet state quenching enables fluorophores, e.g. rhodamines, carbopyronines, SiRs and cyanines (Figure 5A), to be resistant to photobleaching in fixed-cell SRM. 76, 79-80 Two sulfonated cyanines conjugated to COT were coupled to the extracellular side of a membrane receptor and displayed high photostability in live-cell imaging.79 However, these fluorophores have not been applied to live-cell SRM of intracellular targets so far. Another strategy to enhance photostability is the fusion of electron-withdrawing substituents to small-molecule fluorophores.22, 81 The recently reported PhoxBright 430 fluorophore 26 includes electron-withdrawing phosphine oxide

Brightness Brightness is the fluorescence output per molecule, which is proportional to the product of the extinction coefficient (at the relevant excitation wavelength) and the fluorescence quantum yield. In live-cell SRM, fluorophores with higher brightness are advantageous as they allow lower laser intensities to be used, thereby reducing phototoxicity. In addition, brighter fluorophores result in more precise localizations in SMLM and therefore in higher resolutions.21 Thus, researchers have proposed different approaches to improve the brightness of fluorophores.61, 72 In general, the design strategy is to suppress non-radiative pathways. For example, replacement of N,Ndimethylamino substituents in TMR 19a by four-membered azetidines (compound 21a) mitigates formation of a twisted internal charge transfer state, which relaxes non-radiatively.24 The substitution resulted in a 2.7-fold increase in brightness and also an increased photostability, while other photophysical properties and cell-permeability were not significantly affected. Importantly, the favorable effect of replacing alkylated amines with azetidines is not limited to rhodamines but was established as a general strategy to increase the brightness of other fluorophores, including coumarines and SiRs.24

Photostability Photostability describes the property of a fluorophore to give a stable fluorescent signal over time and to withstand light irradiation without bleaching. Many small-molecule fluorophores, e.g. rhodamines, coumarins, and cyanines, show acceptable photostability in conventional microscopy. However, the high power densities used in certain SRM techniques can easily cause photobleaching. To avoid photobleaching, researchers have developed several different approaches.35, 74 One of these approaches is the use of triplet state quenchers (TSQ) as additives in fixed cell imaging. Whereas some TSQ can increase the fluorescence output already at micromolar

Figure 5. (A) Representative structure of TSQ-conjugated Cy5 derivative 22 along with the structures of TSQs. (B) Phosphine oxide-fused photostable fluorophore 26. X in 26 represents chemical groups increasing hydrophilicity. (C) Fluorinated photostable rhodamine 27. R groups show where the fluorophores are substituted for labeling purposes.

as part of the fluorophore, resulting in excellent photostability and allowing long-term STED imaging.82 Albeit, the relatively low brightness and low solubility of the fluorophore limits its utility for live-cell imaging. Introduction of CH2CF3 at the nitrogen atom or fluorine atoms at positions 2’ and 7’ of the xanthene in rhodamines (structure 27, Figure 5C) can slow down radical formation and photobleaching.81 As probes based on fluorophore 27 are cell-permeable, they were successfully applied to live-cell STED imaging of tubulin filaments.27 Furthermore, the increased photostability of rhodamines comprising azetidines allowed their successful use in live-cell single molecule tracking, underscoring the utility of azetidines for increasing the performance of small-molecule based fluorescence probes for live-cell imaging (Figure 4A).24, 30 It is worth noting that the requirement for photostability in SMLM is different from that required for STED. STED microscopy requires fluorophores that are very resistant to



photobleaching. However, such fluorophores could be too stable to be switched to an ‘off’ state in SMLM. Several SMLM dyes use mechanisms associated with photobleaching to transition to ‘off’ states, which is further explained in the next section.11, 17

Switching Switching small-molecule fluorophores between an ‘on’ (fluorescent) and an ‘off’ (dark) state is a prerequisite for their use in SRM. In STED microscopy this is achieved by the use of a depletion laser. In SMLM, stochastic switching/blinking of fluorescent probes is commonly based on pushing molecules to a dark state or photoactivation.11, 17 Pushing molecules to a dark state for subsequent stochastic blinking can be achieved by changing the transition probability to triplet states and fluorophore radicals, generating reduced/oxidized forms of fluorophores, and switching between ground state fluorophore isomers.16, 35, 74 Most of these approaches rely on using redox additives and/or oxygen depletion systems that are often not compatible for use with live-cells.17 In addition, high intensity laser irradiation is used to transition fluorophores from bright to dark states, which can induce phototoxicity when applied to live-cells. Furthermore, the best performing small-molecule fluorophores used in SMLM are often not cell-permeable.23 However, scientists have used various approaches to circumvent these limitations. Here we discuss approaches, which were used to obtain suitable blinking kinetics in livecells. Electroporation was used to permeabilize cells and label SNAP-tagged proteins with the non-permeable small-molecule fluorophore Alexa647 29 (Figure 6A).23 To achieve desired blinking properties of the cyanine fluorophore, thiols and an oxygen scavenger system has been used. Albeit, these methods can only be used to observe structures and processes that are not affected by additives and cell permeabilization. Some fluorophores such as ATTO655 31 (Figure 6B) can blink in the presence of glutathione and oxygen as redox system in the cellular environment. The photoswitching cycle is divided in two processes: reduction of the excited fluorophore by glutathione to its non-fluorescent leuco form 30. The leuco form is then oxidized by oxygen which recovers the fluorescent form 31 and completes the photoswitching cycle.17, 83 An important advance in the field was the design of fluorescent probes for live-cell SMLM that can undergo spontaneous blinking in the absence of any additives and high intensity light irradiation.33 Replacing the carboxylate in SiR-carboxyl with a hydroxymethyl group generated a fluorophore that predominantly exists in a non-fluorescent spirocyclic form (HMSiR). However, the molecule can spontaneously switch between its dark form 32 and the bright, open form 33 (Figure 6C). This allows its imaging over extended periods of time at minimal light intensities. Besides stochastically blinking fluorophores, switching can be based on the photoactivation of caged fluorophores. Various irreversibly photoactivatable probes were developed and used in SMLM, but only few of these proved to be compatible with live-cell imaging.84 ortho-Nitrobenzyl and its derivatives are some of the most widely used caging groups. However, fluorophores caged with ortho-nitrobenzyl groups showed low cell-permeability and solubility.85 Consequently, such caged probes where mostly used in fixed-cell SRM.86-88 An attractive alternative is the introduction of a diazoketone group, a very

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small caging group, to rhodamines or SiRs (Figure 6D).30, 89 The diazoketone group locks the fluorophore in a non-fluorescent, spirocyclic form 34, but can be uncaged upon irradiation with light in a two-step process. The initial Wolff-rearrangement gives 35, which efficiently undergoes photoinduced decarboxylation.30

Applications of small-molecule fluorophores in live-cell SRM In the following we highlight some of the applications that demonstrate the potential of small-molecule fluorescent probes for live-cell SRM. The photostable, far-red fluorogenic SiR probes have become particularly popular for live-cell STED imaging. They can be easily combined with the commercially available 775 nm depletion laser, which reduces phototoxicity compared to lower wavelength lasers.10 SiR-tubulin is a fusion of docetaxel to SiR-carboxyl and permits the specific staining of microtubules in living cells. The apparent microtubule diameter measured with SiR-tubulin in U2OS cells was 40 nm. The resolution obtained was two times higher than when staining microtubule with a microtubule-binding protein fused to a protein tag. The higher resolution achieved with SiRtubulin highlights the potential of direct labeling of biological structures with fluorescent probes. SiR-tubulin also has been successfully used for live-cell SRM of the centrosome (Figure 7A). The centrosome is the major microtubule-organizing centre in the cell that is built around the centriole, a cylindrical structure composed of nine triplets of microtubules (Figure 7B). Live-cell STED of centrosomes revealed a cylindrical structure

Figure 6. Fluorophores used for photoswitching or photoactivation. (A) Alexa647. (B) Atto655. (C) HMSiR. (D) Diazoketone rhodamine. X-SH represents thiol-containing


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compounds. R groups show where the fluorophores are substituted for labeling purposes.

Figure 7. Live-cell STED imaging with small-molecule fluorescent probes. (A) STED image of centrosomal microtubules stained with SiRtubulin. The modulation in brightness along the rim of the cylinder (with polar angle  of 40°) is a consequence of the nine-fold symmetry of centrioles. Scale bar, 200 nm. (B) Cartoon of structure of a centriole with nine-fold symmetry structure. (C) Live-cell STED image of rat hippocampal neurons stained with SiR-actin, revealing the presence of actin rings. Scale bar, 1 μm. (D) Cartoon of the periodic, ring-like actin structure found in neuronal cells. (E) COS-7 cells expressing Halo-Sec61b (ER membrane) and SNAP-KDEL (ER lumen) fusion proteins labeled with ATTO590 and SiR respectively. Scale bar, 2 μm. (F) Scheme of ER structure showing ER lumen and membrane. (G) Two-color image of body wall muscle microtubule network stained with carbopyronine-tubulin and tracheoles stained with SiR-carboxyl. Insert shows zoom-in image of the region shown in panel as a white rectangle. Image shows microtubule network following tracheole. Scale bars: 5 mm in the large field of view and 1 mm in the zoom-in image. (H) Cartoon of tracheole structure. A and C: Adapted by permission from ref 48 Springer Nature Copyright 2014. E: Adapted from ref 71 published by Springer Nature under CC BY 4.0. G: Adapted from ref 27 published by the Royal Society of Chemistry under CC BY-NC 3.0.

with a diameter of 176 nm, which is in agreement with previous data obtained through electron microscopy (EM). Importantly, SRM allows the observation of live-cells, which not only enables to observe dynamic phenomena in real time inaccessible to EM but also avoids pitfalls of fixation procedures required for EM.90 Furthermore, STED images revealed a modulation in fluorescence intensity along the perimeter of the ring with measured polar angles between neighbouring maxima of approximately 40°, consistent with the nine-fold symmetry of the centriole. SiR-actin, which specifically labels F-actin, is another probe that together with STED imaging has provided unprecedented insights into the organization of the cytoskeleton in living cells. Specifically, STED-imaging of primary rat hippocampal neurons stained with SiR-actin revealed ring-shaped structures at the rim of axons, evenly spaced along axonal shafts with a periodicity of 180 nm (Figure 7C, D).48 These live-cell images confirmed earlier studies on fixed samples.91 Furthermore, subsequent live-cell STED with SiR-actin revealed that this periodic subcortical cytoskeletal structure is a general feature of cells of the nervous system.92-93 Spectroscopically distinguishable fluorescent probes have also been used for live-cell multicolor STED. Using a combination of Halo and SNAP-tagged proteins labeled with SiR and ATTO590, the dynamic interaction of different organelles such as the Golgi, endoplasmic reticulum (ER) or

mitochondria was followed over minutes by live-cell, dualcolor STED (Figure 7E, F).71 With the same combination of small-molecule fluorophores, it was later found that the small GTPase ARF1 is involved in the formation of long and thin tubular carries involved in Golgi transport.94 Another example of dual-color, live-cell STED is the imaging of microtubules and tracheoles in tissues of fruit flies. Interestingly, simple SiRcarboxyl without any ligand attached was used as fluorescent probe for the staining of tracheoles. This serendipitous finding is reminder that useful fluorescent probes for live-cell imaging can arise from careful analysis of perceived background signals (Figure 7G, H).27 Finally, the development of techniques such as two-photon STED and advances in the labeling of tissues as well as living animals will ultimately further expand the applicability of STED.95-96 Live-cell SMLM was also successfully used for characterizing biological processes and structures with high spatial and temporal resolution. An early example was the characterization of clathrin-mediated endocytosis of transferrin clusters.23 In these experiments, transferrin was chemically labeled with Alexa568 and clathrin-coated pits were labeled via SNAP-tag and Alexa647 (introduced by electroporation). Twocolor SMLM images revealed the morphology of the clathrin coat and the enclosed transferrin within 30 s (Figure 8A, B). Transferrin alone was imaged with time resolutions as high as 0.5 s revealing cluster formation and internalization.23



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Photoswitchable membrane-specific small-molecule fluorophores made it possible to observe structural dynamics of

Figure 8. Live-cell SMLM imaging with small-molecule fluorescent probes. (A) Dual-color SMLM image of clathrin coated pits labeled with Alexa647 (magenta) and transferrin labeled with Alexa568 (green). Top image xy projection. Lower left xy cross section, lower middle xz cross section through the middle of the pit, lower right cross section through the middle of the pit clathrin channel only. Scale bars, 500 nm and 100 nm respectively. (B) Cartoon of an invaginating clathrin coated pit together with its transferrin cargo. (C) SMLM image of the plasma membrane labeled with DiI in a hippocampal neuron. Scale bar, 1 µm. (D) Cartoon of hippocampal neuron showing dendritic spine necks. (E) 𝛃-Tubulin–HaloTag fusion proteins expressed in Vero cells were labeled with HMSiR–Halo. Sequential acquisition of superresolution images of microtubules at 0 min (white), 31 min (yellow) and 63 min (green). Scale bar, 2 µm. (F) Cartoon of tubulin structure forming with 𝛂-tubulin and 𝛃-tubulin. (G) SMLM image of the ER in HeLa cells. Upper left corner shows the diffraction limited image. Color indicates the time between 0 and 500 s of the localization. Averaged line profiles from the four yellow lines are shown in an insert. The profiles show FWHM values of 50 ± 3 nm (super-resolution image, yellow) and 367 ± 38 nm (diffraction-limited image, white dashed). Scale bar, 1 μm. (H) Cartoon of ER with periphery tubules. A: Adapted by permission from ref 23 Springer Nature Copyright 2011. C: Adapted from ref 31 published by the National Academy of Sciences under CC BY-NC-ND. E: Adapted by permission from ref 33 Springer Nature Copyright 2014. G: Adapted by permission from ref 97 Springer Nature Copyright 2017.

the plasma membrane, mitochondria, ER, lysosomes and the Golgi. 31, 50 For example, the lipophilic membrane dye 1,1'dioctadecyl-3,3,3',3'-tetramethylindocarbo-cyanine (DiI) was used to investigate the structure and dynamics of the plasma membrane of hippocampal neurons. The obtained resolution allowed to determine the width of dendritic spine necks in a snapshot taken within 15 s (Figure 8C, D). In addition, the dynamics of filopodia and dendritic spines were studied in a series of 10 s snapshots, revealing retraction and extension of different regions.31 The introduction of the previously discussed HMSiR 32, which switches without high intensity laser irradiation or the need for additives, allowed to perform SMLM over longer time periods.33 The decrease in irradiation intensity to about 10% of what is required for standard fluorophores in SMLM dramatically decreased photobleaching and photodamage. This allowed to monitor the movement of microtubules in 30 s snapshots every 10 min over a time span of one hour (Figure 8E, F).33 HMSiR was further used in combination with click chemistry, through which the fluorophore was localized to different membranes. As observed for other HIDE probes,63 the lipophilic membrane environment shifts the ‘on’/‘off’ ratio of HMSiR further to the ‘off’ state enabling to image densely labeled structures more easily. A HMSiR-based HIDE probe for the ER revealed the formation of 50 nm wide ER tubules at the

periphery of cells, which is consistent with previous EM data (Figure 8G, H).97 While these examples demonstrate the power of live-cell SMLM, broadening its scope will require advances in speed of acquisition, better image reconstruction algorithms and brighter, buffer independent small-molecule fluorophores.98-99

Conclusion and Outlook Live-cell SRM can provide insights into biological phenomena with unprecedented spatiotemporal resolution. As we have outlined above, the specific labeling of biomolecules with probes that possess suitable spectroscopic properties has become a major bottleneck to unleash the full potential of livecell SRM. It is instructive to summarize again the properties that a small-molecule fluorescent probe for live-cell imaging ideally should possess. The ideal probe would have (i) exquisite brightness and photostability, (ii) far-red excitation and emission wavelengths, (iii) appropriate switching properties, (iv) display fluorogenicity for no-wash labeling, (v) could be specifically attached to the biomolecule of interest without affecting its function or increasing its size, (vi) possesses good cell permeability, and (vii) displays minimal background staining. Unfortunately, a probe that fulfills all of these criteria has not yet been developed. The challenge in improving existing probes is that optimization for one of the properties listed above must not go on the expense of the others. Ideally,


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structural modifications will be identified that, for example improve brightness and photostability, without decreasing permeability or increase unspecific binding. The replacement of the dimethylamino group, which is ubiquitously found in fluorophores, by azetidines is an excellent example of such a modification: Replacing a dimethlyamino group with an azetidine increases brightness and photostability in various fluorophores while only adding one CH2 group to the structure. Chemists should also not be discouraged by the hurdles that will have to be overcome to create a perfect probe, as even probes that only partially fulfill the wish-list above can have a significant impact on biology. SiR-actin and SiR-tubulin, two probes for fluorescence labeling of the cytoskeleton are good examples: since their introduction in 2014, they have already been used in more than 250 publications. Surely, chemists will come up with even more powerful probes in the future. Furthermore, new labeling strategies are required that only minimally disturb the tagged biomolecule. Self-labeling protein based approaches profit form the advances in genome editing, which make cell lines with endogenously tagged proteins readily available, thereby avoiding overexpression artefacts. However, this does not circumvent the challenges imposed by the size of the tag, which can disturb the natural function of the protein. Hence, smaller protein tags with the same kind of specificity as the currently used tags would be needed. Methods that employ unnatural amino acids or chemical ligands that directly bind to proteins of interest have to be made more generally applicable. Currently, these approaches require extensive optimization for each protein and ligand. We can also expect that the ingenuity of physicists will continue to challenge probe developers. For example, in 2017, MINFLUX, a new concept of SRM was introduced.100 Here, the fluorophore is probed with a local minimum of excitation light, which minimizes the number of photons needed for SRM. MINFLUX increases the localization precision of SRM to ~1 nm and was used to resolve molecules which were only 6 nm apart. This resolution approaches the size of the currently used labeling tags, highlighting once more the need for new labeling chemistry. MINFLUX makes use of photoactivatable fluorophores and was first demonstrated using mEOS3.2 and Alexa647. However, to exploit its potential for live-cell SRM, new photoswitchable fluorophores would be highly welcomed. Finally, the peculiarities of MINFLUX, exploiting the induced absence of emission of a fluorophore for its localization, requires to minimize fluorescence signal arising from unspecific binding, which further raises the bar for the performance of small-molecule fluorescent probes. In summary, the development of new small-molecule fluorescent probes for live-cell SRM requires the solution of complex, interdependent problems. These probes then create the exciting opportunity to characterize biology on the nanoscale.

AUTHOR INFORMATION Corresponding Author * [email protected].

Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.

Funding Sources The authors acknowledge funding of the Max Planck Society and the École Polytechnique Fédérale de Lausanne (EPFL). LW is recipient of an Alexander von Humboldt Fellowship. AS is a recipient of a fellowship from Boehringer Ingelheim Fonds.

Notes KJ is inventor on patents on siliconrhodamines and SNAP-tag, filed by EPFL.

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