GMars-T Enabling Multimodal Subdiffraction Structural and Functional Fluorescence Imaging in Live Cells.

The BiFC (bimolecular fluorescence complementation) assay and BiFC combined with FRET (fluorescence resonance energy transfer) technique have become important tools for molecular interaction studies in live cells. However, the real detection and cellular imaging performances of most existing red fluorescent protein-derived BiFC assays still suffer from relatively low ensemble brightness, high cytotoxicity, the red fluorescent proteins being prone-to-aggregation or severe residual dimerization, inefficient complementation and slow maturation at 37 °C physiological temperature in live mammalian cells. We developed a BiFC assay based on a recently evolved truly monomeric red fluorescent protein (FP) mScarlet-I with excellent cellular performances such as low cytotoxicity, fast and efficient chromophore maturation and the highest in-cell brightness among all previously reported monomeric red fluorescent proteins. In this work, a classic β-Fos/β-Jun constitutive heterodimerization model and a rapamycin-inducible FRB/FKBP interaction system were used to establish and test the performance of the mScarlet-I-based BiFC assay in live mammalian cells. Furthermore, simply by adopting the large-Stokes-shift fluorescent protein mAmetrine as the donor, β-Jun-β-Fos-NFAT1 ternary protein complex formation could be readily and efficiently detected and visualized with minimal spectral cross-talk in live HeLa cells by combining live-cell sensitized-emission FRET measurement with the mScarlet-I-based BiFC assay. The currently established BiFC assay in this work was also shown to be able to detect and visualize various protein-protein interactions (PPIs) at different subcellular compartments with high specificity and sensitivity at 37 °C physiological temperature in live mammalian cells.

Genetic code expansion and bioorthogonal labeling provide for the first time a way for direct, site-specific labeling of proteins with fluorescent-dyes in live cells. Although the small size and superb photophysical parameters of fluorescent-dyes offer unique advantages for high-resolution microscopy, this approach has yet to be embraced as a tool in live cell imaging. Here we evaluated the feasibility of this approach by applying it for α-tubulin labeling. After a series of calibrations, we site-specifically labeled α-tubulin with silicon rhodamine (SiR) in live mammalian cells in an efficient and robust manner. SiR-labeled tubulin successfully incorporated into endogenous microtubules at high density, enabling video recording of microtubule dynamics in interphase and mitotic cells. Applying this labeling approach to structured illumination microscopy resulted in an increase in resolution, highlighting the advantages in using a smaller, brighter tag. Therefore, using our optimized assay, genetic code expansion provides an attractive tool for labeling proteins with a minimal, bright tag in quantitative high-resolution imaging.

Over the past years, fluorescent proteins (e.g., green fluorescent proteins) have been widely utilized to visualize recombinant protein expression and localization in live cells. Although powerful, fluorescent protein tags are limited by their relatively large sizes and potential perturbation to protein function. Alternatively, site-specific labeling of proteins with small-molecule organic fluorophores using bioorthogonal chemistry may provide a more precise and less perturbing method. This approach involves site-specific incorporation of unnatural amino acids (UAAs) into proteins via genetic code expansion, followed by bioorthogonal chemical labeling with small organic fluorophores in living cells. While this approach has been used to label extracellular proteins for live cell imaging studies, site-specific bioorthogonal labeling and fluorescence imaging of intracellular proteins in live cells is still challenging. Herein, we systematically evaluate site-specific incorporation of diastereomerically pure bioorthogonal UAAs bearing stained alkynes or alkenes into intracellular proteins for inverse-electron-demand Diels-Alder cycloaddition reactions with tetrazine-functionalized fluorophores for live cell labeling and imaging in mammalian cells. Our studies show that site-specific incorporation of axial diastereomer of trans-cyclooct-2-ene-lysine robustly affords highly efficient and specific bioorthogonal labeling with monosubstituted tetrazine fluorophores in live mammalian cells, which enabled us to image the intracellular localization and real-time dynamic trafficking of IFITM3, a small membrane-associated protein with only 137 amino acids, for the first time. Our optimized UAA incorporation and bioorthogonal labeling conditions also enabled efficient site-specific fluorescence labeling of other intracellular proteins for live cell imaging studies in mammalian cells.

In situ spatiotemporal microRNA (miRNA) imaging in mammal cells plays an essential role in illustrating its structures and biological functions. Herein, we proposed a near-infrared (NIR) light-activated nanoprobe for high-sensitive in situ controllable miRNA imaging in living cells. The NIR-activated nanoprobe employed an upconversion nanoparticle that acted as a NIR-to-UV transducer to trigger the following photocleavage toward a dumbbell DNA probe tethered on the surface of the nanoparticle. The structure change of the dumbbell probe then induced a catalytic hairpin assembly of target miRNAs, by which in situ readout of the amplified fluorescence signal was enabled. Additionally, both intracellular miRNA imaging and accurate quantification in live cells were realized without damaging the cell membranes. Compared with conventional in situ strategies, the proposed approach remarkedly increases imaging efficiency by eliminating those unfavored intercellular molecular imaging backgrounds. We assured that the proposed NIR-activated miRNA sensing strategy will add to the advancement for bioanalysis in living systems, which is of crucial importance in the diagnosis of various human diseases, especially cancers.

Proper membrane localization of ion channels is essential for the function of neuronal cells. Particularly, the computational ability of dendrites depends on the localization of different ion channels in specific subcompartments. However, the molecular mechanisms that control ion channel localization in distinct dendritic subcompartments are largely unknown. Here, we developed a quantitative live cell imaging method to analyze protein sorting and post-Golgi vesicular trafficking. We focused on two dendritic voltage-gated potassium channels that exhibit distinct localizations: Kv2.1 in proximal dendrites and Kv4.2 in distal dendrites. Our results show that Kv2.1 and Kv4.2 channels are sorted into two distinct populations of vesicles at the Golgi apparatus. The targeting of Kv2.1 and Kv4.2 vesicles occurred by distinct mechanisms as evidenced by their requirement for specific peptide motifs, cytoskeletal elements, and motor proteins. By live cell and super-resolution imaging, we identified a novel trafficking machinery important for the localization of Kv2.1 channels. Particularly, we identified non-muscle myosin II as an important factor in Kv2.1 trafficking. These findings reveal that the sorting of ion channels at the Golgi apparatus and their subsequent trafficking by unique molecular mechanisms are crucial for their specific localizations within dendrites.

By introducing beam-scanning multifocal multiphoton 4Pi-confocal microscopy, we have attained fast fluorescence imaging of live cells with axial super resolution. Rapid scanning of up to 64 pairs of interfering high-angle fields and subsequent confocal detection enabled us to perform three to five times finer optical sectioning than confocal microscopy. In conjunction with nonlinear image restoration, we demonstrate, to our knowledge for the first time, three-dimensional imaging of live eukaryotic cells at an equilateral resolution of approximately 100 nm. This imaging mode allowed us to reveal the morphology and size of the green fluorescent protein-labeled mitochondrial compartment of live Saccharomyces cerevisiae (bakers' yeast) growing on different carbon sources. Our studies show that mitochondria of cells grown on medium containing glycerol as the only carbon source, as opposed to glucose-grown cells, exhibit a strongly branched tubular reticulum. We determine the average tubular diameter and find that it increases from 339 +/- 5 nm to 360 +/- 4 nm when changing from glucose to glycerol, that is, from a fermentable to a nonfermentable carbon source. Moreover, this change is associated with a 2.8-fold increase of the surface of the reticulum, resulting in an average increase in volume of the mitochondrial compartment by a factor of 3.0 +/- 0.2.

Development of a method for the detection of Cu2+ in the environment and live cells using a synthesized spider web-like fluorescent probe

Deficiency of the lysosomal glycoside hydrolase glucocerebrosidase (GCase) leads to abnormal accumulation of glucosyl ceramide in lysosomes and the development of the lysosomal storage disease known as Gaucher's disease. More recently, mutations in the GBA1 gene that encodes GCase have been uncovered as a major genetic risk factor for Parkinson's disease (PD). Current therapeutic strategies to increase GCase activity in lysosomes involve enzyme replacement therapy (ERT) and molecular chaperone therapy. One challenge associated with developing and optimizing these therapies is the difficulty in determining levels of GCase activity present within the lysosomes of live cells. Indeed, visualizing the activity of endogenous levels of any glycoside hydrolases, including GCase, has proven problematic within live mammalian cells. Here we describe the successful modular design and synthesis of fluorescence-quenched substrates for GCase. The selection of a suitable fluorophore and quencher pair permits the generation of substrates that allow convenient time-dependent monitoring of endogenous GCase activity within cells as well as localization of activity within lysosomes. These efficiently quenched (∼99.9%) fluorescent substrates also permit assessment of GCase inhibition in live cells by either confocal microscopy or high content imaging. Such substrates should enable improved understanding of GCase in situ as well the optimization of small-molecule chaperones for this enzyme. These findings also suggest routes to generate fluorescence-quenched substrates for other mammalian glycoside hydrolases for use in live cell imaging.

Quantum dots (QDs) and other nanoparticles require delivery and targeting for most intracellular applications. Despite many advances, intracellular delivery and targeting remains inefficient with many QDs remaining bound to the plasma membrane rather than internalized into the cell. The fluorescence resulting from these extracellular QDs results in a background signal that competes with intracellular QDs of interest. We present two methods for the reduction and discrimination of signal resulting from plasma membrane-bound QDs. The first method, a photophysical approach, uses an extracellular quencher to greatly reduce the fluorescence signal from extracellular QDs. This method is compatible with fast, widefield, fluorescence imaging in live cells. Results are presented for two extracellular quenchers, QSY-21 and trypan blue, used in combination with 655 nm emitting QDs. The use of an extracellular quencher can be extended to a wide variety of fluorophores. The second method uses transmission electron microscopy (TEM) to image thin (60-70 nm) slices of resin-embedded cells. The use of sectioned cells and high-resolution TEM makes it possible to discriminate between plasma membrane-bound and intracellular QDs. To overcome the difficulties associated with using TEM to image individual QDs in cells, we have utilized a silver enhancement method that significantly improves the contrast of QDs in TEM images.

Live-Cell Super-Resolution Fluorescence Imaging at High Spatiotemporal Resolutions

Force generation by actin assembly shapes cellular membranes. An experimentally constrained multiscale model shows that a minimal branched actin network is sufficient to internalize endocytic pits against membrane tension. Around 200 activated Arp2/3 complexes are required for robust internalization. A newly developed molecule-counting method determined that ~200 Arp2/3 complexes assemble at sites of clathrin-mediated endocytosis in human cells. Simulations predict that actin self-organizes into a radial branched array with growing ends oriented toward the base of the pit. Long actin filaments bend between attachment sites in the coat and the base of the pit. Elastic energy stored in bent filaments, whose presence was confirmed by cryo-electron tomography, contributes to endocytic internalization. Elevated membrane tension directs more growing filaments toward the base of the pit, increasing actin nucleation and bending for increased force production. Thus, spatially constrained actin filament assembly utilizes an adaptive mechanism enabling endocytosis under varying physical constraints.

Live cell imaging using confocal microscope is an important and popular technique used by biologists to observe and understand biological events occuring in a living cell. It allows simultaneous understanding of the dynamics and functions of many cellular processes in living cells. The principles of live cell imaging are different from that of fixed cell imaging as in live cells, pigments and fluorescent biomolecules are present in their functional state unlike in fixed cell imaging where they are removed. This poses various challenges in live cell imaging, such as maintaining cell viability under high photobleaching conditions and the use of optimal fluorescent components to overcome the artifacts. Given the multitude of advantages of live cell imaging over conventional microscopy, the purpose of this chapter is to provide a basic understanding of the approaches used to visualize plant cells using confocal microscopy, discuss some common challenges encountered during live cell imaging and provide suggestions to overcome them.

Genetically encoded red fluorescent proteins with a large Stokes shift (LSSRFPs) can be efficiently co-excited with common green FPs both under single- and two-photon microscopy, thus enabling dual-color imaging using a single laser. Recent progress in protein development resulted in a great variety of novel LSSRFPs; however, the selection of the right LSSRFP for a given application is hampered by the lack of a side-by-side comparison of the LSSRFPs’ performance. In this study, we employed rational design and random mutagenesis to convert conventional bright RFP mScarlet into LSSRFP, called LSSmScarlet, characterized by excitation/emission maxima at 470/598 nm. In addition, we utilized the previously reported LSSRFPs mCyRFP1, CyOFP1, and mCRISPRed as templates for directed molecular evolution to develop their optimized versions, called dCyRFP2s, dCyOFP2s and CRISPRed2s. We performed a quantitative assessment of the developed LSSRFPs and their precursors in vitro on purified proteins and compared their brightness at 488 nm excitation in the mammalian cells. The monomeric LSSmScarlet protein was successfully utilized for the confocal imaging of the structural proteins in live mammalian cells and multicolor confocal imaging in conjugation with other FPs. LSSmScarlet was successfully applied for dual-color two-photon imaging in live mammalian cells. We also solved the X-ray structure of the LSSmScarlet protein at the resolution of 1.4 Å that revealed a hydrogen bond network supporting excited-state proton transfer (ESPT). Quantum mechanics/molecular mechanics molecular dynamic simulations confirmed the ESPT mechanism of a large Stokes shift. Structure-guided mutagenesis revealed the role of R198 residue in ESPT that allowed us to generate a variant with improved pH stability. Finally, we showed that LSSmScarlet protein is not appropriate for STED microscopy as a consequence of LSSRed-to-Red photoconversion with high-power 775 nm depletion light.

Despite its small size of 5-8 mum - only one order of magnitude above the wavelength of visible light - yeast has developed into an attractive system for light microscopic analysis. First, the ease of genetic manipulation and integrative transformation have opened numerous experimental strategies for genome-wide tagging approaches, e.g., with fluorescent proteins (as discussed in several chapters of this issue). Second, the large number of cells that can be simultaneously visualized provides an excellent basis for statistical image analysis, resulting in reliable morphological or localization information. Third, the flexibility of yeast cultivation in terms of biochemical manipulation, rapid cellular growth, mutant isolation or drug susceptibility offers an unprecedented spectrum of possibilities for in vivo functional studies, and analysis of cellular dynamics and organelle inheritance. Although yeast in itself is an interesting cellular system, its "prototype character" in understanding cellular metabolism, physiology, and signaling in eukaryotes accounts for its popular use in technology development and biomedical research.Here we discuss experimental strategies for live yeast cell imaging, geared towards imaging-based large-scale screens. Major emphasis is on the methods for immobilizing cells under "physiological" conditions, with minimum impact on yeast. We also point out potential pitfalls resulting from live cell imaging that once again stresses the necessity for extremely careful experimental design and interpretation of data resulting from imaging experiments. It goes without saying that these problems are not restricted to yeast and are also highly relevant to "large" cells. If an image tells more than a thousand (perhaps misleading?) words, the ease of obtaining "images" thus rather suggests analyzing many thousands of images, to come up with one relevant and biologically significant conclusion.

The widespread application of fluorescence microscopy to study live cells has led to a greater understanding of numerous biological processes. Many techniques have been developed to uniquely label structures and track metabolic pathways using fluorophores in live cells. However, the photochemistry of nonnative compounds and the deposition of energy into the cell during imaging can result in unexpected and unwanted side effects. Herein, we examine potential live cell damage by first discussing common imaging considerations and modalities in fluorescence microscopy. We then consider several mechanisms by which various photochemical and photophysical phenomena cause cellular damage and introduce techniques that have leveraged these phenomena to intentionally create damage inside cells. Reviewing conditions under which intentional damage occurs can allow one to better predict when unintentional damage may be important. Finally, we delineate ways of checking for and reducing photochemical and photophysical damage.