Long-Lived Intracellular Single-Molecule Fluorescence Using Electroporated Molecules

Studies of biomolecules in vivo are crucial to understand their function in a natural, biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins to study their expression, localization, and action; however, the scope of such studies would be increased considerably by using organic fluorophores, which are smaller and more photostable than their fluorescent protein counterparts. Here, we describe a straightforward, versatile, and high-throughput method to internalize DNA fragments and proteins labeled with organic fluorophores into live Escherichia coli by employing electroporation. We studied the copy numbers, diffusion profiles, and structure of internalized molecules at the single-molecule level in vivo, and were able to extend single-molecule observation times by two orders of magnitude compared to green fluorescent protein, allowing continuous monitoring of molecular processes occurring from seconds to minutes. We also exploited the desirable properties of organic fluorophores to perform single-molecule Förster resonance energy transfer measurements in the cytoplasm of live bacteria, both for DNA and proteins. Finally, we demonstrate internalization of labeled proteins and DNA into yeast Saccharomyces cerevisiae, a model eukaryotic system. Our method should broaden the range of biological questions addressable in microbes by single-molecule fluorescence.     

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisiæ, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.     

Single-Molecule Imaging at High Fluorophore Concentrations by Local Activation of Dye

Single-molecule fluorescence microscopy is a powerful tool for observing biomolecular interactions with high spatial and temporal resolution. Detecting fluorescent signals from individual labeled proteins above high levels of background fluorescence remains challenging, however. For this reason, the concentrations of labeled proteins in in vitro assays are often kept low compared to their in vivo concentrations. Here, we present a new fluorescence imaging technique by which single fluorescent molecules can be observed in real time at high, physiologically relevant concentrations. The technique requires a protein and its macromolecular substrate to be labeled each with a different fluorophore. Making use of short-distance energy-transfer mechanisms, only the fluorescence from those proteins that bind to their substrate is activated. This approach is demonstrated by labeling a DNA substrate with an intercalating stain, exciting the stain, and using energy transfer from the stain to activate the fluorescence of only those labeled DNA-binding proteins bound to the DNA. Such an experimental design allowed us to observe the sequence-independent interaction of Cy5-labeled interferon-inducible protein 16 with DNA and the sliding via one-dimensional diffusion of Cy5-labeled adenovirus protease on DNA in the presence of a background of hundreds of nanomolar Cy5 fluorophore.     

Lucky Imaging: Improved Localization Accuracy for Single Molecule Imaging

We apply the astronomical data-analysis technique, Lucky imaging, to improve resolution in single molecule fluorescence microscopy. We show that by selectively discarding data points from individual single-molecule trajectories, imaging resolution can be improved by a factor of 1.6 for individual fluorophores and up to 5.6 for more complex images. The method is illustrated using images of fluorescent dye molecules and quantum dots, and the in vivo imaging of fluorescently labeled linker for activation of T cells.     

DNA binding fluorescent proteins for the direct visualization of large DNA molecules

Fluorescent proteins that also bind DNA molecules are useful reagents for a broad range of biological applications because they can be optically localized and tracked within cells, or provide versatile labels for in vitro experiments. We report a novel design for a fluorescent, DNA-binding protein (FP-DBP) that completely ‘paints’ entire DNA molecules, whereby sequence-independent DNA binding is accomplished by linking a fluorescent protein to two small peptides (KWKWKKA) using lysine for binding to the DNA phosphates, and tryptophan for intercalating between DNA bases. Importantly, this ubiquitous binding motif enables fluorescent proteins (K_d = 14.7 μM) to confluently stain DNA molecules and such binding is reversible via pH shifts. These proteins offer useful robust advantages for single DNA molecule studies: lack of fluorophore mediated photocleavage and staining that does not perturb polymer contour lengths. Accordingly, we demonstrate confluent staining of naked DNA molecules presented within microfluidic devices, or localized within live bacterial cells.     

Optimized delivery of fluorescently labeled proteins in live bacteria using electroporation

Studying the structure and dynamics of proteins in live cells is essential to understanding their physiological activities and mechanisms, and to validating in vitro characterization. Improvements in labeling and imaging technologies are starting to allow such in vivo studies; however, a number of technical challenges remain. Recently, we developed an electroporation-based protocol for internalization, which allows biomolecules labeled with organic fluorophores to be introduced at high efficiency into live E. coli (Crawford et al. in Biophys J 105 (11):2439–2450, 2013). Here, we address important challenges related to internalization of proteins, and optimize our method in terms of (1) electroporation buffer conditions; (2) removal of dye contaminants from stock protein samples; and (3) removal of non-internalized molecules from cell suspension after electroporation. We illustrate the usability of the optimized protocol by demonstrating high-efficiency internalization of a 10-kDa protein, the ω subunit of RNA polymerase. Provided that suggested control experiments are carried out, any fluorescently labeled protein of up to 60 kDa could be internalized using our method. Further, we probe the effect of electroporation voltage on internalization efficiency and cell viability and demonstrate that, whilst internalization increases with increased voltage, cell viability is compromised. However, due to the low number of damaged cells in our samples, the major fraction of loaded cells always corresponds to non-damaged cells. By taking care to include only viable cells into analysis, our method allows physiologically relevant studies to be performed, including in vivo measurements of protein diffusion, localization and intramolecular dynamics via single-molecule Förster resonance energy transfer.     

Automated Analysis of Single-Molecule Photobleaching Data by Statistical Modeling of Spot Populations

The number of fluorophores within a molecule complex can be revealed by single-molecule photobleaching imaging. A widely applied strategy to analyze intensity traces over time is the quantification of photobleaching step counts. However, several factors can limit and bias the detection of photobleaching steps, including noise, high numbers of fluorophores, and the possibility that several photobleaching events occur almost simultaneously. In this study, we propose a new approach, to our knowledge, to determine the fluorophore number that correlates the intensity decay of a population of molecule complexes with the decay of the number of visible complexes. We validated our approach using single and fourfold Atto-labeled DNA strands. As an example we estimated the subunit stoichiometry of soluble CD95L using GFP fusion proteins. To assess the precision of our method we performed in silico experiments showing that the estimates are not biased for experimentally observed intensity fluctuations and that the relative precision remains constant with increasing number of fluorophores. In case of fractional fluorescent labeling, our simulations predicted that the fluorophore number estimate corresponds to the product of the true fluorophore number with the labeling fraction. Our method, denoted by spot number and intensity correlation (SONIC), is fully automated, robust to noise, and does not require the counting of photobleaching events.     

In?Vivo Nonlinear Optical Imaging of Immune Responses: Tissue Injury and Infection

In this study, we demonstrate a noninvasive imaging approach based on multimodal nonlinear optical microscopy to in vivo image the responses of immune cells (neutrophils) to the tissue injury and bacterial infection in a zebrafish model. Specifically, the second harmonic generation from myosin thick filaments in sarcomere enabled a clear visualization of the muscle injury and infection. Two-photon excited fluorescence was used to track the behavior of the neutrophils that were transgenically labeled by red fluorescent protein. The corresponding reduced nicotinamide adenine dinucleotide (NADH) two-photon excited fluorescence images revealed a detailed morphological transformation process of individual neutrophils during muscle tissue injury and bacterial infection. The analysis of time-resolved NADH signals from the neutrophils provided important biological insights of the cellular energy metabolism during the immune responses. We found a significant increase of free/protein-bound NADH ratios in activated neutrophils in bacterial-infected tissue. In this study, we also discovered that, under 720 nm excitation, two wild-type strains (DH5α and BL21) of bacteria Escherichia coli emitted distinct endogenous fluorescence of double-peak at ∼450 and ∼520 nm, respectively. We demonstrated that the double-peak fluorescence signal could be used to differentiate the E. coli from surrounding tissues of dominant NADH signals, and to achieve label-free tracking of E. coli bacteria in vivo.     

Autofluorescent proteins in single-molecule research: applications to live cell imaging microscopy

The spectral and photophysical characteristics of the autofluorescent proteins were analyzed and compared to flavinoids to test their applicability for single-molecule microscopy in live cells. We compare 1) the number of photons emitted by individual autofluorescent proteins in artificial and in vivo situations, 2) the saturation intensities of the various autofluorescent proteins, and 3) the maximal emitted photons from individual fluorophores in order to specify their use for repetitive imaging and dynamical analysis. It is found that under relevant conditions and for millisecond integration periods, the autofluorescent proteins have photon emission rates of approximately 3000 photons/ms (with the exception of DsRed), saturation intensities from 6 to 50 kW/cm2, and photobleaching yields from 10(-4) to 10(-5). Definition of a detection ratio led to the conclusion that the yellow-fluorescent protein mutant eYFP is superior compared to all the fluorescent proteins for single-molecule studies in vivo. This finding was subsequently used for demonstration of the applicability of eYFP in biophysical research. From tracking the lateral and rotational diffusion of eYFP in artificial material, and when bound to membranes of live cells, eYFP is found to dynamically track the entity to which it is anchored.     

New strategies for fluorescently labeling proteins in the study of amyloids

Amyloid proteins are widely studied, both for their unusual biophysical properties and their association with disorders such as Alzheimer’s and Parkinson’s disease. Fluorescence-based methods using site-specifically labeled proteins can provide information on the details of their structural dynamics and their roles in specific biological processes. Here, we describe the application of different labeling methods and novel fluorescent probe strategies to the study of amyloid proteins, both for in vitro biophysical experiments and for in vivo imaging. These labeling tools can be elegantly used to answer important questions on the function and pathology of amyloid proteins.     

Dynamics of the Serine Chemoreceptor in the Escherichia coli Inner Membrane: A High-Speed Single-Molecule Tracking Study

We investigated the mobility of the polar localized serine chemoreceptor, Tsr, labeled by the fluorescent protein Venus in the inner membrane of live Escherichia coli cells at observation rates up to 1000 Hz. A fraction (7%) of all Tsr molecules shows free diffusion over the entire cell surface with an average diffusion coefficient of 0.40 ± 0.01 μm² s⁻¹. The remaining molecules were found to be ultimately confined in compartments of size 290 ± 15 nm and showed restricted diffusion at an inner barrier found at 170 ± 10 nm. At the shortest length-scales (<170 nm), all Tsr molecules diffuse equally. Disruption of the cytoskeleton and rounding of the cells resulted in an increase in the mobile fraction of Tsr molecules and a fragmenting of the previously polar cluster of Tsr consistent with a curvature-based mechanism of Tsr cluster maintenance.     

Solution-Based Single-Molecule FRET Studies of K+ Channel Gating in a Lipid Bilayer

Ion channels are dynamic multimeric proteins that often undergo multiple unsynchronized structural movements as they switch between their open and closed states. Such structural changes are difficult to measure within the context of a native lipid bilayer and have often been monitored via macroscopic changes in Förster resonance energy transfer (FRET) between probes attached to different parts of the protein. However, the resolution of this approach is limited by ensemble averaging of structurally heterogeneous subpopulations. These problems can be overcome by measurement of FRET in single molecules, but this presents many challenges, in particular the ability to control labeling of subunits within a multimeric protein with acceptor and donor fluorophores, as well as the requirement to image large numbers of individual molecules in a membrane environment. To address these challenges, we randomly labeled tetrameric KirBac1.1 potassium channels, reconstituted them into lipid nanodiscs, and performed single-molecule FRET confocal microscopy with alternating-laser excitation as the channels diffused in solution. These solution-based single-molecule FRET measurements of a multimeric ion channel in a lipid bilayer have allowed us to probe the structural changes that occur upon channel activation and inhibition. Our results provide direct evidence of the twist-to-shrink movement of the helix bundle crossing during channel gating and demonstrate how this method might be applied to real-time structural studies of ion channel gating.     

Condensed Mitotic Chromosome Structure at Nanometer Resolution Using PALM and EGFP- Histones

Photoactivated localization microscopy (PALM) and related fluorescent biological imaging methods are capable of providing very high spatial resolutions (up to 20 nm). Two major demands limit its widespread use on biological samples: requirements for photoactivatable/photoconvertible fluorescent molecules, which are sometimes difficult to incorporate, and high background signals from autofluorescence or fluorophores in adjacent focal planes in three-dimensional imaging which reduces PALM resolution significantly. We present here a high-resolution PALM method utilizing conventional EGFP as the photoconvertible fluorophore, improved algorithms to deal with high levels of biological background noise, and apply this to imaging higher order chromatin structure. We found that the emission wavelength of EGFP is efficiently converted from green to red when exposed to blue light in the presence of reduced riboflavin. The photon yield of red-converted EGFP using riboflavin is comparable to other bright photoconvertible fluorescent proteins that allow <20 nm resolution. We further found that image pre-processing using a combination of denoising and deconvolution of the raw PALM images substantially improved the spatial resolution of the reconstruction from noisy images. Performing PALM on Drosophila mitotic chromosomes labeled with H2AvD-EGFP, a histone H2A variant, revealed filamentous components of ∼70 nm. This is the first observation of fine chromatin filaments specific for one histone variant at a resolution approximating that of conventional electron microscope images (10–30 nm). As demonstrated by modeling and experiments on a challenging specimen, the techniques described here facilitate super-resolution fluorescent imaging with common biological samples.     

KERA: analysis tool for multi-process,multi-state single-molecule data

Molecular machines within cells dynamically assemble, disassemble and reorganize. Molecular interactions between their components can be observed at the single-molecule level and quantified using colocalization single-molecule spectroscopy, in which individual labeled molecules are seen transiently associating with a surface-tethered partner, or other total internal reflection fluorescence microscopy approaches in which the interactions elicit changes in fluorescence in the labeled surface-tethered partner. When multiple interacting partners can form ternary, quaternary and higher order complexes, the types of spatial and temporal organization of these complexes can be deduced from the order of appearance and reorganization of the components. Time evolution of complex architectures can be followed by changes in the fluorescence behavior in multiple channels. Here, we describe the kinetic event resolving algorithm (KERA), a software tool for organizing and sorting the discretized fluorescent trajectories from a range of single-molecule experiments. KERA organizes the data in groups by transition patterns, and displays exhaustive dwell time data for each interaction sequence. Enumerating and quantifying sequences of molecular interactions provides important information regarding the underlying mechanism of the assembly, dynamics and architecture of the macromolecular complexes. We demonstrate KERA’s utility by analyzing conformational dynamics of two DNA binding proteins: replication protein A and xeroderma pigmentosum complementation group D helicase.     

Single-Molecule Force Spectroscopy Study on the Mechanism of RNA Disassembly in Tobacco Mosaic Virus

To explore the disassembly mechanism of tobacco mosaic virus (TMV), a model system for virus study, during infection, we have used single-molecule force spectroscopy to mimic and follow the process of RNA disassembly from the protein coat of TMV by the replisome (molecular motor) in vivo, under different pH and Ca²⁺ concentrations. Dynamic force spectroscopy revealed the unbinding free-energy landscapes as that at pH 4.7 the disassembly process is dominated by one free-energy barrier, whereas at pH 7.0 the process is dominated by one barrier and that there exists a second barrier. The additional free-energy barrier at longer distance has been attributed to the hindrance of disordered loops within the inner channel of TMV, and the biological function of those protein loops was discussed. The combination of pH increase and Ca²⁺ concentration drop could weaken RNA-protein interactions so much that the molecular motor replisome would be able to pull and disassemble the rest of the genetic RNA from the protein coat in vivo. All these facts provide supporting evidence at the single-molecule level, to our knowledge for the first time, for the cotranslational disassembly mechanism during TMV infection under physiological conditions.     

Fluorophores for live cell imaging of AGT fusion proteins across the visible spectrum

O6-alkylguanine-DNA alkyltransferase (AGT) fusion proteins can be specifically and covalently labeled with fluorescent O6-benzylguanine (O6-BG) derivatives for multicolor live cell imaging approaches. Here, we characterize several new BG fluorophores suitable for in vivo AGT labeling that display fluorescence emission maxima covering the visible spectrum from 472 to 673 nm, thereby extending the spectral limits set by fluorescent proteins. We show that the photostability of the cell-permeable dyes BG Rhodamine Green (BG505) and CP tetramethylrhodamine (CP-TMR) is in the range of enhanced green fluorescent protein (EGFP) and monomeric red fluorescent protein (mRFP), and that BG diethylaminomethyl coumarin (BGDEAC), a derivative of coumarin, is even more stable than enhanced cyan fluorescent protein (ECFP). Due to the increasing number of new BG derivatives with interesting fluorescence properties, such as far-red emission, fluorescence labeling of AGT fusion proteins is becoming a versatile alternative to existing live cell imaging approaches.     

High-temperature single-molecule kinetic analysis of thermophilic archaeal MCM helicases

The minichromosome maintenance (MCM) complex is the replicative helicase responsible for unwinding DNA during archaeal and eukaryal genome replication. To mimic long helicase events in the cell, a high-temperature single-molecule assay was designed to quantitatively measure long-range DNA unwinding of individual DNA helicases from the archaeons Methanothermobacter thermautotrophicus (Mth) and Thermococcus sp. 9°N (9°N). Mth encodes a single MCM homolog while 9°N encodes three helicases. 9°N MCM3, the proposed replicative helicase, unwinds DNA at a faster rate compared to 9°N MCM2 and to Mth MCM. However, all three MCM proteins have similar processivities. The implications of these observations for DNA replication in archaea and the differences and similarities among helicases from different microorganisms are discussed. Development of the high-temperature single-molecule assay establishes a system to comprehensively study thermophilic replisomes and evolutionary links between archaeal, eukaryal, and bacterial replication systems.     

Single-Cell Quantitative Proteomic Analysis of Human Oocyte Maturation Revealed High Heterogeneity in In Vitro–Matured Oocytes

Oocyte maturation is pertinent to the success of in vitro maturation (IVM), which is used to overcome female infertility, and produced over 5000 live births worldwide. However, the quality of human IVM oocytes has not been investigated at single-cell proteome level. Here, we quantified 2094 proteins in human oocytes during in vitro and in vivo maturation (IVO) by single-cell proteomic analysis and identified 176 differential proteins between IVO and germinal vesicle oocytes and 45 between IVM and IVO oocytes including maternal effect proteins, with potential contribution to the clinically observed decreased fertilization, implantation, and birth rates using human IVM oocytes. IVM and IVO oocytes showed separate clusters in principal component analysis, with higher inter-cell variability among IVM oocytes, and have little correlation between mRNA and protein changes during maturation. The patients with the most aberrantly expressed proteins in IVM oocytes had the lowest level of estradiol per mature follicle on trigger day. Our data provide a rich resource to evaluate effect of IVM on oocyte quality and study mechanism of oocyte maturation.