Sensitive detection of histamine using fluorescently labeled oxido-reductases

A detection scheme is described by which the histamine contents of biological samples can be established. The scheme is based on the use of methylamine dehydrogenase (MADH) which converts primary amines into the corresponding aldehydes and ammonia. The generated reducing equivalents are subsequently transferred to the physiological partner of MADH, amicyanin, which thereby is converted from the oxidized blue-colored form into the reduced colorless form. The change in absorption is detected by monitoring the fluorescence of a covalently attached Cy5 dye label whose fluorescence is (partly) quenched by Förster resonance energy transfer (FRET) to the Cu-site of the amicyanin. The quenching efficiency and, thereby, the label fluorescence, depends on the oxidation state of the amicyanin. When adding histamine to the assay mixture the proportionality between the substrate concentration and the observed rate of the fluorescence increase has enabled this assay as a sensor method with high sensitivity. The MADH and amicyanin composition can be tuned so that the sensor can be adapted over a broad range of histamine concentrations (13 nM–225 μM). The lowest concentration detected so far is 13 nM of histamine. The sensor retained its linearity up to 225 μM with a coefficient of variation of 11% for 10 measurements of 100 nM histamine in a 100 μL sample volume. The use of a label fluorescing around 660 nm helps circumventing the interference from background fluorescence in biological samples. The sensor has been tested to detect histamine in biological fluids such as fish extracts and blood serum.     

Binding of protein kinase substrates by fluorescently labeled calmodulin

A synthetic protein kinase substrate, PRO-LEU-SER-ARG-THR-LEU-SER-VAL-SER-SER-NH₂, undergoes calcium-dependent binding by calmodulin. Phosphorylation of the peptide decreases its affinity for calmodulin with the dissociation constant increasing from 2.4 to $\text{ca}$. 7 mM. The results are consistent with the suggestion that calmodulin and the cAMP-dependent protein kinase can act on common recognition sequences.     

Quantitative characterization of the binding of fluorescently labeled colchicine to tubulin in vitro using fluorescence correlation spectroscopy

Fluorescence correlation spectroscopy (FCS) is a new technique that allows the determination of the diffusion constant of a fluorescent molecule in solution. Also, the binding of the fluorescent molecule to a target can be analyzed, if the difference in the diffusion coefficients of the free and bound ligand is sufficiently large. With FCS, the interaction between fluorescein-colchicine (FC) and tubulin has been studied in vitro. A fast and reversible binding is observed with an association constant at room temperature of (3.9 +/- 0.1) x 10(4) M-1. No competition with colchicine is seen, indicating that FCS reveals the existence of a new binding site on tubulin. FCS is not able to show the binding of FC to the original colchicine binding site, even though it exists, because the fluorescence of FC is strongly quenched upon binding to this site. This quenching is evident in spectrofluorometry experiments, revealing a slow binding of FC to tubulin that is subject to competition with colchicine. FCS allows the determination of the diffusion coefficients of both free and bound fluorescent colchicine which were found to be (2.6 +/- 0.2) x 10(-)10 and (2.0 +/- 0.2) x 10(-)11 m2 s-1, respectively. It can be concluded that fluorescent labeling, especially of small molecules, can interfere considerably with the binding behavior that is being studied. Although general qualitative effects in vivo are similar for colchicine and its fluorescein derivative, this quantitative study of the binding to tubulin presents a nuanced view, and the existence of a second binding site for FC can even explain some conflicting indications in the literature.     

Analysis of cell division using fluorescently labeled actin and myosin in living PtK2 cells

Actin and the light chains of myosin were labeled with fluorescent dyes and injected into interphase PtK2 cells in order to study the changes in distribution of actin and myosin that occurred when the injected cells subsequently entered mitosis and divided. The first changes occurred when stress fibers in prophase cells began to disassemble. During this process, which began in the center of the cell, individual fibers shortened, and in a few fibers, adjacent bands of fluorescent myosin could be seen to move closer together. In most cells, stress fiber disassembly was complete by metaphase, resulting in a diffuse distribution of the fluorescent proteins throughout the cytoplasm with the greatest concentration present in the mitotic spindle. The first evidence of actin and myosin concentration in a cleavage ring occurred at late anaphase, just before furrowing could be detected. Initially, the intensity of fluorescence and the width of the fluorescent ring increased as the ring constricted. In cells with asymmetrically positioned mitotic spindles, both protein concentration and furrowing were first evident in the cortical regions closest to the equator of the mitotic spindle. As cytokinesis progressed in such asymmetrically dividing cells, fluorescent actin and myosin appeared at the opposite side of the cell just before furrowing activity could be seen there. At the end of cytokinesis, myosin and actin were concentrated beneath the membrane of the midbody and subsequently became organized in two rings at either end of the midbody.     

Visualization of intermediate filaments in living cells using fluorescently labeled desmin

Fluorescently labeled desmin was incorporated into intermediate filaments when microinjected into living tissue culture cells. The desmin, purified from chicken gizzard smooth muscle and labeled with the fluorescent dye iodoacetamido rhodamine, was capable of forming a network of 10-nm filaments in solution. The labeled protein associated specifically with the native vimentin filaments in permeabilized, unfixed interphase and mitotic PtK2 cells. The labeled desmin was microinjected into living, cultured embryonic skeletal myotubes, where it became incorporated in straight fibers aligned along the long axis of the myotubes. Upon exposure to nocodazole, microinjected myotubes exhibited wavy, fluorescent filament bundles around the muscle nuclei. In PtK2 cells, an epithelial cell line, injected desmin formed a filamentous network, which colocalized with the native vimentin intermediate filaments but not with the cytokeratin networks and microtubular arrays. Exposure of the injected cells to nocadazole or acrylamide caused the desmin network to collapse and form a perinuclear cap that was indistinguishable from vimentin caps in the same cells. During mitosis, labeled desmin filaments were excluded from the spindle area, forming a cage around it. The filaments were partitioned into two groups either during anaphase or at the completion of cytokinesis. In the former case, the perispindle desmin filaments appeared to be stretched into two parts by the elongating spindle. In the latter case, a continuous bundle of filaments extended along the length of the spindle and appeared to be pinched in two by the contracting cleavage furrow. In these cells, desmin filaments were present in the midbody where they gradually were removed as the desmin filament network became redistributed throughout the cytoplasm of the spreading daughter 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.     

Analysis of myofibrillar structure and assembly using fluorescently labeled contractile proteins

To study how contractile proteins become organized into sarcomeric units in striated muscle, we have exposed glycerinated myofibrils to fluorescently labeled actin, alpha-actinin, and tropomyosin. In this in vitro system, alpha-actinin bound to the Z-bands and the binding could not be saturated by prior addition of excess unlabeled alpha-actinin. Conditions known to prevent self-association of alpha-actinin, however, blocked the binding of fluorescently labeled alpha-actinin to Z-bands. When tropomyosin was removed from the myofibrils, alpha-actinin then added to the thin filaments as well as the Z-bands. Actin bound in a doublet pattern to the regions of the myosin filaments where there were free cross-bridges i.e., in that part of the A-band free of interdigitating native thin filaments but not in the center of the A- band which lacks cross-bridges. In the presence of 0.1-0.2 mM ATP, no actin binding occurred. When unlabeled alpha-actinin was added first to myofibrils and then labeled actin was added fluorescence occurred not in a doublet pattern but along the entire length of the myofibril. Tropomyosin did not bind to myofibrils unless the existing tropomyosin was first removed, in which case it added to the thin filaments in the l-band. Tropomyosin did bind, however, to the exogenously added tropomyosin-free actin that localizes as a doublet in the A-band. These results indicate that the alpha-actinin present in Z-bands of myofibrils is fully complexed with actin, but can bind exogenous alpha- actinin and, if actin is added subsequently, the exogenous alpha- actinin in the Z-band will bind the newly formed fluorescent actin filaments. Myofibrillar actin filaments did not increase in length when G-actin was present under polymerizing conditions, nor did they bind any added tropomyosin. These observations are discussed in terms of the structure and in vivo assembly of myofibrils.     

Generation of a fluorescently labeled endogenous protein library in living human cells

We present a protocol to tag proteins expressed from their endogenous chromosomal locations in individual mammalian cells using central dogma tagging. The protocol can be used to build libraries of cell clones, each expressing one endogenous protein tagged with a fluorophore such as the yellow fluorescent protein. Each round of library generation produces 100-200 cell clones and takes about 1 month. The protocol integrates procedures for high-throughput single-cell cloning using flow cytometry, high-throughput cDNA generation and 3' rapid amplification of cDNA ends, semi-automatic protein localization screening using fluorescent microscopy and freezing cells in 96-well format.     

Detection of protein oxidation in rat-1 fibroblasts by fluorescently labeled tyramine.

Oxidative damage to proteins has been postulated as a major cause of various degenerative diseases including the loss of functional capacity during aging. A prominent target for oxidation by reactive oxygen species (ROS) is the tyrosine residue. Here we present a highly sensitive method for the detection of tyrosyl radical formation in cells. The method is based on the fluorescein-labeled tyrosine analogue, tyramine, which upon oxidation may couple to proteins carrying a tyrosyl radical. Coupling of the probe (denoted TyrFluo) to standard proteins could be induced by generating ROS with horseradish peroxidase/hydrogen peroxide, SIN-1 or with peroxides (cumene or hydrogen peroxide) in combination with a transition metal. TyrFluo added to rat-1 fibroblasts remained outside the cell, whereas the acetylated form (acetylTyrFluo) was membrane-permeable and accumulated in the cell. Exposure of the cells to oxidative stress in the presence of either TyrFluo or acetylTyrFluo gave a cellular labeling characteristic for each probe. Western blot analysis confirmed that each probe labeled a specific set of proteins. This new method for the detection of ROS-induced oxidation of proteins may mimic the tendency of oxidized proteins to form dityrosine bonds.     

Tracking expression and subcellular localization of RNA and protein species using high-throughput single cell imaging flow cytometry

We report a high-throughput application of multispectral imaging flow cytometry (MIFC) for analyzing the expression and localization of both RNA and protein molecules in a heterogeneous population of cells. The approach was developed using polyadenylated nuclear (PAN) RNA, an abundant, noncoding RNA expressed by Kaposi's sarcoma-associated herpesvirus (KSHV) during the lytic phase of infection. High levels of PAN RNA are, in part, dependent on its interaction with poly(A)-binding protein C1 (PABPC1), which relocalizes from the cytoplasm to the nucleus of lytically infected cells. We quantitatively tracked the cytoplasmic to nuclear translocation of PABPC1 and examined how this translocation relates to the expression and localization of viral RNA and protein molecules in KSHV-infected cells. This high-throughput approach will be useful for other systems in which changes in subcellular localization of RNA and protein molecules need to be monitored simultaneously.     

Localization of fluorescently labeled structures in frozen-hydrated samples using integrated light electron microscopy

Correlative light and electron microscopy is an increasingly popular technique to study complex biological systems at various levels of resolution. Fluorescence microscopy can be employed to scan large areas to localize regions of interest which are then analyzed by electron microscopy to obtain morphological and structural information from a selected field of view at nm-scale resolution. Previously, an integrated approach to room temperature correlative microscopy was described. Combined use of light and electron microscopy within one instrument greatly simplifies sample handling, avoids cumbersome experimental overheads, simplifies navigation between the two modalities, and improves the success rate of image correlation. Here, an integrated approach for correlative microscopy under cryogenic conditions is presented. Its advantages over the room temperature approach include safeguarding the native hydrated state of the biological specimen, preservation of the fluorescence signal without risk of quenching due to heavy atom stains, and reduced photo bleaching. The potential of cryo integrated light and electron microscopy is demonstrated for the detection of viable bacteria, the study of in vitro polymerized microtubules, the localization of mitochondria in mouse embryonic fibroblasts, and for a search into virus-induced intracellular membrane modifications within mammalian cells.     

Synthesis of rhodamine 6G-based compounds for the ATRP synthesis of fluorescently labeled biocompatible polymers

Facile derivatization of rhodamine 6G in the 2' position by direct reaction with secondary amines is reported. If the secondary amine contains a hydroxy group, the hydroxyl-functional intermediate can be readily esterified to give either fluorescent initiators for atom transfer radical polymerization (ATRP) or a fluorescent methacrylic comonomer. In contrast to rhodamine dyes functionalized using primary amines, which are only fluorescent at low pH, these compounds are highly fluorescent at physiological pH. These new compounds were subsequently used to prepare a range of fluorescently labeled biocompatible polymers based on the biomimetic monomer, 2-(methacryloyloxy)ethyl phosphorylcholine (MPC), for biomedical studies.     

Characterization and imaging of A6 epithelial cell clones expressing fluorescently labeled ENaC subunits

A6 model renal epithelialcells were stably transfected with enhanced green fluorescent protein(EGFP)-tagged - or -subunits of the epithelial Na⁺channel (ENaC). Transfected RNA and proteins were both expressed in lowabundance, similar to the endogenous levels of ENaC in native cells. Inliving cells, laser scanning confocal microscopy revealed apredominately subapical distribution of EGFP-labeled subunits,suggesting a readily accessible pool of subunits available toparticipate in Na⁺ transport. The basal level ofNa⁺ transport in the clonal lines was enhanced two- tofourfold relative to the parent line. Natriferic responses to insulinor aldosterone were similar in magnitude to the parent line, whileforskolin-stimulated Na⁺ transport was 64% greater thancontrol in both the - and -transfected lines. In response toforskolin, EGFP-labeled channel subunits traffic to the apicalmembrane. These data suggest that channel regulators, not the channelper se, form the rate-limiting step in response to insulin oraldosterone stimulation, while the number of channel subunits isimportant for basal as well as cAMP-stimulated Na⁺ transport.

     

Determination of the stoichiometry of protein complexes using liquid chromatography with fluorescence and mass spectrometric detection of fluorescently labeled proteolytic peptides

A method for the determination of the stoichiometry of protein complexes has been developed, which is based on proteolytic digestion of the complex, labeling with a fluorescent reagent, specific for amino or sulfhydryl groups, and separation by liquid chromatography with fluorescence and mass spectrometric detection. The intensity of the fluorescence signal of the labeled peptides resulting from different proteins is directly proportional to the stoichiometry of these proteins in the complex. The performance of the method was evaluated with standard peptides and proteins to ensure that accurate molar ratios can be obtained from the fluorescence chromatogram. Standard deviations of the measured molar ratio from the expected molar ratio were below 10% for both peptides and proteins. The method was finally employed for the determination of the stoichiometry of the 1:1 complex of sFc gamma RIII and hFc1. Using the described methodology, a stoichiometry of 1:1.1 was measured, which agrees well with a 1:1 complex.