Fluorescence Lifetime Imaging of Molecular Rotors in Living Cells

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular events. Viscosity is one of the key parameters affecting the diffusion of molecules and proteins, and changes in viscosity have been linked to disease and malfunction at the cellular level.^1-3 While methods to measure the bulk viscosity are well developed, imaging microviscosity remains a challenge. Viscosity maps of microscopic objects, such as single cells, have until recently been hard to obtain. Mapping viscosity with fluorescence techniques is advantageous because, similar to other optical techniques, it is minimally invasive, non-destructive and can be applied to living cells and tissues.Fluorescent molecular rotors exhibit fluorescence lifetimes and quantum yields which are a function of the viscosity of their microenvironment.^4,5 Intramolecular twisting or rotation leads to non-radiative decay from the excited state back to the ground state. A viscous environment slows this rotation or twisting, restricting access to this non-radiative decay pathway. This leads to an increase in the fluorescence quantum yield and the fluorescence lifetime. Fluorescence Lifetime Imaging (FLIM) of modified hydrophobic BODIPY dyes that act as fluorescent molecular rotors show that the fluorescence lifetime of these probes is a function of the microviscosity of their environment.^6-8 A logarithmic plot of the fluorescence lifetime versus the solvent viscosity yields a straight line that obeys the Förster Hoffman equation.⁹ This plot also serves as a calibration graph to convert fluorescence lifetime into viscosity.Following incubation of living cells with the modified BODIPY fluorescent molecular rotor, a punctate dye distribution is observed in the fluorescence images. The viscosity value obtained in the puncta in live cells is around 100 times higher than that of water and of cellular cytoplasm.^6,7 Time-resolved fluorescence anisotropy measurements yield rotational correlation times in agreement with these large microviscosity values. Mapping the fluorescence lifetime is independent of the fluorescence intensity, and thus allows the separation of probe concentration and viscosity effects. In summary, we have developed a practical and versatile approach to map the microviscosity in cells based on FLIM of fluorescent molecular rotors.     

Preparation of Living Isolated Vertebrate Photoreceptor Cells for Fluorescence Imaging

In the vertebrate retina, phototransduction, the conversion of light to an electrical signal, is carried out by the rod and cone photoreceptor cells^1-4. Rod photoreceptors are responsible for vision in dim light, cones in bright light. Phototransduction takes place in the outer segment of the photoreceptor cell, a specialized compartment that contains a high concentration of visual pigment, the primary light detector. The visual pigment is composed of a chromophore, 11-cis retinal, attached to a protein, opsin. A photon absorbed by the visual pigment isomerizes the chromophore from 11-cis to all-trans. This photoisomerization brings about a conformational change in the visual pigment that initiates a cascade of reactions culminating in a change in membrane potential, and bringing about the transduction of the light stimulus to an electrical signal. The recovery of the cell from light stimulation involves the deactivation of the intermediates activated by light, and the reestablishment of the membrane potential. Ca²⁺ modulates the activity of several of the enzymes involved in phototransduction, and its concentration is reduced upon light stimulation. In this way, Ca²⁺ plays an important role in the recovery of the cell from light stimulation and its adaptation to background light.Another essential part of the recovery process is the regeneration of the visual pigment that has been destroyed during light-detection by the photoisomerization of its 11-cis chromophore to all-trans^5-7. This regeneration begins with the release of all-trans retinal by the photoactivated pigment, leaving behind the apo-protein opsin. The released all-trans retinal is rapidly reduced in a reaction utilizing NADPH to all- trans retinol, and opsin combines with fresh 11-cis retinal brought into the outer segment to reform the visual pigment. All-trans retinol is then transferred out of the outer segment and into neighboring cells by the specialized carrier Interphotoreceptor Retinoid Binding Protein (IRBP).Fluorescence imaging of single photoreceptor cells can be used to study their physiology and cell biology. Ca²⁺-sensitive fluorescent dyes can be used to examine in detail the interplay between outer segment Ca²⁺ changes and response to light^8-12 as well as the role of inner segment Ca²⁺ stores in Ca²⁺ homeostasis^13,14. Fluorescent dyes can also be used for measuring Mg²⁺ concentration¹⁵, pH, and as tracers of aqueous and membrane compartments¹⁶. Finally, the intrinsic fluorescence of all-trans retinol (vitamin A) can be used to monitor the kinetics of its formation and removal in single photoreceptor cells^17-19.Download video file.^{(70M, mov)}     

Intravital Fluorescence Excitation in Whole-Animal Optical Imaging

Whole-animal fluorescence imaging with recombinant or fluorescently-tagged pathogens or cells enables real-time analysis of disease progression and treatment response in live animals. Tissue absorption limits penetration of fluorescence excitation light, particularly in the visible wavelength range, resulting in reduced sensitivity to deep targets. Here, we demonstrate the use of an optical fiber bundle to deliver light into the mouse lung to excite fluorescent bacteria, circumventing tissue absorption of excitation light in whole-animal imaging. We present the use of this technology to improve detection of recombinant reporter strains of tdTomato-expressing Mycobacterium bovis BCG (Bacillus Calmette Guerin) bacteria in the mouse lung. A microendoscope was integrated into a whole-animal fluorescence imager to enable intravital excitation in the mouse lung with whole-animal detection. Using this technique, the threshold of detection was measured as 10³ colony forming units (CFU) during pulmonary infection. In comparison, the threshold of detection for whole-animal fluorescence imaging using standard epi-illumination was greater than 10⁶ CFU.     

pH敏感的荧光探针及在活细胞中的应用

细胞内的pH是细胞内多种酶活性和生理活动的重要调节因素,准确、动态的监测细胞内pH变化对研究细胞内的活动至关重要。一些荧光小分子可以感应pH的变化,同时具有较高的灵敏度和特异性,对细胞损伤较小且标记操作简单,已逐渐发展成为一种监测细胞内pH变化的有效方法。本文主要介绍目前常用pH敏感的荧光探针及其在活细胞研究中的进展。     

Detecting Subtle Plasma Membrane Perturbation in Living Cells Using Second Harmonic Generation Imaging

The requirement of center asymmetry for the creation of second harmonic generation (SHG) signals makes it an attractive technique for visualizing changes in interfacial layers such as the plasma membrane of biological cells. In this article, we explore the use of lipophilic SHG probes to detect minute perturbations in the plasma membrane. Three candidate probes, Di-4-ANEPPDHQ (Di-4), FM4-64, and all-trans-retinol, were evaluated for SHG effectiveness in Jurkat cells. Di-4 proved superior with both strong SHG signal and limited bleaching artifacts. To test whether rapid changes in membrane symmetry could be detected using SHG, we exposed cells to nanosecond-pulsed electric fields, which are believed to cause formation of nanopores in the plasma membrane. Upon nanosecond-pulsed electric fields exposure, we observed an instantaneous drop of ∼50% in SHG signal from the anodic pole of the cell. When compared to the simultaneously acquired fluorescence signals, it appears that the signal change was not due to the probe diffusing out of the membrane or changes in membrane potential or fluidity. We hypothesize that this loss in SHG signal is due to disruption in the interfacial nature of the membrane. The results show that SHG imaging has great potential as a tool for measuring rapid and subtle plasma membrane disturbance in living cells.     

Fluorescence Fluctuation Spectroscopy of mCherry in Living Cells

The red fluorescent protein mCherry is of considerable interest for fluorescence fluctuation spectroscopy (FFS), because the wide separation in color between mCherry and green fluorescent protein provides excellent conditions for identifying protein interactions inside cells. This two-photon study reveals that mCherry exists in more than a single brightness state. Unbiased analysis of the data needs to account for the presence of multiple states. We introduce a two-state model that successfully describes the brightness and fluctuation amplitude of mCherry. The properties of the two states are characterized by FFS and fluorescence lifetime experiments. No interconversion between the two states was observed over the experimentally probed timescales. The effect of fluorescence resonance energy transfer between enhanced green fluorescent protein (EGFP) and mCherry is incorporated into the two-state model to describe protein hetero-oligomerization. The model is verified by comparing the predicted and measured brightness and fluctuation amplitude of several fusion proteins that contain mCherry and EGFP. In addition, hetero-fluorescence resonance energy transfer between mCherry molecules in different states is detected, but its influence on FFS parameters is small enough to be negligible. Finally, the two-state model is applied to study protein oligomerization in living cells. We demonstrate that the model successfully describes the homodimerization of nuclear receptors. In addition, we resolved a mixture of interacting and noninteracting proteins labeled with EGFP and mCherry. These results provide the foundation for quantitative applications of mCherry in FFS studies.     

Detecting Subtle Plasma Membrane Perturbation in Living Cells Using Second Harmonic Generation Imaging

The requirement of center asymmetry for the creation of second harmonic generation (SHG) signals makes it an attractive technique for visualizing changes in interfacial layers such as the plasma membrane of biological cells. In this article, we explore the use of lipophilic SHG probes to detect minute perturbations in the plasma membrane. Three candidate probes, Di-4-ANEPPDHQ (Di-4), FM4-64, and all-trans-retinol, were evaluated for SHG effectiveness in Jurkat cells. Di-4 proved superior with both strong SHG signal and limited bleaching artifacts. To test whether rapid changes in membrane symmetry could be detected using SHG, we exposed cells to nanosecond-pulsed electric fields, which are believed to cause formation of nanopores in the plasma membrane. Upon nanosecond-pulsed electric fields exposure, we observed an instantaneous drop of ∼50% in SHG signal from the anodic pole of the cell. When compared to the simultaneously acquired fluorescence signals, it appears that the signal change was not due to the probe diffusing out of the membrane or changes in membrane potential or fluidity. We hypothesize that this loss in SHG signal is due to disruption in the interfacial nature of the membrane. The results show that SHG imaging has great potential as a tool for measuring rapid and subtle plasma membrane disturbance in living cells.As the epicenter for many cellular functions, understanding the dynamics of the plasma membrane is important to monitoring biological phenomena. External forces acting upon the plasma membrane (e.g., electric, mechanical) have been shown to cause rapid disturbances, often resulting in dramatic changes in cell physiology (1–3). To understand this interaction, a minimally invasive, highly sensitive imaging technique that enables monitoring the structure of the plasma membrane is needed. Lipophilic dyes, which embed themselves into lipid membranes, are sensitive to the surrounding electric field and, therefore, report changes in membrane fluidity as well as voltage due to the capacitive nature of the membranes (4,5). This sensitivity is typically detected as a shift in the fluorescence emission spectrum. Localization of the fluorescence signal to only the plasma membrane is difficult because the probes also label internal membrane structures. Thus, to overcome this lack of spatial selectivity, second harmonic generation (SHG) has been used as an alternative to fluorescence for membrane imaging (6,7).In SHG, a second-order nonlinear polarization is induced by electronic disruption of a probe molecule from the electromagnetic field of the incident laser beam. This polarization generates oscillating dipole moments that reradiate light at twice the energy of the excitation beam. The induction of this dipole is sensitive to the static electric field surrounding the probe and the steady-state molecular polarization of the probe molecule. These properties make SHG probes useful for monitoring changes in biological membranes.First, as the voltage potential across the membrane changes, the static electric field around the probe also shifts, making the probe sensitive to these variations (7). Several SHG probes have, therefore, been employed to monitor plasma membrane potential (7,8).Second, because the dipole is affected by the steady-state molecular polarization of the probe itself, a SHG signal is only produced in materials that lack a center of inversion symmetry. In the centrosymmetric case, any emitted radiation is cancelled out by destructive interference. The properties of an interfacial environment, such as a cellular plasma membrane, not only provide the necessary asymmetry, but cause the polarized lipophilic dyes to be aligned in respect to the interface, instead of being randomly distributed as they would in a bulk environment. This alignment allows the generation of a coherent SHG signal from the plasma membrane while the rest of the cell remains nearly signal-free (6,7).We investigated whether the alignment sensitivity of the SHG response could be used to detect minute changes in the organization of the plasma membrane. Jurkat clone E6-1 human T-lymphocytes with a spherical morphology were selected for optimum signal clarity and cultured as directed by American Type Culture Collection (ATCC, Manassas, VA) with 1 I.U./mL penicillin and 0.1 μg/mL streptomycin. Cells were added to 35-mm poly-L-lysine-coated glass-bottomed dishes (MatTek, Ashland, MA) and incubated for 1 h in growth media to allow adherence. Before loading, the cells were rinsed with a buffer consisting of 135 mM NaCl, 5 mM KCl, 2 mM MgCl2, 10 mM HEPES, 10 mM glucose, 2 mM CaCl2, pH 7.4, 290–310 mOsm. SHG probes, Di-4-ANEPPDHQ (Di-4) (5 μM final concentration), FM4-64 (15 μM) or ATR (100 μM, 1 mg/mL BSA) were added to the buffer solution and incubated for 1 h. Cellular imaging was performed in the labeling buffer to limit diffusion of the probe molecules out of the cell membranes.A Ti:sapphire oscillator at 980 nm (Coherent Chameleon, 130 fs, 80 MHz, ∼15 mW at the sample; Coherent Laser, Santa Clara, CA) was coupled through the scan head of a modified model No. TCS SP5 II (Leica Geosystems, Norcross, GA) for SHG and multiphoton-excited fluorescence imaging (40×, water, 1.1 NA) using resonant scanning. SHG signal was collected in transmission by a photomultiplier tube after 680-nm shortpass and 485/25-nm bandpass filters; simultaneous fluorescence signal was collected in the epi-direction by two non-descanned photomultiplier tubes with 540/60-nm and 650/60-nm bandpass filters.Three SHG probes previously used to monitor voltage or membrane order in living cells were tested (8–10). Although ATR is reported to be effective in monitoring membrane voltage, we obtained nearly no SHG signal, despite successful loading as indicated by the fluorescence signal (Fig. 1). When FM4-64 and Di-4 were loaded to similar fluorescence intensities, nearly equivalent SHG signal was collected. Di-4 did appear to have a greater internalization of the dye. However, after the first frame, the FM4-64 signal dropped considerably (Fig. 1 b), an observation reported as a membrane voltage-independent bleaching effect (8). This drop in signal recovered after excitation was blocked for several seconds, but quantification of the response was difficult. Di-4 did not suffer as dramatic a drop in signal upon excitation, and still had sufficient SHG signal/noise after several seconds, so it was used in all further experiments.Open in a separate windowFigure 1(a) Fluorescence (top) and SHG (bottom) images for the three probes. (b) Signal/noise for the fluorescence and SHG for the initial frame and shortly after beginning acquisition. Error bars represent the mean ± SE (n = 10). Scale bar is 10 μm.To test whether Di-4 would report a rapid change in membrane organization, we applied a single nanosecond-duration pulsed electric field (nsPEF) to the labeled cell. These ultrabrief, high-intensity (MV/m) pulses differ from longer (μs-ms), lower-intensity (kV/m) pulses traditionally associated with electroporation in induced cellular response (3,11,12). Through selective uptake of small ions (Ca²⁺, Ti⁺) with limited uptake of propidium iodide, nsPEF have been previously postulated to cause nanopores (<2 nm diameter) in the plasma membrane. In contrast with a previous study observing poration resulting from traditional electroporation (13), the brevity of this apparently novel cellular insult allows for the decoupling of the mechanical effects of the pulse on the membrane from the electrical effects of the pulse itself. A single pulse, generated by a custom pulse generator, was delivered to the cells using a pair of 125-μm diameter tungsten electrodes, separated edge-to-edge by 150 μm, as previously described in Ibey et al. (14). For maximum visualization of changes in the SHG signal, a half-wave plate was placed before the scan head to align the polarization of the laser such that the brightest signals from the plasma membrane were at the poles facing the electrodes.The Di-4 SHG signal in response to a single 16.7 kV/cm, 600-ns nsPEF is shown in Fig. 2. Before the pulse, the intensity of the SHG signal is high at each of these poles. Immediately after the pulse, the SHG intensity drops by ∼50% on the side of the cell facing the anodic electrode, whereas little intensity is lost at the other pole. This response is plotted in Fig. 2 (pulse applied at 2 s), where it is apparent that the response is near instantaneous with little recovery in signal in the 5 s postexposure. The SHG response matches the previously observed effect of this stimulus, where ion uptake displayed a polar dependence and persisted for a number of minutes (11,12). Images taken 5 min after an nsPEF exposure are also shown in Fig. 2. These images confirm the eventual recovery of the cell and the corresponding return of SHG signal to preexposure levels.Open in a separate windowFigure 2(a) SHG images showing drop in signal on the anodic (or A-pole) of the cell. (b) Time trace of SHG response with the electrical pulse applied at 2 s that shows a near-instantaneous drop in the SHG signal at the anodic pole of the cell. (c) SHG image preexposure, immediately postexposure, and then 5-min postexposure showing recovery of the SHG signal.To decouple membrane disturbance from environmental changes around the membrane, we compared the SHG response to the simultaneously acquired fluorescence signal. Because fluorescence is not subject to the strict orientation requirement of SHG, the plasma membrane fluorescence signal provides an indication of the membrane fluidity and/or potential. Despite the dramatic shift in SHG intensity on the anodic pole upon the electrical pulse exposure (Fig. 3 a), the fluorescence channels display little response from the equivalent membrane sections with the exception of photobleaching and a slight increase in signal in both emission bands on the anodic side (Fig. 3, b and c). The shading in these graphs represents the mean ± SE for six cells. Although this slight increase may indicate that a small amount of dye is simply diffusing in or out of the membrane upon exposure, the fluorescence response is not as rapid or as lasting as the SHG response. Change in membrane fluidity or voltage can also be quantized using these fluorescence signals and a value known as the generalized polarization (GP) (4),GP=I515−570−I620−680I515−570+I620−680.(1)As with the raw intensity of the individual signals, the GP value for the membrane (Fig. 3 d) shows no significant shift, indicating that the membrane is likely not transitioning between a more raft- and fluidlike state. Thus, it seems likely that the dye was initially aligned in the tightly-packed ordered membrane so that the probes were able to generate a SHG photon. As shown in Fig. 3 e, we postulate that upon electrical pulse exposure, the membrane was disrupted by the formation with nanopores giving the probe molecules the flexibility to disorient within the membrane. The resulting alignment of the probes is more isotropic in nature, thereby limiting the probes probability of producing a SHG photon. The fluorescence signal remained, however, indicating that the probes remained active in the membrane.Open in a separate windowFigure 3(a) Average SHG signal showing the dramatic drop in signal on the anodic pole at the pulse application (2 s). (b and c) Simultaneous TPF signals showing nearly no instantaneous change at the pulse application. (d) GP showing no observable changes in the membrane potential or fluidity after the pulse. (Shaded areas) Fit to the mean ± SE for each trace (n = 6). (e) Conceptualization of the hypothesized membrane disruption underlying the observed change in SHG response.Thus, by taking advantage of the selection criteria of SHG, we were able to successfully use the SHG probe, Di-4, to monitor rapid disruption of the plasma membrane. Because SHG can only be generated when the probes are aligned in the plasma membrane, the SHG signal diminishes significantly upon disruption. The simultaneous collection of the multiphoton-excited fluorescence signal was advantageous in that it demonstrated that the probes did not simply diffuse out of the membrane, did not appear to be energetically disrupted by the electric pulse, and showed that the membrane changes were not simply a change in lipid order. We believe that this technique holds tremendous potential for use in the study of how external stimuli interact with and change the orientation of biological membranes. Such knowledge may allow for further understanding of how manipulation of cells and biological systems can be achieved using external stimuli.     

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases ¹. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression ¹. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer''s disease and multiple sclerosis ¹. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. ² In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.Download video file.^{(134M, mp4)}     

Bimolecular Fluorescence Complementation; Lighting-Up Tau-Tau Interaction in Living Cells

Abnormal tau aggregation is a pathological hallmark of many neurodegenerative disorders and it is becoming apparent that soluble tau aggregates play a key role in neurodegeneration and memory impairment. Despite this pathological importance, there is currently no single method that allows monitoring soluble tau species in living cells. In this regard, we developed a cell-based sensor that visualizes tau self-assembly. By introducing bimolecular fluorescence complementation (BiFC) technique to tau, we were able to achieve spatial and temporal resolution of tau-tau interactions in a range of states, from soluble dimers to large aggregates. Under basal conditions, tau-BiFC cells exhibited little fluorescence intensity, implying that the majority of tau molecules exist as monomers. Upon chemically induced tau hyperphosphorylation, BiFC fluorescence greatly increased, indicating an increased level of tau-tau interactions. As an indicator of tau assembly, our BiFC sensor would be a useful tool for investigating tau pathology.     

Fast Label-Free Cytoskeletal Network Imaging in Living Mammalian Cells

We present a full-field technique that allows label-free cytoskeletal network imaging inside living cells. This noninvasive technique allows monitoring of the cytoskeleton dynamics as well as interactions between the latter and organelles on any timescale. It is based on high-resolution quantitative phase imaging (modified Quadriwave lateral shearing interferometry) and can be directly implemented using any optical microscope without modification. We demonstrate the capability of our setup on fixed and living Chinese hamster ovary cells, showing the cytoskeleton dynamics in lamellipodia during protrusion and mitochondria displacement along the cytoskeletal network. In addition, using the quantitative function of the technique, along with simulation tools, we determined the refractive index of a single tubulin microtubule to be n_tubu = 2.36 ± 0.6 at λ = 527 nm.The cytoskeleton is mainly composed of an actin and tubulin microtubule network, and it has many important roles at the cellular scale (1). It allows the cell to modify its shape, is implied in cell migration and adhesion processes, and is used as a support for organelle displacement inside cells.Optical microscopy is useful for dynamic studies in which the cytoskeletal network is reorganizing quickly. However, due to the poor native interaction between light and this network fluorescence labeling is commonly used to image the cytoskeleton (2). Anisotropic approaches are also used on this kind of structure, as the cytoskeletal filaments may present refractive index anisotropy. Based on this property, polarized microscopy has been used to reveal the cytoskeleton (3). However, this technique is relatively slow compared to the cytoskeleton dynamics and requires perfectly stressless optics and a nondepolarizing sample. Differential interference contrast (DIC) approaches enhance the contrast in unstained cytoskeletal fibers (4) but also require precise light polarization control of both samples and optical components (for example, no plastic elements can be used for standard DIC). Moreover, the image has a gradient shape that induces loss of resolution and makes the images hard to interpret, especially in complex biological environments. Some DIC-based developments have been proposed that would make it possible to retrieve quantitative information from the sample and/or minimize the effects of depolarizing elements (5–8). Nonlinear interactions in second-harmonic generation (SHG) (9) that are sensitive to orientation and anisotropic refractive index are also applied to cytoskeleton imaging. Label-free imaging is thus obtained, but it requires a powerful laser and a scanning approach that may be too slow when fast dynamics need to be studied.Although light interaction with a nonlabeled cytoskeleton is weak, with barely any absorption, there is a signature on the beam that travels through the structure even with nonpolarized illumination/detection. Indeed, as tubulin microtubules and actin filaments are denser than the cytoplasm, their respective refractive indices are also higher (10). This means that the light is slightly delayed by the cytoskeleton, leading to a possible contrast when looking at the phase component of light. In this article, we consider quantitative phase microscopy (QPM) based on quadriwave lateral shearing interferometry (QWLSI) (11). QWLSI makes it possible to image nonlabeled cells with a conventional transillumination microscope equipped with a halogen lamp. We propose a modified version of the QWLSI presented in our previous publication (11) that allows the fast, sensitive, and highly resolved imaging required to reveal cytoskeletal network dynamics in living cells. After discussing the signal/noise ratio (SNR) of our approach, we compare QPM with immunostaining of actin and tubulin microtubules on Chinese hamster ovary (CHO) cells, demonstrating the capability of QPM to visualize the cytoskeleton. Living wild-type (wt) CHO cells are then imaged at a high frame rate (2.5 Hz) to illustrate the spatiotemporal resolution of the technique for cytoskeleton imaging.     

Fast Label-Free Cytoskeletal Network Imaging in Living Mammalian Cells

We present a full-field technique that allows label-free cytoskeletal network imaging inside living cells. This noninvasive technique allows monitoring of the cytoskeleton dynamics as well as interactions between the latter and organelles on any timescale. It is based on high-resolution quantitative phase imaging (modified Quadriwave lateral shearing interferometry) and can be directly implemented using any optical microscope without modification. We demonstrate the capability of our setup on fixed and living Chinese hamster ovary cells, showing the cytoskeleton dynamics in lamellipodia during protrusion and mitochondria displacement along the cytoskeletal network. In addition, using the quantitative function of the technique, along with simulation tools, we determined the refractive index of a single tubulin microtubule to be n_tubu=2.36±0.6$n_{tubu}=2.36\pm 0.6$ at λ=527$\lambda =527$ nm.     

Video-Rate Confocal Microscopy for Single-Molecule Imaging in Live Cells and Superresolution Fluorescence Imaging

There is no confocal microscope optimized for single-molecule imaging in live cells and superresolution fluorescence imaging. By combining the swiftness of the line-scanning method and the high sensitivity of wide-field detection, we have developed a, to our knowledge, novel confocal fluorescence microscope with a good optical-sectioning capability (1.0 μm), fast frame rates (<33 fps), and superior fluorescence detection efficiency. Full compatibility of the microscope with conventional cell-imaging techniques allowed us to do single-molecule imaging with a great ease at arbitrary depths of live cells. With the new microscope, we monitored diffusion motion of fluorescently labeled cAMP receptors of Dictyostelium discoideum at both the basal and apical surfaces and obtained superresolution fluorescence images of microtubules of COS-7 cells at depths in the range 0–85 μm from the surface of a coverglass.     

动物活体内细胞光操控的研究进展

动物活体环境下单细胞的光操控对于研究细胞的结构和功能,细胞与组织之间的相互作用,以及细胞病变机理、血栓形成机制和肿瘤细胞迁移等生物医学问题具有重要意义。2013年,光镊技术首次应用于活体动物内单细胞的捕获和操控,开辟了活体动物内光学操控新领域。本文就该领域涉及的活体操控技术及近来取得的重要研究进展进行概述,简要分析了实现深度组织内细胞操控所遇到的技术瓶颈并讨论了解决方案。     

Analytic 3D Imaging of Mammalian Nucleus at Nanoscale Using Coherent X-Rays and Optical Fluorescence Microscopy

Despite the notable progress that has been made with nano-bio imaging probes, quantitative nanoscale imaging of multistructured specimens such as mammalian cells remains challenging due to their inherent structural complexity. Here, we successfully performed three-dimensional (3D) imaging of mammalian nuclei by combining coherent x-ray diffraction microscopy, explicitly visualizing nuclear substructures at several tens of nanometer resolution, and optical fluorescence microscopy, cross confirming the substructures with immunostaining. This demonstrates the successful application of coherent x-rays to obtain the 3D ultrastructure of mammalian nuclei and establishes a solid route to nanoscale imaging of complex specimens.     

Analytic 3D Imaging of Mammalian Nucleus at Nanoscale Using Coherent X-Rays and Optical Fluorescence Microscopy

Despite the notable progress that has been made with nano-bio imaging probes, quantitative nanoscale imaging of multistructured specimens such as mammalian cells remains challenging due to their inherent structural complexity. Here, we successfully performed three-dimensional (3D) imaging of mammalian nuclei by combining coherent x-ray diffraction microscopy, explicitly visualizing nuclear substructures at several tens of nanometer resolution, and optical fluorescence microscopy, cross confirming the substructures with immunostaining. This demonstrates the successful application of coherent x-rays to obtain the 3D ultrastructure of mammalian nuclei and establishes a solid route to nanoscale imaging of complex specimens.