Evaluation of few-cycle laser pulses for label-free metabolic imaging on live cells

Is it possible to assess label-free live cell metabolic imaging during early oocyte and embryo development? Label-free metabolic imaging can be systematically used during early development, showing no differences between controls and illuminated oocytes and embryos in terms of early development, blastocyst formation, and embryo outgrowth. Non-invasive methods that are reliable to assess oocyte and embryo quality are a significant aim for ARTs. Changes in metabolic activity could lead to cell death or altered early development and low implantation potential. This could potentially be predicted by incorporating non-invasive measurements of metabolism. Metabolic imaging has been investigated through complex methodologies; however, scientific evidence for its utility during early oocyte and embryo development requires further investigation to assess potential translation in clinical settings. Measurements of metabolic activity could be a useful tool, as the autofluorescence of molecules such as nicotinamide adenine dinucleotide phosphate hydrogen (NAD(P)H) and flavin adenine dinucleotide (FAD) are a straightforward representation of mitochondrial function. Female mice (n = 15) and super-ovulated female mice (n = 30) were used to produce oocytes and embryos, respectively. Oocytes and in-vivo produced embryos were divided into the control group, sham control group, and illuminated group. Illuminated samples were assessed for both NAD(P)H and FAD levels in oocytes and NAD(P)H levels during early embryo development every 3 h using arbitrary units of autofluorescence (AU). Produced blastocysts were assessed for total cell and inner-cell-mass (ICM) number (by immunostaining for Oct4) and embryo outgrowth assays. Furthermore, safety live birth studies were also conducted. F1 (C57BL6/CBA) mouse strain was used. NAD(P)H and FAD autofluorescence levels were measured during oocyte and embryo development using confocal microscopy (Olympus FV1200). A confocal Z-stacking function was used to record 15 focal planes, using a 20×/0.95 NA air objective of the entire oocytes and embryos and opening the confocal pinhole system completely. Images were then collected and analysed using FIJI software (version: 2.0.0-rc-69/1.52n; ImageJ). Developmental rates, blastocyst cell numbers, outgrowth rates (for 4 days post blastocyst formation), and live birth rates were assessed. Oocyte IVM and embryo culture experiments showed no significant differences in developmental rates between study groups (P > 0.05). Similarly, the total number of cells from blastocysts (control: 82.9 ± 5.6; sham: 76.5 ± 3.3; Illuminated: 77.1 ± 4.2; ± SEM) and ICM cells (control: 10.8 ± 1.3; sham: 9.4 ± 0.7; Illuminated: 11.9 ± 0.8; ± SEM) did not differ between groups (P > 0.05). Outgrowth assays of the study groups presented similar outgrowth areas during Days 5-8 (post) blastocyst development (P > 0.05). Illumination of oocytes demonstrated a significant increase in metabolic activity during IVM, measured by the optical redox ratio (ORR: FAD/NAD(P)H + FAD; P < 0.001). Illumination of embryos demonstrated significantly different NAD(P)H activity levels during embryo development, particularly between the two-cell stage (987.1 ± 36.2 AU), morula stage (1226.0 ± 31.5 AU) and blastocyst stage (649 ± 42.9 AU; ± SEM; P < 0.05). Additionally, embryos that did not form blastocysts also presented significantly decreased NAD(P)H activity levels at the two-cell stage (normal development: 987.1 ± 36.2; no blastocyst: 726.9 ± 121.7 AU; P < 0.05) to the morula stage (normal development: 1226.0 ± 31.5; no blastocyst: 886.0 ± 150.4 AU; P < 0.05) when compared with normally developing embryos. Our study indicated that metabolic imaging during early oocyte and embryo development presents no negative effects on developmental rates, blastocyst quality, and embryo outgrowths. Subsequently, live birth rates and offspring health showed no differences between controls and illuminated embryos at the blastocyst stage. Current results provide significant useful information about metabolic activity during live cell imaging as a potential method for timelapse metabolic imaging. N/A. The study was conducted using a mouse model and focused on early oocyte and embryo development, embryo outgrowths, live birth, and early offspring health. Thus, further studies of long-term offspring health are required to fully assess safety and to further validate potential wider applications. Validation in ageing models is also required to assess potential applications for embryo selection. Measurements of metabolic activity could be applied to determine oocyte and embryo metabolic activity using a variety of microscopy technology with low energy doses as described in this study. Further applications could link the use of metabolic imaging with timelapse technology and artificial intelligence applications to monitor culture conditions. This study was funded in part by a research/educational grant from Ferring Pharmaceuticals, awarded from the Fertility Society of Australia and New Zealand (FSANZ). Funding was also provided in part by the Education Program in Reproduction and Development (EPRD), Department of Obstetrics and Gynaecology, Monash University. F.H. and M.H.-T. have applied for a patent in the topic of metabolic imaging. R.B.G. declares speakers' fees from Gedeon Richter and Ferring. The other authors have nothing to declare.

BackgroundImmunotherapy is revolutionizing cancer treatment from conventional radiotherapies and chemotherapies to immune checkpoint inhibitors which use patients’ immune system to recognize and attack cancer cells. Despite the huge clinical success...

<h3>Background</h3> Intravital multiphoton autofluorescence microscopy provides <i>in vivo</i>, label free, single cell imaging of metabolic changes. These metabolic changes are quantified via the metabolic coenzymes NAD(P)H and FAD which are...

Site-specific fluorescent labeling of proteins inside live mammalian cells has been achieved by employing Streptolysin O, a bacterial toxin which forms temporary pores in the membrane and allows delivery of virtually any fluorescent probes, ranging from labeled IgG’s to small ligands, with high efficiency (>85% of cells). The whole process, including recovery, takes 30 min, and the cell is ready to be imaged immediately. A variety of cell viability tests were performed after treatment with SLO to ensure that the cells have intact membranes, are able to divide, respond normally to signaling molecules, and maintains healthy organelle morphology. When combined with Oxyrase, a cell-friendly photostabilizer, a ~20x improvement in fluorescence photostability is achieved. By adding in glutathione, fluorophores are made to blink, enabling super-resolution fluorescence with 20–30 nm resolution over a long time (~30 min) under continuous illumination. Example applications in conventional and super-resolution imaging of native and transfected cells include p65 signal transduction activation, single molecule tracking of kinesin, and specific labeling of a series of nuclear and cytoplasmic protein complexes.DOI: http://dx.doi.org/10.7554/eLife.20378.001

Aggregation-Induced Fluorogens in Bio-Detection, Tumor Imaging, and Therapy: A Review

P55 Selective turn-on fluorescent probes for detection of hydrogen sulfide in living cells

The Promise of Advanced Imaging Techniques for the Detection of Hepatitis C Virus Antigens in the Infected Liver

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.

Tissues of many marine invertebrates of class Anthozoa contain intensely fluorescent or brightly colored pigments. These pigments belong to a family of photoactive proteins closely related to Green Fluorescent Protein (GFP), and their emissions range from blue to red wavelengths. The great diversity of these pigments has only recently been realized. To investigate the role of these proteins in corals, we have performed an in vivo fluorescent pigment (FP) spectral and cellular distribution analyses in live coral cells using single and multi-photon laser scanning imaging and microspectroscopy. These analyses revealed that even single color corals contain spectroscopically heterogeneous pigment mixtures, with 2-5 major color types in the same area of tissue. They were typically arranged in step-wise light emission energy gradients (e.g. blue, green, yellow, red). The successive overlapping emission-excitation spectral profiles of differently colored FPs suggested that they were suited for sequential energy coupling. Traces of red FPs (emission = 570-660 nm) were present, even in non-red corals. We confirmed that radiative energy transfer could occur between separate granules of blue and green FPs and that energy transfer was inversely proportional to the square of the distance between them. Multi-photon micro-spectrofluorometric analysis gave significantly improved spectral resolution by restricting FP excitation to a single point in the focal plane of the sample. Pigment heterogeneity at small scales within granules suggested that fluorescence resonance energy transfer (FRET) might be occurring, and we confirmed that this was the case. Thus, energy transfer can take place both radiatively and by FRET, probably functioning in photoprotection by dissipation of excessive solar radiation.

Genetically encoded temperature indicators (GETIs) allow for real-time measurement of subcellular temperature dynamics in live cells. However, GETIs have suffered from poor temperature sensitivity, which may not be sufficient to resolve small heat production from a biological process. Here, we develop a highly-sensitive GETI, denoted as ELP-TEMP, comprised of a temperature-responsive elastin-like polypeptide (ELP) fused with a cyan fluorescent protein (FP), mTurquoise2 (mT), and a yellow FP, mVenus (mV), as the donor and acceptor, respectively, of Förster resonance energy transfer (FRET). At elevated temperatures, the ELP moiety in ELP-TEMP undergoes a phase transition leading to an increase in the FRET efficiency. In HeLa cells, ELP-TEMP responded to the temperature from 33 to 40 °C with a maximum temperature sensitivity of 45.1 ± 8.1%/°C, which was the highest ever temperature sensitivity among hitherto-developed fluorescent nanothermometers. Although ELP-TEMP showed sensitivity not only to temperature but also to macromolecular crowding and self-concentration, we were able to correct the output of ELP-TEMP to achieve accurate temperature measurements at a subcellular resolution. We successfully applied ELP-TEMP to accurately measure temperature changes in cells induced by a local heat spot, even if the temperature difference was as small as < 1 °C, and to visualize heat production from stimulated Ca2+ influx in live HeLa cells induced by a chemical stimulation. Furthermore, we investigated temperatures in the nucleus and cytoplasm of live HeLa cells and found that their temperatures were almost the same within the temperature resolution of our measurement. Our study would contribute to better understanding of cellular temperature dynamics, and ELP-TEMP would be a useful GETI for the investigation of cell thermobiology.

A series of new fluorescent probes were developed to carry out live cell super-resolution imaging with low STED laser power or suitable STORM working conditions. And STED-FLIM imaging of microtubules labeled with ATTO647N inside HeLa cells and the mitosis process was obtained, which provides new insight into the cell structure and functions.Finally, stochastic optical reconstruction microscopy (STORM) super-resolution imaging of mitochondrial membrane in live HeLa cells was obtained by the implementation of new fluorescent probes, improved imaging system and optimized single molecule localization algorithm. This provided an important tool and strategy for studying dynamic events and complex functions in living cells.

Few naturally-occurring plasmids are maintained in mammalian cells. Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus (EBV) and Kaposi's Sarcoma-associated herpesvirus (KSHV), which cause multiple human malignancies (1-3). These two genomes are replicated in a licensed manner, each using a single viral protein and cellular replication machinery, and are passed to daughter cells during cell division despite their lacking traditional centromeres (4-8). Much work has been done to characterize the replications of these plasmid genomes using methods such as Southern blotting and fluorescence in situ hybridization (FISH). These methods are limited, though. Quantitative PCR and Southern blots provide information about the average number of plasmids per cell in a population of cells. FISH is a single-cell assay that reveals both the average number and the distribution of plasmids per cell in the population of cells but is static, allowing no information about the parent or progeny of the examined cell. Here, we describe a method for visualizing plasmids in live cells. This method is based on the binding of a fluorescently tagged lactose repressor protein to multiple sites in the plasmid of interest (9). The DNA of interest is engineered to include approximately 250 tandem repeats of the lactose operator (LacO) sequence. LacO is specifically bound by the lactose repressor protein (LacI), which can be fused to a fluorescent protein. The fusion protein can either be expressed from the engineered plasmid or introduced by a retroviral vector. In this way, the DNA molecules are fluorescently tagged and therefore become visible via fluorescence microscopy. The fusion protein is blocked from binding the plasmid DNA by culturing cells in the presence of IPTG until the plasmids are ready to be viewed. This system allows the plasmids to be monitored in living cells through several generations, revealing properties of their synthesis and partitioning to daughter cells. Ideal cells are adherent, easily transfected, and have large nuclei. This technique has been used to determine that 84% of EBV-derived plasmids are synthesized each generation and 88% of the newly synthesized plasmids partition faithfully to daughter cells in HeLa cells. Pairs of these EBV plasmids were seen to be tethered to or associated with sister chromatids after their synthesis in S-phase until they were seen to separate as the sister chromatids separated in Anaphase(10). The method is currently being used to study replication of KSHV genomes in HeLa cells and SLK cells. HeLa cells are immortalized human epithelial cells, and SLK cells are immortalized human endothelial cells. Though SLK cells were originally derived from a KSHV lesion, neither the HeLa nor SLK cell line naturally harbors KSHV genomes(11). In addition to studying viral replication, this visualization technique can be used to investigate the effects of the addition, removal, or mutation of various DNA sequence elements on synthesis, localization, and partitioning of other recombinant plasmid DNAs.

Fluorescence lifetime-resolved imaging microscopy (FLIM) has been used to monitor the enzymatic activity of a proteolytic enzyme, Membrane Type 1 Matrix Metalloproteinase (MT1-MMP), with a recently developed FRET-based biosensor in vitro and in live HeLa and HT1080 cells. MT1-MMP is a collagenaise that is involved in the destruction of extra-cellular matrix (ECM) proteins, as well as in various cellular functions including migration. The increased expression of MT1-MMP has been positively correlated with the invasive potential of tumor cells. However, the precise spatiotemporal activation patterns of MT1-MMP in live cells are still not well-established. The activity of MT1-MMP was examined with our biosensor in live cells. Imaging of live cells was performed with full-field frequency-domain FLIM. Image analysis was carried out both with polar plots and phase differential enhancement. Phase differential enhancement, which is similar to phase suppression, is shown to facilitate the differentiation between different conformations of the MT1-MMP biosensor in live cells when the lifetime differences are small. FLIM carried out in differential enhancement or phase suppression modes, requires only two acquired phase images, and permits rapid imaging of the activity of MT1-MMP in live cells.

Septins are filamentous GTPases that associate with cell membranes and the cytoskeleton and play essential roles in cell division and cellular morphogenesis. Septins are implicated in many human diseases including cancer and neuropathies. Small molecules that reversibly perturb septin organization and function would be valuable tools for dissecting septin functions and could be used for therapeutic treatment of septin-related diseases. Forchlorfenuron (FCF) is a plant cytokinin previously shown to disrupt septin localization in budding yeast. However, it is unknown whether FCF directly targets septins and whether it affects septin organization and functions in mammalian cells. Here, we show that FCF alters septin assembly in vitro without affecting either actin or tubulin polymerization. In live mammalian cells, FCF dampens septin dynamics and induces the assembly of abnormally large septin structures. FCF has a low level of cytotoxicity, and these effects are reversed upon FCF washout. Significantly, FCF treatment induces mitotic and cell migration defects that phenocopy the effects of septin depletion by small interfering RNA. We conclude that FCF is a promising tool to study mammalian septin organization and functions.

Editor's evaluation: Autofluorescence imaging permits label-free cell type assignment and reveals the dynamic formation of airway secretory cell associated antigen passages (SAPs)