Probing Deviation of Adhered Membrane Dynamics between Reconstituted Liposome and Cellular System.

The dynamics of cell-cell adhesion are complicated due to complexities in cellular interactions and intra-membrane interactions. In the present work, we have reconstituted a liposome-based model system to mimic the cell-cell adhesion process. Our model liposome system consists of one fluorescein-tagged and one TRITC (tetramethyl-rhodamine isothiocyanate)-tagged liposome, adhered through biotin-neutravidin interaction. We monitored the adhesion process in liposomes using Förster Resonance Energy Transfer (FRET) between fluorescein (donor) and TRITC (acceptor). Occurrence of FRET is confirmed by the decrease in donor lifetime as well as distinct rise time of the acceptor fluorescence. Interestingly, the acceptor's emission exhibits fluctuations in the range of ≈3±1 s. This may be attributed to structural oscillations associated in two adhered liposomes arising from the flexible nature of biotin-neutravidin interaction. We have compared the dynamics in a cell-mimicking liposome system with that in an in vitro live cell system. In the adhered live cell system, we used CPM (7-diethylamino-3-(4-maleimido-phenyl)-4-methylcoumarin, donor) and nile red (acceptor), which are known to stain the membrane of CHO (Chinese Hamster Ovary) cells. The dynamics of the adhered membranes of two live CHO cells were observed through FRET between CPM and nile red. The acceptor fluorescence intensity exhibits an oscillation in the time-scale of ≈1±0.75 s, which is faster compared to the reconstituted liposome system, indicating the contributions and involvement of multiple dynamic protein complexes around the cell membrane. This study offers simple reconstituted model systems to understand the complex membrane dynamics using a FRET-based physical chemistry approach.

Physical chemistry in a single live cell: confocal microscopy.

A live cell is a complex, yet extremely important container. Understanding the dynamics in a selected intracellular component is a challenging task. We have recently made significant progress in this direction using a confocal microscope as a tool. The smallest size of the focused spot in a confocal microscope is ∼0.2 µm (200 nm). This is nearly one hundred times smaller than the size of a live cell. Thus, one can selectively study different intracellular components/organelles in a live cell. In this paper, we discuss how one can image different intracellular components/organelles, record fluorescence spectra and decay at different locations, ascertain local polarity and viscosity, and monitor the dynamics of solvation, proton transfer, red-ox and other phenomena at specified locations/organelles inside a cell. We will highlight how this knowledge enriched us in differentiating between cancer and non-cancer cells, 3D tumor spheroids and towards drug delivery.

Effect of alcohol on the structure of cytochrome C: FCS and molecular dynamics simulations.

Effect of ethanol on the size and structure of a protein cytochrome C (Cyt C) is investigated using fluorescence correlation spectroscopy (FCS) and molecular dynamics (MD) simulations. For FCS studies, Cyt C is covalently labeled with a fluorescent probe, alexa 488. FCS studies indicate that on addition of ethanol, the size of the protein varies non-monotonically. The size of Cyt C increases (i.e., the protein unfolds) on addition of alcohol (ethanol) up to a mole fraction of 0.2 (44.75% v/v) and decreases at higher alcohol concentration. In order to provide a molecular origin of this structural transition, we explore the conformational free energy landscape of Cyt C as a function of radius of gyration (Rg) at different compositions of water-ethanol binary mixture using MD simulations. Cyt C exhibits a minimum at Rg ∼ 13 Å in bulk water (0% alcohol). Upon increasing ethanol concentration, a second minimum appears in the free energy surface with gradually larger Rg up to χEtOH ∼ 0.2 (44.75% v/v). This suggests gradual unfolding of the protein. At a higher concentration of alcohol (χEtOH > 0.2), the minimum at large Rg vanishes, indicating compaction. Analysis of the contact map and the solvent organization around protein indicates a preferential solvation of the hydrophobic residues by ethanol up to χEtOH = 0.2 (44.75% v/v) and this causes the gradual unfolding of the protein. At high concentration (χEtOH = 0.3 (58% v/v)), due to structural organization in bulk water-ethanol binary mixture, the extent of preferential solvation by ethanol decreases. This causes a structural transition of Cyt C towards a more compact state.

Cytochrome†c-Capped Fluorescent Gold Nanoclusters: Imaging of Live Cells and Delivery of Cytochrome†c.

Cytochrome c-capped fluorescent gold nanoclusters (Au-NCs) are used for imaging of live lung and breast cells. Delivery of cytochrome c inside the cells is confirmed by covalently attaching a fluorophore (Alexa Fluor 594) to cytochrome c-capped Au-NCs and observing fluorescence from Alexa 594 inside the cell. Mass spectrometry studies suggest that in bulk water, addition of glutathione (GSH) to cytochrome c-capped Au-NCs results in the formation of glutathione-capped Au-NCs and free apo-cytochrome c. Thus glutathione displaces cytochrome c as a capping agent. Using confocal microscopy, the emission spectra and decay of Au-NCs are measured in live cells. From the position of the emission maximum it is shown that the Au-NCs exist as Au8 in bulk water and as Au13 inside the cells. Fluorescence resonance energy transfer from cytochrome c-Au-NC (donor) to Mitotracker Orange (acceptor) indicates that the Au-NCs localise in the mitochondria of live cells.

Cancer Cell Imaging Using in Situ Generated Gold Nanoclusters.

In situ generated fluorescent gold nanoclusters (Au-NCs) are used for bio-imaging of three human cancer cells, namely, lung (A549), breast (MCF7), and colon (HCT116), by confocal microscopy. The amount of Au-NCs in non-cancer cells (WI38 and MCF10A) is 20-40 times less than those in the corresponding cancer cells. The presence of a larger amount of glutathione (GSH) capped Au-NCs in the cancer cell is ascribed to a higher glutathione level in cancer cells. The Au-NCs exhibit fluorescence maxima at 490-530 nm inside the cancer cells. The fluorescence maxima and matrix-assisted laser desorption ionization (MALDI) mass spectrometry suggest that the fluorescent Au-NCs consist of GSH capped clusters with a core structure (Au8-13). Time-resolved confocal microscopy indicates a nanosecond (1-3 ns) lifetime of the Au-NCs inside the cells. This rules out the formation of aggregated Au-thiolate complexes, which typically exhibit microsecond (≈1000 ns) lifetimes. Fluorescence correlation spectroscopy (FCS) in live cells indicates that the size of the Au-NCs is ≈1-2 nm. For in situ generation, we used a conjugate consisting of a room-temperature ionic liquid (RTIL, [pmim][Br]) and HAuCl4. Cytotoxicity studies indicate that the conjugate, [pmim][AuCl4], is non-toxic for both cancer and non-cancer cells.

Intermittent Fluorescence Oscillations in Lipid Droplets in a Live Normal and Lung Cancer Cell: Time-Resolved Confocal Microscopy.

Intermittent structural oscillation in the lipid droplets of live lung cells is monitored using time-resolved confocal microscopy. Significant differences are observed between the lung cancer cell (A549) and normal (nonmalignant) lung cell (WI38). For this study, the lipid droplets are covalently labeled with a fluorescent dye, coumarin maleimide (7-diethylamino-3-(4-maleimido-phenyl)-4-methylcoumarin, CPM). The number of lipid droplets in the cancer cell is found to be ∼20-fold higher than that in the normal (nonmalignant) cell. The fluctuation in the fluorescence intensity of the dye (CPM) is attributed to the red-ox processes and periodic formation/rupture of the S-CPM bond. The amount of reactive oxygen species (ROS) is much higher in a cancer cell. This is manifested in faster oscillations (0.9 ± 0.3 s) in cancer cells compared to that in the normal cells (2.8 ± 0.7 s). Solvation dynamics in the lipid droplets of cancer cells is slower compared to that in the normal cell.