Name: rejection of fluorescence background in resonance and spontaneous raman microspectroscopy
Uploaded: 2011-05-18T12:00:00
Description: Watch this Scientific Journal Video about Rejection of Fluorescence Background in Resonance and Spontaneous Raman Microspectroscopy at JoVE.com

This work was funded by NSF award DBI 0852891. Part of this work was also funded by the Center for Biophotonics Science and Technology, a designated NSF Science and Technology Center managed by the University of California, Davis, under Cooperative Agreement No. PHY0120999.

1.	Some care must be taken in preparing and placing a Raman sample within this system.
<ol>
	<li>Because the system typically makes use of very high numerical aperture objectives with very short working distances, the samples are placed on a coverslip. Biological samples are typically placed on a No. 1 thickness coverslip mounted in an Attofluor cell chamber (Invitrogen, Carlsbad, CA). </li>
 <li>Liquid samples, particularly those toxic to humans, are placed in a small glass bottle with a coverslip cem...

<ol>
	<li>Savitzky, A., Golay, M. J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Smoothing+and+differentiation+of+data+by+simplified+least+squares+procedures.">Smoothing and differentiation of data by simplified least squares procedures.</a> Analytical Chemistry. 36, 1627-1639 (1964).</li><li>Lieber, C. A., Mahadevan-Jansen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+method+for+subtraction+of+fluorescence+from+biological+Raman+spectra.">Automated method for subtraction of fluorescence from biological Raman spectra.</a> Applied Spectroscopy. 57, 1363-1367 (2003).</li><li>Gniadecka, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Melanoma+diagnosis+by+Raman+spectroscopy+and+neural+networks:+Structure+alterations+in+proteins+and+lipids+in+intact+cancer+tissue.">Melanoma diagnosis by Raman spectroscopy and neural networks: Structure alterations in proteins and lipids in intact cancer tissue.</a> Journal of Investiga-tive Dermatology. 122, 443-449 (2004).</li><li>Lieber, C. A., Majumder, S. K., Billheimer, D., Ellis, D. L., Mahadevan-Jansen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Raman+microspectroscopy+for+skin+cancer+detection+in+vitro.">Raman microspectroscopy for skin cancer detection in vitro.</a> Journal of Biomedical Optics. 13, 024013-024013 (2008).</li><li>Chen, K., Qin, Y., Zheng, F., Sun, M., Shi, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diagnosis+of+colorectal+cancer+using+Raman+spectroscopy+of+laser-trapped+single+living+epithelial+cells.">Diagnosis of colorectal cancer using Raman spectroscopy of laser-trapped single living epithelial cells.</a> Optics Letters. 31, 2015-2017 (2006).</li><li>Chan, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nondestructive+identification+of+individual+leukemia+cells+by+laser+trapping+Raman+spectroscopy.">Nondestructive identification of individual leukemia cells by laser trapping Raman spectroscopy.</a> Analytical Chemistry. 80, 2180-2187 (2008).</li><li>Zhu, Q. Y., Quivey, R. G., Berger, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+bacterial+concentration+fractions+in+polymicrobial+mixtures+by+Raman+microspectroscopy.">Measurement of bacterial concentration fractions in polymicrobial mixtures by Raman microspectroscopy.</a> Journal of Biomedical Optics. 9, 1182-1186 (2004).</li><li>R&#246;sch, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemotaxonomic+identification+of+single+bacteria+by+micro-Raman+spectroscopy:+Application+to+clean-room-relevant+biological+contaminations.">Chemotaxonomic identification of single bacteria by micro-Raman spectroscopy: Application to clean-room-relevant biological contaminations.</a> Applied and Environmental Microbiology. 71, 1626-1637 (2005).</li><li>Moritz, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Raman+spectroscopic+signatures+of+the+metabolic+states+of+escherichia+coli+cells+and+their+dependence+on+antibiotics+treatment.">Raman spectroscopic signatures of the metabolic states of escherichia coli cells and their dependence on antibiotics treatment.</a> Biophysical Journal. 98, 742a-742a (2010).</li><li>Dehring, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identifying+chemical+changes+in+subchondral+bone+taken+from+murine+knee+joints+using+Raman+spectroscopy.">Identifying chemical changes in subchondral bone taken from murine knee joints using Raman spectroscopy.</a> Applied Spectroscopy. 60, 1134-1141 (2006).</li><li>Berger, A. J., Koo, T. W., Itzkan, I., Horowitz, G., Feld, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multicomponent+blood+analysis+by+near-infrared+Raman+spectroscopy.">Multicomponent blood analysis by near-infrared Raman spectroscopy.</a> Applied Optics. 38, 2916-2926 (1999).</li><li>Qi, D., Berger, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+concentration+measurement+in+blood+serum+and+urine+samples+using+liquid-core+optical+fiber+Raman+spectroscopy.">Chemical concentration measurement in blood serum and urine samples using liquid-core optical fiber Raman spectroscopy.</a> Applied Spectroscopy. 46, 1726-1734 (2007).</li><li>Beier, B. D., Berger, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Method+for+automated+background+subtraction+from+Raman+spectra+containing+known+contaminants.">Method for automated background subtraction from Raman spectra containing known contaminants.</a> The Analyst. 134, 1198-1202 (2009).</li><li>De Luca, A. C., Mazilu, M., Riches, A., Herrington, C. S., Dholakia, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Online+fluorescence+suppression+in+modulated+Raman+spectroscopy.">Online fluorescence suppression in modulated Raman spectroscopy.</a> Analytical Chemistry. 82, 738-745 (2010).</li><li>Evans, C. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+imaging+of+tissue+in+vivo+with+video-rate+coherent+anti-Stokes+Raman+scattering+microscopy.">Chemical imaging of tissue in vivo with video-rate coherent anti-Stokes Raman scattering microscopy.</a> Proceedings of the National Academy of Sciences of the United States of America. 102, 16807-16812 (2005).</li><li>Jones, W. J., Stoiche, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inverse+raman+spectra:+Induced+absorption+at+optical+frequencies.">Inverse raman spectra: Induced absorption at optical frequencies.</a> Physical Review Letters. 13, 657-659 (1964).</li><li>Freudiger, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-Free+et+Imaging+with+High+Sensitivity+by+Stimulated+Raman+Scattering+Microscopy.">Label-Free et Imaging with High Sensitivity by Stimulated Raman Scattering Microscopy.</a> Science. 322, 1857-1861 (2008).</li><li>Cui, M., Bachler, B. R., Ogilvie, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparing+coherent+and+spontaneous+Raman+scattering+under+biological+imaging+conditions.">Comparing coherent and spontaneous Raman scattering under biological imaging conditions.</a> Optics Letters. 34, 773-775 (2009).</li><li>Matousek, P., Towrie, M., Stanley, A., Parker, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+rejection+of+fluorescence+from+Raman+spectra+using+picosecond+Kerr+gating.">Efficient rejection of fluorescence from Raman spectra using picosecond Kerr gating.</a> Applied Spectroscopy. 53, 1485-1489 (1999).</li><li>Matousek, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+suppression+in+resonance+Raman+spectroscopy+using+a+high-performance+picosecond+Kerr+gate.">Fluorescence suppression in resonance Raman spectroscopy using a high-performance picosecond Kerr gate.</a> Journal of Raman Spectroscopy. 32, 983-988 (2001).</li><li>Knorr, F., Smith, Z. J., Wachsmann-Hogiu, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+time-gated+system+for+Raman+spectroscopy+of+biological+samples.">Development of a time-gated system for Raman spectroscopy of biological samples.</a> Optics Express. 18, 20049-20058 (2010).</li></ol>

The field of biomedical Raman spectroscopy has seen increasing interest over the past several years as a result of its demonstrated potential for solving several difficult challenges in biological diagnostics. For example, Raman spectra have been shown to have diagnostic value in cancer detection 3, 4, 5, 6. Raman spectroscopy has also been used in bacterial quantitation 7, 8 and bacterial drug response 9. It has also found application in a broad range of other biomedical applications ran...

<table border="1">
	<tbody>
 <tr>
 	<th>Name</th>
 <th>Company</th>
 <th>Catalog Number</th>
 <th>Comments</th>
 </tr>
 <tr>
 	<td>Lenses</td>
 <td>ThorLabs</td>
 <td>Various</td>
 <td>All lenses coated to have maximum transmission losses of 1% each</td>
 </tr>
 <tr>
 	<td>Tunable Ti:Sapph laser</td>
 <td>Coherent</td>
 <td>Chameleon</td>
 <td>30 nJ, 200 fs, 80 MHz</td>
 </tr>
 <tr>
 	<td>40X oil immersion objective</td>
 <td>Olympus</td>
 <td>UApo/340</td>
 <td>NA = 1.35</td>
 </tr>
 <tr>
 	<td>Inverted microscope</td>
 <td>Olympus</td>
 <td>IX-71</td>
 <td>Modified to remove all lenses in side port</td>
 </tr>
 <tr>
 	<td>Half wave plate</td>
 <td>Thorlabs</td>
 <td>AHWP05M-600</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Glan-Thompson polarizer</td>
 <td>Thorlabs</td>
 <td>GTH10M</td>
 <td>&tilde;10% transmission loss</td>
 </tr>
 <tr>
 	<td>Spectrometer</td>
 <td>PI Acton</td>
 <td>SP2300i</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>CCD</td>
 <td>PI Acton</td>
 <td>Pixis 100B</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Mathmatical software</td>
 <td>The MathWorks</td>
 <td>MATLAB</td>
 <td>version 2008a</td>
 </tr>
 <tr>
 	<td>Faraday isolator</td>
 <td>EOT</td>
 <td>BB8-5I</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Piezo-electric mirror</td>
 <td>Newport</td>
 <td>AG-M100</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>BBO crystal</td>
 <td>CASIX</td>
 <td>custom</td>
 <td>1 mm thickness</td>
 </tr>
 <tr>
 	<td>Bandpass filter 1</td>
 <td>Andover</td>
 <td>008FC14</td>
 <td>808 &plusmn; 0.4 nm</td>
 </tr>
 <tr>
 	<td>Dichroic mirror</td>
 <td>Semrock</td>
 <td>FF662-FDI01</td>
 <td>band edge at 662 nm</td>
 </tr>
 <tr>
 	<td>Long-pass filter</td>
 <td>Semrock</td>
 <td>BLP01-405R</td>
 <td>band edge at 417 nm</td>
 </tr>
 <tr>
 	<td>Bandpass filter 2</td>
 <td>Semrock</td>
 <td>FF02-447/60</td>
 <td>417-447 nm</td>
 </tr>
 <tr>
 	<td>CS2</td>
 <td>Sigma-Aldrich</td>
 <td>335266</td>
 <td>99% purity</td>
 </tr>
 <tr>
 	<td>Coumarin 30</td>
 <td>Sigma-Aldrich</td>
 <td>546127</td>
 <td>99% purity</td>
 </tr>
 <tr>
 	<td>Immersion oil</td>
 <td>Cargille</td>
 <td>16242</td>
 <td>Type DF</td>
 </tr>
 
 </tbody>
</table>

rejection of fluorescence background in resonance and spontaneous raman microspectroscopy

Raman spectroscopy is often plagued by a strong fluorescent background, particularly for biological samples. If a sample is excited with a train of ultrafast pulses, a system that can temporally separate spectrally overlapping signals on a picosecond timescale can isolate promptly arriving Raman scattered light from late-arriving fluorescence light. Here we discuss the construction and operation of a complex nonlinear optical system that uses all-optical switching in the form of a low-power optical Kerr gate to isolate Raman and fluorescence signals. A single 808 nm laser with 2.4 W of average power and 80 MHz repetition rate is split, with approximately 200 mW of 808 nm light being converted to &lt; 5 mW of 404 nm light sent to the sample to excite Raman scattering. The remaining unconverted 808 nm light is then sent to a nonlinear medium where it acts as the pump for the all-optical shutter. The shutter opens and closes in 800 fs with a peak efficiency of approximately 5%. Using this system we are able to successfully separate Raman and fluorescence signals at an 80 MHz repetition rate using pulse energies and average powers that remain biologically safe. Because the system has no spare capacity in terms of optical power, we detail several design and alignment considerations that aid in maximizing the throughput of the system. We also discuss our protocol for obtaining the spatial and temporal overlap of the signal and pump beams within the Kerr medium, as well as a detailed protocol for spectral acquisition. Finally, we report a few representative results of Raman spectra obtained in the presence of strong fluorescence using our time-gating system.

Watch this Scientific Journal Video about Rejection of Fluorescence Background in Resonance and Spontaneous Raman Microspectroscopy at JoVE.com

Rejection of Fluorescence Background in Resonance and Spontaneous Raman Microspectroscopy

We discuss the construction and operation of a complex nonlinear optical
system that uses ultrafast all-optical switching to isolate Raman from fluorescence
signals. Using this system we are able to successfully separate Raman and fluorescence
signals utilizing pulse energies and average powers that remain biologically safe.

Raman spectroscopy is often plagued by a strong fluorescent background, particularly for biological samples. If a sample is excited with a train of ...

rejection-fluorescence-background-resonance-spontaneous-raman

Research

JoVE Journal

Bioengineering

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 F&ouml;rster 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.

Fluorescence Lifetime Imaging (FLIM) has emerged as a key technique to image the environment and interaction of specific proteins and dyes in living cells. FLIM of fluorescent molecular rotors allows mapping of viscosity 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 ...

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

3D Orbital Tracking in a Modified Two-photon Microscope: An Application to the Tracking of Intracellular Vesicles

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a modified two-photon microscope1. As opposed to conventional approaches (raster scan or wide field based on a stack of frames), the 3D orbital tracking allows to localize and follow with a high spatial (10 nm accuracy) and temporal resolution (50 Hz frequency response) the 3D displacement of a moving fluorescent particle on length-scales of hundreds of microns2. The method is based on a feedback algorithm that controls the hardware of a two-photon laser scanning microscope in order to perform a circular orbit around the object to be tracked: the feedback mechanism will maintain the fluorescent object in the center by controlling the displacement of the scanning beam3-5. To demonstrate the advantages of this technique, we followed a fast moving organelle, the lysosome, within a living cell6,7. Cells were plated according to standard protocols, and stained using a commercially lysosome dye. We discuss briefly the hardware configuration and in more detail the control software, to perform a 3D orbital tracking experiment inside living cells. We discuss in detail the parameters required in order to control the scanning microscope and enable the motion of the beam in a closed orbit around the particle. We conclude by demonstrating how this method can be effectively used to track the fast motion of a labeled lysosome along microtubules in 3D within a live cell. Lysosomes can move with speeds in the range of 0.4-0.5 &#181;m/sec, typically displaying a directed motion along the microtubule network8.

In this video protocol we track - at high speed and in three dimensions - fluorescently labeled lysosomes within living cells, using the orbital tracking method in a modified two-photon microscope.

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a ...

Luminescence Resonance Energy Transfer to Study Conformational Changes in Membrane Proteins Expressed in Mammalian Cells

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range of 10-100 &#197;. By measuring the distances under various ligated conditions, conformational changes of the protein can be easily assessed. With LRET, a lanthanide, most often chelated terbium, is used as the donor fluorophore, affording advantages such as a longer donor-only emission lifetime, the flexibility to use multiple acceptor fluorophores, and the opportunity to detect sensitized acceptor emission as an easy way to measure energy transfer without the risk of also detecting donor-only signal. Here, we describe a method to use LRET on membrane proteins expressed and assayed on the surface of intact mammalian cells. We introduce a protease cleavage site between the LRET fluorophore pair. After obtaining the original LRET signal, cleavage at that site removes the specific LRET signal from the protein of interest allowing us to quantitatively subtract the background signal that remains after cleavage. This method allows for more physiologically relevant measurements to be made without the need for purification of protein.

We describe here an improved Luminescence Resonance Energy Transfer (LRET) method where we introduce a protease cleavage site between the donor and acceptor fluorophore sites. This modification allows us to obtain specific LRET signals arising from membrane proteins of interest, allowing for the study of membrane proteins without protein purification.

Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range ...

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Light-sheet Fluorescence Microscopy to Capture 4-Dimensional Images of the Effects of Modulating Shear Stress on the Developing Zebrafish Heart

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths from the myocardium. Genetic program defects in the Notch signaling cascade are involved in ventricular defects such as Left Ventricular Non-Compaction Cardiomyopathy or Hypoplastic Left Heart Syndrome. Using this protocol, it can be determined that shear stress driven trabeculation and Notch signaling are related to one another. Using Light-sheet Fluorescence Microscopy, visualization of the developing zebrafish heart was possible. In this manuscript, it was assessed whether hemodynamic forces modulate the initiation of trabeculation via Notch signaling and thus, influence contractile function occurs. For qualitative and quantitative shear stress analysis, 4-D (3-D+time) images were acquired during zebrafish cardiac morphogenesis, and integrated light-sheet fluorescence microscopy with 4-D synchronization captured the ventricular motion. Blood viscosity was reduced via gata1a-morpholino oligonucleotides (MO) micro-injection to decrease shear stress, thereby, down-regulating Notch signaling and attenuating trabeculation. Co-injection of Nrg1 mRNA with gata1a MO rescued Notch-related genes to restore trabeculation. To confirm shear stress driven Notch signaling influences trabeculation, cardiomyocyte contraction was further arrested via tnnt2a-MO to reduce hemodynamic forces, thereby, down-regulating Notch target genes to develop a non-trabeculated myocardium. Finally, corroboration of the expression patterns of shear stress-responsive Notch genes was conducted by subjecting endothelial cells to pulsatile flow. Thus, the 4-D light-sheet microscopy uncovered hemodynamic forces underlying Notch signaling and trabeculation with clinical relevance to non-compaction cardiomyopathy.

Here, we present a protocol to visualize developing hearts in zebrafish in 4-Dimensions (4-D). 4-D imaging, via light-sheet fluorescence microscopy (LSFM), takes 3-Dimensional (3-D) images over time, to reconstruct developing hearts. We show qualitatively and quantitatively that shear stress activates endocardial Notch signaling during chamber development, which promotes cardiac trabeculation.

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths ...

Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense lipidic nanotubes that are, in terms of morphology and dynamics, reminiscent of ER. The networks are derived from phospholipid double bilayer membrane patches adhering to a transparent Al2O3 substrate. The adhesion is mediated by Ca2+ in the ambient buffer. Subsequent depletion of Ca2+ by means of BAPTA/EDTA causes retraction of the membrane, resulting in spontaneous lipid nanotube network formation. The method only comprises phospholipids and microfabricated surfaces for simple formation of an ER model and does not require the addition of proteins or chemical energy (e.g., GTP or ATP). In contrast to the 3D morphology of the cellular endoplasmic reticulum, the model is two-dimensional (albeit the nanotube dimensions, geometry, structure, and dynamics are maintained). This unique in vitro ER model consists of only a few components, is easy to construct, and can be observed under a light microscope. The resulting structure can be further decorated for additional functionality, such as the addition of ER-associated proteins or particles to study transport phenomena among the tubes. The artificial networks described here are suitable structural models for the cellular ER, whose unique characteristic morphology has been shown to be related to its biological function, whereas details regarding formation of the tubular domain and rearrangements within are still not completely understood. We note that this method uses Al2O3 thin-film-coated microscopy coverslips, which are commercially available but require special orders. Therefore, it is advisable to have access to a microfabrication facility for preparation.

Solid-supported, protein-free, double phospholipid bilayer membranes (DLBM) can be transformed into complex and dynamic lipid nanotube networks and can serve as 2D bottom-up models of the endoplasmic reticulum.

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense ...

In vivo Imaging of Biological Tissues with Combined Two-Photon Fluorescence and Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy enables label-free imaging of the biological tissues in its natural microenvironment based on intrinsic molecular vibration, thus providing a perfect tool for in vivo study of biological processes at subcellular resolution. By integrating two-photon excited fluorescence (TPEF) imaging into the SRS microscope, the dual-modal in vivo imaging of tissues can acquire critical biochemical and biophysical information from multiple perspectives which helps understand the dynamic processes involved in cellular metabolism, immune response and tissue remodeling, etc. In this video protocol, the setup of a TPEF-SRS microscope system as well as the in vivo imaging method of the animal spinal cord is introduced. The spinal cord, as part of the central nervous system, plays a critical role in the communication between the brain and peripheral nervous system. Myelin sheath, abundant in phospholipids, surrounds and insulates the axon to permit saltatory conduction of action potentials. In vivo imaging of myelin sheaths in the spinal cord is important to study the progression of neurodegenerative diseases and spinal cord injury. The protocol also describes animal preparation and in vivo TPEF-SRS imaging methods to acquire high-resolution biological images.

In vivo Imaging of Biological Tissues with Combined Two-Photon Fluorescence and Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows label-free imaging of biomolecules based on their intrinsic vibration of specific chemical bonds. In this protocol, the instrumental setup of an integrated SRS and two-photon fluorescence microscope is described to visualize cellular structures in the spinal cord of live mice.

Stimulated Raman scattering (SRS) microscopy enables label-free imaging of the biological tissues in its natural microenvironment based on intrinsic ...

Real-Time, Two-Color Stimulated Raman Scattering Imaging of Mouse Brain for Tissue Diagnosis

Stimulated Raman scattering (SRS) microscopy has emerged as a powerful optical imaging technique for tissue diagnosis. In recent years, two-color SRS has been shown to be able to provide hematoxylin and eosin (H&amp;E)-equivalent images that allow fast and reliable diagnosis of brain cancer. Such capability has enabled exciting intraoperative cancer diagnosis applications. Two-color SRS imaging of tissue can be done with either a picosecond or femtosecond laser source. Femtosecond lasers have the advantage of enabling flexible imaging modes, including fast hyperspectral imaging and real-time, two-color SRS imaging. A spectral-focusing approach with chirped laser pulses is typically used with femtosecond lasers to achieve high spectral resolution. 
Two-color SRS acquisition can be realized with orthogonal modulation and lock-in detection. The complexity of pulse chirping, modulation, and characterization is a bottleneck for the widespread adoption of this method. This article provides a detailed protocol to demonstrate the implementation and optimization of spectral-focusing SRS and real-time, two-color imaging of mouse brain tissue in the epi-mode. This protocol can be used for a broad range of SRS imaging applications that leverage the high speed and spectroscopic imaging capability of SRS.

Stimulated Raman scattering (SRS) microscopy is a powerful, nondestructive, and label-free imaging technique. One emerging application is stimulated Raman histology, where two-color SRS imaging at the protein and lipid Raman transitions are used to generate pseudo-hematoxylin and eosin images. Here, we demonstrate a protocol for real-time, two-color SRS imaging for tissue diagnosis.

Stimulated Raman scattering (SRS) microscopy has emerged as a powerful optical imaging technique for tissue diagnosis. In recent years, two-color SRS has ...