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 cemented to the opening by means of a silicone epoxy. The bottle is then inverted for measurement. </li>
 <li>Because our system depends on precise timing of the pump and signal pulses, we are constrained not to use the conventional focus adjustment of our microscope, which translates the objective up and down. This adds and subtracts optical path from our system. Instead, we place our samples on a secondary stage mounted on top of the microscope stage that has its own independent focus control.</li>
</ol>


2.	In order to take time-gated Raman spectra, the excitation beam must be properly prepared. 
<ol> 
 
 <li>We start with light emerging from a 2.4 W tunable pulsed Ti:Sapph laser (Chameleon, Coherent Systems, Santa Clara, CA). Each pulse in the 80 MHz pulse train has 30 nJ of energy, has a temporal width of 140 fs, and has a spectrum centered at 808 nm with a spectral bandwidth of ~6 nm. </li>
 <li>To prevent back-reflections from re-entering the laser cavity, the light is passed through a Faraday isolator. A half-wave plate placed before the Faraday isolator allows continuous tuning of the total power sent into the system. </li>
 <li>Because 6 nm is too broad a bandwidth to resolve most Raman modes, the beam is sent through a very narrow (0.8 nm FWHM, Andover Corporation, Salem, NH) bandpass filter centered at 808 nm. </li>
 <li>The light is then focused by an achromatic doublet (f = 100 mm) onto a 5 mm &beta;-Barium Borate (BBO) crystal (CASIX, Fuzhou, Fujian, P. R. China) to halve the wavelength from 808 nm to 404 nm. The BBO crystal is placed in a mount with tip and tilt controls, and also mounted on a translation stage. To maximize the efficiency of the wavelength conversion, the crystal must be placed precisely symmetric about the focus of the doublet, and with its crystal axis aligned to the polarization of the incoming beam. </li>
 <li>Because efficiency of the wavelength conversion is polarization-dependent, control over the amount of light sent to the sample can be obtained by placing a second half-wave plate after the Faraday isolator. By rotating this waveplate, the intensity of light sent to the sample can be adjusted independently of the intensity sent in the pump beam (to be discussed below).</li>
 <li>The wavelength-converted light is recollimated by a second achromatic doublet (f = 500 mm) chosen such that the exiting beam is large enough to fill the back aperture of the microscope objective, and directed into an inverted microscope (IX-71, Olympus, Center Valley, PA) by means of two steering mirrors. </li>
 <li>The microscope objective, being the optical element housed within the most monolithic component of the system, defines the optic axis. To align the excitation beam to this axis, a mirror is placed in the sample plane of the microscope. The two steering mirrors are then iteratively tuned while observing the back-reflected laser beam on a CCD cam-era (&mu;Eye, IDS, Obersulm, Germany) attached to the imaging port of the microscope. Assuming that the image on the camera is centered on the microscope's field-of-view, the beam is on-axis when the focal spot is centered on the microscope chip and translation of the objective along the z-axis does not change the center point of the defocused beam. </li>
 <li>When a sample is placed in the sample plane and irradiated with laser light Raman scattering occurs. A dichroic filter placed below the microscope objective separates the wavelength-shifted Raman scattered light (and fluorescence) from the excitation beam, directing the Raman scattered light to the side port of the microscope. The microscope was modified to remove any lenses within this path, such that the signal light emerges from the microscope collimated. </li>
 <li>Because the signal beam emerging from the microscope is larger than the clear aperture of the Glan-thompson polarizers, a 0.47x telescope constructed of two achromatic doublets (f1 = 75 mm, f2 = 35 mm) is used to shrink the beam. </li>
 <li>The signal light is then polarized by a Glan-Thompson polarizer oriented at 0&deg; with respect to vertical in the lab frame, and directed to a dichroic mirror where it is recombined with the pump beam. </li>
</ol>
3.	In order for the optical gate to operate at peak efficiency, care must be taken in the preparation of the pump beam (Kerr beam) as well. 

<ol>
	<li>The pump pulse is first magnified by 0.35x with a telescope constructed with two achromatic doublets (f1 = 35 mm, f2 = 100 mm) to match the final size of the signal beam. </li>
 <li>The pump beam is then sent into a delay line composed of a right angle prism placed on a linear translation stage that can be tuned to ensure temporal overlap of the pump and signal pulses (tuning to be discussed below).</li>
 <li>After the delay line, the beam is sent through a half-wave plate and polarizer oriented at 45&deg; with respect to vertical in the lab frame. This ensures the proper polarization state of the pump beam when it reaches the nonlinear medium. </li>
 <li>The light is then reflected off of two steering mirrors, one with piezo-electric controls, that are used to finely adjust the position of the pump beam such that it overlaps spatially with the signal beam. The overlap is obtained by observing the pump and signal beams at two locations, one close and one far from the dichroic mirror where the beams are combined. By using the first steering mirror to overlap the two beams at the near point, and the piezo mirror to overlap the beams at the far point, the pump beam can be made exactly collinear with the signal beam. </li>
 <li>With the two beams combined, the Kerr gate and collection system are set up to maximize the collected time-gated signal. </li>
</ol>
4.	With the two beams combined, the Kerr gate and collection system are set up to maximize the collected time-gated signal. 
<ol> 
 <li>The pump and signal beams are first passed through a dichroic filter that has an OD of 6 at 404 nm to prevent any residual excitation light from exciting Raman scattering within the nonlinear medium. </li>
 <li>The pump and signal beams are then both focused by an achromatic doublet (f = 35 mm) into a 1 cm pathlength quartz cuvette containing the nonlinear material. Any nonlinear material having a suitably high nonlinear index (higher than CS2), and suitably short temporal response (&lt;2 ps), can be utilized here. For these experiments, we use carbon disulfide, CS2 , that has a nonlinear index n2 = 3.1 x 10&#x2212;18 m2/W. The light is then recollimated by a second doublet with a focal length identical to the first.</li>
 <li>The beams are then passed through and Glan-Thompson analyzer on a rotating mount, and then through a set of absorption and interference filters that combined have an OD of 10 at 808 nm. </li>
 <li>Finally, the signal light is focused by an achromatic doublet (f = 35 mm) into a 50 &mu;m multimode optical fiber, where the fiber is mounted in a stage that allows translation in x, y, and z. The fiber is then coupled to a commercial imaging spectrograph with attached CCD camera (SP2300i and Pixis 100B, respectively, both manufactured by Princeton Instruments, Trenton, NJ). </li>
 <li>To align the collection system to maximize collected signal, the analyzer is set to 0&deg; and a test sample of toluene is placed in the sample plane. By adjusting the x, y, and z controls of the fiber mount, the collected Raman signal is optimized. </li>
 <li>To ensure proper spatial and temporal overlap of the pump and signal beams, a mirror is placed in the sample plane of the microscope. The 404 nm filter is removed from the system. With the analyzer rotated to 90&deg; the retroreflected 404 nm beam is sent into the spectrograph with the intensity adjusted such that it does not saturate the camera. With the pump beam off, the analyzer is rotated to minimize the transmitted 404 nm signal. With the pump beam turned back on, the delay stage is slowly tuned until the transmission of the 404 nm light begins to increase. Then, by iteratively tuning the delay stage, the piezo-electric mirror, and the x, y, and z controls of the fiber, one maximizes the signal.</li>
 <li>Because the retro-reflected 404 nm beam and the Raman scattered light may take slightly different paths through the system, final adjustments are made by placing a strong Raman scatterer (such as toluene) on the sample stage and the slightly tweaking the alignment by changing the delay stage, piezeo-electric mirror, and x, y, and z controls of the fiber to optimize the Raman signal. </li>
</ol>

5.	Collection of a single time-gated Raman spectrum requires acquisition of several spectra to correct for system artifacts.

<ol>
	<li>With the analyzer set to 0&deg;, the excitation beam on and the pump beam off, an "ungated" spectrum is taken, representing the spectrum that would be acquired without the benefit of the time-gating system. </li>
 <li>With the analyzer set to 90&deg;, the excitation beam and the pump beam still off, a back-ground spectrum is taken, representing the amount of stray light leaking through the polarizers and other elements in the system. </li>
 <li>With the analyzer remaining at 90&deg; and all beams on, the "gated" spectrum is taken, representing the Raman scattered light allowed through the crossed polarizers by the presence of the pump beam. </li>
 <li>Finally, with the analyzer still at 90&deg;, the excitation beam off and the pump beam on, a second background spectrum is taken, representing the contribution to the "gated" spectrum of residual amounts of 808 nm light entering the spectrograph. </li>
 <li>Additionally, a spectrum is taken with all lasers off, characterizing the baseline "dark" signal level of the camera and electronics. </li>
 <li>Because the spectra often require lengthy integration times, it is imperative to break this integration time into several pieces (frames) to allow for proper correction of cosmic rays - a common problem when acquiring data over spans of several minutes. </li>
</ol>
6.	Once acquired, several processing steps are helpful to improve the quality and appearance of the data.
<ol>
	<li>First, all spectra are dark corrected by subtracting off the "dark" spectrum. </li>
 <li>To correct for cosmic rays, the several frames constituting one acquisition are compared to each other on a pixel-by-pixel basis. For any pixel of any frame that falls outside of 2 median deviations from the median value for that pixel across all frames, that pixel's value is replaced by the median value across all frames.</li>
 <li>The cosmic ray corrected frames are then averaged and smoothed by a Savitzky-Golay filter 1. </li>
 <li>To extract the "true" gated spectrum, we subtract the "excitation-only" and "pump-only" spectra from the measured gated spectrum. </li>
 <li>Finally, the gated and ungated spectra are then background corrected by subtracting a 5th-order polynomial, determined using the method of Lieber and Mahadevan-Jansen 2.
</li>
</ol>
7. Representative Results: 
<img src="/files/ftp_upload/2592/2592fig1.jpg" alt="Figure 1"/>
 Figure 1. Schematic diagram of the Kerr gating system. The pump beam path is shown in red, while the SHG path is shown in blue. The path where Raman and Fluorescence are overlapped is shown in green, while the path where the Fluorescence has been temporally filtered out is shown in yellow. Abbreviations as follows: BPF, band pass filter; CCD, charge-coupled device; DCM, dichroic mirror; FI, Faraday isolator; &lambda;/2, half wave plate; LPF, long-pass filter; NLM, nonlinear medium; P, polarizer; SHG, second harmonic generation crystal.
<img src="/files/ftp_upload/2592/2592fig2.jpg" alt="Figure 2"/> 
Figure 2. Top: Raw spectra of coumarin dissolved in immersion oil. Red curve shows the spectrum taken with the gate held open (analyzer set for maximum transmission). Black curve shows the spectrum taken with the analyzer aligned for minimum transmission and a pump beam applied (the gated spectrum). Blue curve shows the spectum taken with the analyzer aligned for minimum transmission and no pump beam applied (gate held closed). Green curve shows the spectrum taken with only the pump beam applied. Dashed magenta lines indicate spectral region shown in panel below. All spectra have been smoothed with a 11 point, 3rd order Savitzky-Golay filter. Bottom: Spectra of coumarin dissolved in immersion oil after fluorescence background subtraction. Red curve is the spectrum with the gate held open, and the blue curve is the gated spectrum. The gated spectrum clearly shows the convoluted high-wavenumber peak characteristic of oils.

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No conflicts of interest declared.

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.

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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 ...

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13.0K Views.  University of California, Davis. 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.

Video: Rejection of Fluorescence Background in Resonance and Spontaneous Raman Microspectroscopy

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 ...