Name: real-time, two-color stimulated raman scattering imaging of mouse brain for tissue diagnosis
Uploaded: 2022-02-01T14:40:00
Description: Watch this Scientific Journal Video about Real-Time, Two-Color Stimulated Raman Scattering Imaging of Mouse Brain for Tissue Diagnosis at JoVE.com

This study was supported by NIH R35 GM133435 to D.F.

All experimental animal procedures were conducted with 200 &#181;m, fixed, sectioned mouse brains, in accordance with the protocol (# 4395-01) approved by the Institute of Animal Care and Use Committee (IACUC) of the University of Washington. Wild-type mice (C57BL/6J strain) are euthanized with CO2. Then, a craniotomy is performed to extract their brains for fixation in 4% paraformaldehyde in phosphate-buffered saline. The brains are embedded in a 3% agarose and 0.3% gelatin mixture and sectioned into 200 &#95...

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The two-color SRS imaging scheme presented in this protocol hinges on the proper implementation of one-color SRS imaging. In one-color SRS imaging, the critical steps are spatial alignment, temporal alignment, modulation depth, and phase shift. Spatially combining the two beams is accomplished by a dichroic mirror. Several steering mirrors are used for fine adjustment when sending the beams to the dichroic mirror. Once the beams are combined with the dichroic mirror, spatial alignment can be confirmed by picking off the ...

Traditional tissue diagnostics rely on staining protocols followed by examination under an optical microscope. One common staining method used by pathologists is H&#38;E staining: hematoxylin stains cell nuclei a purplish blue, and eosin stains the extracellular matrix and cytoplasm pink. This simple staining remains the gold standard in pathology for many tissue diagnoses tasks, particularly cancer diagnosis. However, H&#38;E histopathology, particularly the frozen sectioning technique used in an intraoperative setting, still has limitations. The staining procedure is a laborious process involving tissue embedding, sectioning, fixation, and staining1. The typical turnaround time is 20 min or longer. Performing H&#38;E during frozen sectioning can sometimes become more challenging when multiple sections are processed at once due to the need to evaluate cellular features or growth patterns in 3D for margin assessment. Moreover, intraoperative histological techniques require skilled technicians and clinicians. Limitation in the number of board-certified pathologists in many hospitals is a constraint for intraoperative consultation in many cases. Such limitations may be alleviated with the fast development interests in digital pathology and artificial intelligence-based diagnosis2. However, the H&#38;E staining results are variable, depending on the experience of the technician, which presents additional challenges for computer-based diagnosis2.
These challenges can potentially be addressed with label-free optical imaging techniques. One such technique is SRS microscopy. SRS uses synchronized pulsed lasers&#8212;pump and Stokes&#8212;to excite molecular vibrations with high efficiency3. Recent reports have demonstrated that SRS imaging of proteins and lipids can generate H&#38;E-equivalent images (also known as stimulated Raman histology or SRH) with intact fresh tissue, which bypasses the need for any tissue processing, significantly shortens the time needed for diagnosis, and has been adapted intraoperatively4. Moreover, SRS imaging can provide 3D images, which offers additional information for diagnosis when 2D images are insufficient5. SRH is unbiased and generates digital images that are readily available for computer-based diagnosis. It quickly emerges as a possible solution for intraoperative cancer diagnosis and tumor margin analysis, especially in brain cancer6,7,8. More recently, SRS imaging of chemical changes of tissue has also been suggested to provide useful diagnostic information that can further help clinicians stratify different cancer types or stages9.
Despite its tremendous potential in tissue diagnosis applications, SRS imaging is mostly limited to academic laboratories specialized in optics due to the complexity associated with the imaging platform, which includes ultrafast lasers, the laser scanning microscope, and sophisticated detection electronics. This protocol provides a detailed workflow to demonstrate the use of a common femtosecond laser source for real-time, two-color SRS imaging and the generation of pseudo-H&#38;E images from mouse brain tissue. The protocol will cover the following procedures:
Alignment and chirp optimization 
Most SRS imaging schemes use either picosecond or femtosecond lasers as the excitation source. With femtosecond lasers, the bandwidth of the laser is much larger than the Raman linewidth. To overcome this limitation, a spectral focusing approach is used to chirp the femtosecond lasers to a picosecond timescale to achieve narrow spectral resolution10. Optimal spectral resolution is only achieved when the temporal chirp (also known as the group delay dispersion or just dispersion) is properly matched for the pump and the Stokes lasers. The alignment procedure and the steps needed to optimize the dispersion of the laser beams using highly dispersive glass rods are demonstrated here.
Frequency calibration 
An advantage of spectral focusing SRS is that the Raman excitation can be quickly tuned by changing the time delay between the pump and the Stokes lasers. Such tuning affords fast imaging and reliable spectral acquisition compared to tuning laser wavelengths. However, the linear relationship between excitation frequency and time delay requires external calibration. Organic solvents with known Raman peaks are used to calibrate the Raman frequency for spectral focusing SRS.
Real-time, two-color imaging 
It is important to increase the imaging speed in tissue diagnosis applications to shorten the time needed for analyzing large tissue specimens. Simultaneous two-color SRS imaging of lipids and proteins obviates the need to tune the laser or time delay, which increases the imaging speed by more than two-fold. This is achieved by using a novel orthogonal modulation technique and dual-channel demodulation with a lock-in amplifier11. This paper describes the protocol for orthogonal modulation and dual-channel image acquisition.
Epi-mode SRS imaging 
The majority of SRS imaging shown to date is performed in transmission mode. Epi-mode imaging detects backscattered photons from tissue12. For pathology applications, surgical specimens can be quite large. For transmission mode imaging, tissue sectioning is often necessary, which undesirably requires extra time. In contrast, epi-mode imaging can work with intact surgical specimens. Because the same objective is used to collect backscattered light, there is also no need for aligning a high numerical-aperture condenser required for transmission imaging. Epi-mode is also the only option when tissue sectioning is difficult, such as with bone. Previously we have demonstrated that for brain tissue, epi-mode imaging offers superior imaging quality for tissue thickness &#62; 2 mm13. This protocol uses a polarizing beam splitter (PBS) to collect scattered photons depolarized by tissue. It is possible to collect more photons with an annular detector at the expense of the complexity of customized detector assembly12. The PBS approach is simpler to implement (similar to fluorescence), with the standard photodiode already being used for transmission mode detection.
Pseudo-H&#38;E image generation 
Once two-color SRS images are collected, they can be recolored to simulate H&#38;E staining. This paper demonstrates the procedure for converting lipid and protein SRS images to pseudo-H&#38;E SRS images for pathology applications. The experimental protocol details critical steps needed to generate high-quality SRS images. The procedure shown here is not only applicable to tissue diagnosis but also can be adapted for many other hyperspectral SRS imaging applications such as drug imaging and metabolic imaging14,15.
General system requirements 
The laser system for this protocol must be able to output 2 synchronized femtosecond laser beams. Systems ideally feature an Optical Parametric Oscillator (OPO) for broad wavelength tuning of one of the laser beams. The setup in this protocol uses a commercial laser system Insight DS+ that outputs two lasers (one fixed beam at 1,040 nm and one OPO-based tunable beam, ranging from 680 to 1,300 nm) with a repetition rate of 80 MHz. Laser scanning microscopes, either from major microscope manufacturers or home-built, can be used for SRS imaging. The utilized microscope is an upright laser scanning microscope built on top of a commercial upright microscope frame. A pair of 5 mm galvo mirrors are used to scan the laser beam. For users choosing to adopt a homebuilt laser scanning microscope, refer to a previously published protocol for the construction of a laser scanning microscope16.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 mm Achromatic Lens</td>
			<td>THORLABS</td>
			<td>AC254-100-B</td>
			<td>Broadband, 650 - 1,050 nm, achromatic lens focal length, 100 mm</td>
		</tr>
		<tr>
			<td>20 MHz bandpass filter</td>
			<td>Minicircuits</td>
			<td>BBP-21.4+</td>
			<td>Lumped LC Band Pass Filter, 19.2 - 23.6 MHz, 50 &#937;</td>
		</tr>
		<tr>
			<td>200 mm Achromatic Lens</td>
			<td>THORLABS</td>
			<td>AC254-200-B</td>
			<td>Broadband, 650 - 1,050 nm, achromatic lens focal length, 200 mm</td>
		</tr>
		<tr>
			<td>Achromatic Half Waveplate</td>
			<td>Union Optic</td>
			<td>WPA2210-650-1100-M25.4</td>
			<td>Broadband half waveplate</td>
		</tr>
		<tr>
			<td>Achromatic Quarter Waveplate</td>
			<td>Union Optic</td>
			<td>WPA4210-650-1100-M25.4</td>
			<td>Broadband quarter waveplate</td>
		</tr>
		<tr>
			<td>Beam Sampler</td>
			<td>THORLABS</td>
			<td>BSN11</td>
			<td>10:90 Plate Beamsplitter</td>
		</tr>
		<tr>
			<td>Dichroic Mirror</td>
			<td>THORLABS</td>
			<td>DMSP1000</td>
			<td>Other dichroics with a center wavelength around 1,000 nm can be used.</td>
		</tr>
		<tr>
			<td>DMSO (Dimethyl sulfoxide)</td>
			<td>Sigma Aldrich</td>
			<td>472301</td>
			<td>Solvent for calibration of Raman shift. Other solvents with known Raman peaks can be used.</td>
		</tr>
		<tr>
			<td>Electrooptic Amplitude Modulator</td>
			<td>THORLABS</td>
			<td>EO-AM-NR-C1</td>
			<td>Two EOMs are needed for orthogonal modulation and dual-channel imaging. Resonant version is recommended so lower driving voltage can be used.</td>
		</tr>
		<tr>
			<td>False H&#38;E Staining Script</td>
			<td>Matlab</td>
			<td></td>
			<td>https://github.com/TheFuGroup/HE_Staining</td>
		</tr>
		<tr>
			<td>Fanout Buffer</td>
			<td>PRL-414B</td>
			<td>Pulse Research Lab</td>
			<td>1:4 TTL/CMOS Fanout Buffer and Line Driver, for generating the EOM driving frequency and the reference to the lock-in</td>
		</tr>
		<tr>
			<td>Fast Photodiode</td>
			<td>THORLABS</td>
			<td>DET10A2</td>
			<td>Si Detector, 1 ns Rise Time</td>
		</tr>
		<tr>
			<td>Frequency Divider</td>
			<td>PRL-220A</td>
			<td>Pulse Research Lab</td>
			<td>TTL Freq. Divider (f/2, f/4, f/8, f/16), for generating 20MHz from the laser output.</td>
		</tr>
		<tr>
			<td>Highly Dispersive Glass Rods</td>
			<td>Union Optic</td>
			<td>CYLROD01</td>
			<td>High dispersion H-ZF52A Rod lens 120 mm, SF11 Rod lens 100 mm</td>
		</tr>
		<tr>
			<td>Insight DS+</td>
			<td>Newport</td>
			<td></td>
			<td>Laser system capable of outputting two synchronzied pulsed lasers (one fixed beam at 1, 040 nm and one tunable beam, ranging from 680-1,300 nm) with a repetition rate of 80 MHz.&#160;</td>
		</tr>
		<tr>
			<td>Lock-in Amplifier</td>
			<td>Liquid Instruments</td>
			<td>Moku Lab</td>
			<td>Lock-in amplifier to extract SRS signal from the photodiode. A Zurich Instrument HF2LI or similar instrument can be used as well.</td>
		</tr>
		<tr>
			<td>Mirrors</td>
			<td>THORLABS</td>
			<td>BB05-E03-10</td>
			<td>Broadband Dielectric Mirror, 750 - 1,100 nm. Silver mirrors can also be used.</td>
		</tr>
		<tr>
			<td>Motorized Delay Stage</td>
			<td>Zaber</td>
			<td>X-DMQ12P-DE52</td>
			<td>Delay stage for fine control of the temporal overlap of the pump and the Stokes lasers. Any other motorized stage should work.</td>
		</tr>
		<tr>
			<td>Oil Immersion Condensor</td>
			<td>Nikon</td>
			<td>CSC1003</td>
			<td>1.4 NA. Other condensers with NA&#62;1.2 can be used.</td>
		</tr>
		<tr>
			<td>Oscilloscope</td>
			<td>Tektronix</td>
			<td>TDS7054</td>
			<td>Any other oscilloscope with 400 MHz bandwdith or higher should work.</td>
		</tr>
		<tr>
			<td>Phase Shifter</td>
			<td>SigaTek</td>
			<td>SF50A2</td>
			<td>For shifting the phase of the modulation frequency</td>
		</tr>
		<tr>
			<td>Photodiode</td>
			<td>Hamamatsu Corp</td>
			<td>S3994-01</td>
			<td>Silicon PIN diode with large area (10 x 10 cm2). Other diodes with large area and low capacitance can be used.</td>
		</tr>
		<tr>
			<td>Polarizing Beam Splitter</td>
			<td>Union Optic</td>
			<td>PBS9025-620-1000</td>
			<td>Broadband polarizing beamsplitter</td>
		</tr>
		<tr>
			<td>Refactive Index Database</td>
			<td></td>
			<td></td>
			<td>refractiveindex.info</td>
		</tr>
		<tr>
			<td>Retro-reflector</td>
			<td>Edmund Optics</td>
			<td>34-408</td>
			<td>BBAR Right Angle Prism. Other prisms or retroreflector can be used.</td>
		</tr>
		<tr>
			<td>RF Power Amplifier</td>
			<td>Minicircuits</td>
			<td>ZHL-1-2W+</td>
			<td>Gain Block, 5 - 500 MHz, 50 &#937;</td>
		</tr>
		<tr>
			<td>Scan Mirrors</td>
			<td>Cambridge Technologies</td>
			<td>6215H</td>
			<td>We used a 5mm mirror set with silver coating</td>
		</tr>
		<tr>
			<td>ScanImage</td>
			<td>Vidrio</td>
			<td>ScanImage Basic</td>
			<td>Laser scanning microscope control software</td>
		</tr>
		<tr>
			<td>Shortpass Filter</td>
			<td>THORLABS</td>
			<td>FESH1000</td>
			<td>25.0 mm Premium Shortpass Filter, Cut-Off Wavelength: 1,000 nm. For efficient suppression of the Stokes, two filters may be necessary.</td>
		</tr>
		<tr>
			<td>Upright Microscope</td>
			<td>Nikon</td>
			<td>Eclipse FN1</td>
			<td>Any other microscope frame can be used. If a laser scanning microscope is available, it can be used directly. Otherwise, a galvo scanner and scan lens needed to be added to the microscope.</td>
		</tr>
		<tr>
			<td>Water Immersion Objective</td>
			<td>Olympus</td>
			<td>XLPLN25XWMP2</td>
			<td>The multiphoton 25X Objective has a NA of 1.05. Other similar objectives can be used.</td>
		</tr>
	</tbody>
</table>

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.

Optimizing spectral resolution: 
Dispersion through a material is affected by the dispersive medium (length and material) and wavelength. Changing the dispersion rod length affects the spectral resolution and the signal size. It is a give-and-take relationship that can be weighed differently depending on the application. The rods stretch out the beam pulse from being wide in frequency and narrow in time to being narrow in frequency and broad in time. Figure 7 shows the ...

Watch this Scientific Journal Video about Real-Time, Two-Color Stimulated Raman Scattering Imaging of Mouse Brain for Tissue Diagnosis at JoVE.com

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

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 ...

This protocol will demonstrate implementation and optimization of spectral focusing SRS microscopy and how it can be used for real-time stimulated Raman histology applications. The main advantage of this technique is the speed at which samples can be processed once the instrumentation is properly aligned, as SRS does not require tissue processing and staining. This protocol is applicable to other SRS applications, not just limited to histology.Such applications include small molecules, drugs, cells, and tissues. Begin by installing a pair of achromatic lenses for the pump beam to magnify the laser beam size by twofold as a starting point. Place a 100 and 200 millimeter lens at roughly 300 millimeters from each other.Then align the pump beam through the center of both lenses. Next place a mirror after the second lens to send the beam towards a wall more than one meter away. Trace the beam from the mirror to the wall with an IR card and check if the beam changes in size.Collimate the beam if the beam changes in size as a function of distance. Combine the two laser beams by installing a dichroic mirror with several steering mirrors for adjustment. Optimize spatial overlap of the pump and Stokes by monitoring beams after the dichroic mirror at two different positions around one meter apart.Iteratively adjust the steering mirror followed by the dichroic mirror to align the Stokes beam with the pump beam. Ensure that the combined beams are sent to the center of the scan mirrors of the laser scanning microscope by adjusting a pair of steering mirrors when the scan mirrors are in the parked position. After the condenser, use another pair of lenses with focal lengths of 100 millimeters and 30 millimeters, respectively, to relay the transmitted beam onto the photodiode, ensuring both beams are contained within the photodiode.Then install two low pass filters to block out the modulated Stokes beam. Place a beam sampler in the Stokes beam path to pick up 10%of the beam and send it to a fast photodiode to detect the 80 megahertz pulse train. Take one of the outputs of the fanout buffer and filter it with a band pass filter to obtain a 20 megahertz sinusoidal wave.Then use an RF attenuator to adjust the output peak to peak voltage to approximately 500 millivolts. Send the resulting output to a phase shifter, which allows fine adjustment of the RF phase with a voltage source. Send this output to an RF power amplifier and connect the output of the amplifier to the EOM.After unblocking the Stokes beam, optimize the modulation depth of the first EOM by placing a photodiode in the beam path. Adjust the EOM voltage and quarter wave plate until the modulation depth appears satisfactory. Place a photodiode after the dichroic mirror to detect the laser beam.Block the Stokes beam first, then zoom in on one of the pump pulse peaks on the oscilloscope. Place a vertical cursor to mark the temporal position of this peak with the oscilloscope. Now block the pump beam, and unblock the Stokes beam.Translate the delay stage to temporarily match the peak positions on the oscilloscope to the marked position in the previous step. By calculating the delay distance required to match the two beams followed by elongating the beam path of the faster beam or shortening the beam path of the slower beam. Next prepare a microscope slide sample with DMSO and double-sided tape as a spacer to hold the sample between the slide and a coverslip.Place the sample on the microscope with the coverslip side facing the microscope objective and observe the sample from the I piece under brightfield illumination. Find the focus of the sample at both the top and bottom layer of air bubbles at the glass tape interface, then move the focus to be in between the two layers of the tape. Next set the tunable beam output to 798 nanometers.Based on the optical throughput of the condenser, adjust the optical power to be approximately 40 milliwatts each for the pump and Stokes beams at the objective focus. Then open scan image in MATLAB and click on the button labeled focus to start scanning. Adjust the steering mirror before the galvo scanner to center the DC signal on the channel one display.Move the motorized delay stage and closely observe the lock-in output shown on the channel two display. Finally adjust the time delay to maximize the AC signal intensity. Adjust the dichroic mirror to center the SRS signal on the AC channel.Then adjust the phase of the lock-in amplifier to maximize the signal. After mounting the sample slide onto the microscope and adjusting the power of both beams to approximately 40 milliwatts each at sample focus as demonstrated earlier. Open scan image from MATLAB.Then find the maximal SRS signal by scanning through the delay stage corresponding to the 2, 913 inverse centimeter Raman peak of DMSO. Acquire an SRS image corresponding to the 2, 913 inverse centimeter Raman peak of DMSO. Open the image in ImageJ and select a small area in the center of the frame.Use the measure function to calculate the mean and standard deviation of values in the selected area. For frequency access calibration, save a hyperspectral scan with the delay stage range covering the 2, 913 and 2, 994 inverse centimeter Raman peaks of DMSO. Then save the stage positions corresponding to the hyperspectral data set.Perform linear regression for the stage positions and Raman shifts at 2, 913 and 2, 994 inverse centimeters. Using the linear regression equation, convert the delay positions to the corresponding Raman frequencies. For calibration of spectral resolution, estimate the stage position based on the previous position with the increased optical path length due to the insertion of rods.Realign the beams spatial overlap because of the slight deviation of the beam when glass rods are added. Install a polarizing beam splitter, a quarter wave plate, and a second EOM into the soaks beam path after the first EOM. Unplug the signal input to the second EOM, then plug in the signal input to the first EOM, and turn it on.Modulate the Stokes beam at 20 megahertz by sending the beam through the first EOM. Adjust the tilt and position of the first EOM to ensure that the beam is centered through the EOM crystal. Prepare a tissue slide sample with double-sided tape, microscope slide and cover glass.Place the sample on the microscope with the coverslip side facing the microscope objective. For epi-mode SRS imaging, install a half-wave plate before the beam enters the microscope to change the polarization of the beam going into the microscope. Place a polarizing beam splitter above the objective to allow the depolarized back reflected beam to reach the detector.Next, use a 75 millimeter achromate lens and a 30 millimeter aspheric lens to relay the back scattered photons from the back aperture of the objective to the photo detector. Mount the detector to collect the back scattered light directed by the polarizing beam splitter. Then install a filter to block out the modulated beam from entering the detector.Varying the rod lengths affect spectral resolution. When no glass chirping rods are used, the two Raman peaks from DMSO are not resolved at all. An increase in the number of glass chirping rods starts to resolve the two peaks at a satisfactory point.Matched chirping resolves both peaks and can be used to calibrate stage positions to frequency. Two time delayed pulse trains of orthogonal polarization used to image DMSO spectra show the time delay between the SRS excitations with acceptable modulation depth and temporal separation. In contrast, a poor two color SRS phase shift, gives rise to inverted or negative peaks.Real-time two color SRS imaging at 2, 850 and 2, 930 inverse centimeters of ex vivo mouse brain tissue are shown here. The raw lipid and protein images were color coded to produce a single image depicting lipid and protein contributions. False hematoxylin and eosin recoloring, were also performed to mimic hematoxylin and eosin staining for pathological applications.Spatial temporal overlap and spatial resolution optimization are the most important steps for the spatial focusing approach of SRS. This method is generally applicable for other pump probe microscopy experiments such as transient absorption microscopy. The method also allows for imaging of non-fluorescent molecules in cells and tissue.The method presented here speeds up the image acquisition time for future translation of stimulated Raman histology in the clinic.

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Research

JoVE Journal

Bioengineering

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, ...

Universal Hand-held Three-dimensional Optoacoustic Imaging Probe for Deep Tissue Human Angiography and Functional Preclinical Studies in Real Time

The exclusive combination of high optical contrast and excellent spatial resolution makes optoacoustics (photoacoustics) ideal for simultaneously attaining anatomical, functional and molecular contrast in deep optically opaque tissues. While enormous potential has been recently demonstrated in the application of optoacoustics for small animal research, vast efforts have also been undertaken in translating this imaging technology into clinical practice. We present here a newly developed optoacoustic tomography approach capable of delivering high resolution and spectrally enriched volumetric images of tissue morphology and function in real time. A detailed description of the experimental protocol for operating with the imaging system in both hand-held and stationary modes is provided and showcased for different potential scenarios involving functional and molecular studies in murine models and humans. The possibility for real time visualization in three dimensions along with the versatile handheld design of the imaging probe make the newly developed approach unique among the pantheon of imaging modalities used in today&#8217;s preclinical research and clinical practice.

We provide herein a detailed description of the experimental protocol for imaging with a newly developed hand-held optoacoustic (photoacoustic) system for three-dimensional functional and molecular imaging in real time. The demonstrated powerful performance and versatility may define new application areas of the optoacoustic technology in preclinical research and clinical practice.

The exclusive combination of high optical contrast and excellent spatial resolution makes optoacoustics (photoacoustics) ideal for simultaneously ...

Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and other matrix-modifying oxidants/enzymes released during inflammation) are implicated in triggering pathological changes in cellular function, phenotype and viability in a number of diseases. Here, we describe how cell-substrate impedance and live cell imaging approaches can be readily employed to accurately quantify real-time changes in cell adhesion and de-adhesion induced by matrix modification (using endothelial cells and myeloperoxidase as a pathophysiological matrix-modifying stimulus) with high temporal resolution and in a non-invasive manner. The xCELLigence cell-substrate impedance system continuously quantifies the area of cell-matrix adhesion by measuring the electrical impedance at the cell-substrate interface in cells grown on gold microelectrode arrays. Image analysis of time-lapse differential interference contrast movies quantifies changes in the projected area of individual cells over time, representing changes in the area of cell-matrix contact. Both techniques accurately quantify rapid changes to cellular adhesion and de-adhesion processes. Cell-substrate impedance on microelectrode biosensor arrays provides a platform for robust, high-throughput measurements. Live cell imaging analyses provide additional detail regarding the nature and dynamics of the morphological changes quantified by cell-substrate impedance measurements. These complementary approaches provide valuable new insights into how myeloperoxidase-catalyzed oxidative modification of subcellular extracellular matrix components triggers rapid changes in cell adhesion, morphology and signaling in endothelial cells. These approaches are also applicable for studying cellular adhesion dynamics in response to other matrix-modifying stimuli and in related adherent cells (e.g., epithelial cells).

Here, we present a protocol to continuously quantify cell adhesion and de-adhesion processes with high temporal resolution in a non-invasive manner by cell-substrate impedance and live cell imaging analyses. These approaches reveal the dynamics of cell adhesion/de-adhesion processes triggered by matrix modification and their temporal relationship to adhesion-dependent signaling events.

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and ...

Real-time Monitoring of High Intensity Focused Ultrasound (HIFU) Ablation of In Vitro Canine Livers Using Harmonic Motion Imaging for Focused Ultrasound (HMIFU)

Harmonic Motion Imaging for Focused Ultrasound (HMIFU) is a technique that can perform and monitor high-intensity focused ultrasound (HIFU) ablation. An oscillatory motion is generated at the focus of a 93-element and 4.5 MHz center frequency HIFU transducer by applying a 25 Hz amplitude-modulated signal using a function generator. A 64-element and 2.5 MHz imaging transducer with 68kPa peak pressure is confocally placed at the center of the HIFU transducer to acquire the radio-frequency (RF) channel data. In this protocol, real-time monitoring of thermal ablation using HIFU with an acoustic power of 7 W on canine livers in vitro is described. HIFU treatment is applied on the tissue during 2 min and the ablated region is imaged in real-time using diverging or plane wave imaging up to 1,000 frames/second. The matrix of RF channel data is multiplied by a sparse matrix for image reconstruction. The reconstructed field of view is of 90&#176; for diverging wave and 20 mm for plane wave imaging and the data are sampled at 80 MHz. The reconstruction is performed on a Graphical Processing Unit (GPU) in order to image in real-time at a 4.5 display frame rate. 1-D normalized cross-correlation of the reconstructed RF data is used to estimate axial displacements in the focal region. The magnitude of the peak-to-peak displacement at the focal depth decreases during the thermal ablation which denotes stiffening of the tissue due to the formation of a lesion. The displacement signal-to-noise ratio (SNRd) at the focal area for plane wave was 1.4 times higher than for diverging wave showing that plane wave imaging appears to produce better displacement maps quality for HMIFU than diverging wave imaging.

Real-time Monitoring of High Intensity Focused Ultrasound HIFU Ablation of In Vitro Canine Livers Using Harmonic Motion Imaging for Focused Ultrasound HMIFU

This article describes real-time monitoring of HIFU ablation in canine liver with high frame rate ultrasound imaging using diverging and plane wave imaging. Harmonic Motion Imaging for Focused Ultrasound is used to image the decrease of acoustic radiation force induced displacement in the ablated region.

Harmonic Motion Imaging for Focused Ultrasound (HMIFU) is a technique that can perform and monitor high-intensity focused ultrasound (HIFU) ablation. An ...

Hand-held Clinical Photoacoustic Imaging System for Real-time Non-invasive Small Animal Imaging

Translation of photoacoustic imaging into the clinic is a major challenge. Handheld real-time clinical photoacoustic imaging systems are very rare. Here, we report a combined photoacoustic and clinical ultrasound imaging system by integrating an ultrasound probe with light delivery for small animal imaging. We demonstrate this by showing sentinel lymph node imaging in small animals along with minimally invasive real-time needle guidance. A clinical ultrasound platform with access to raw channel data allows the integration of photoacoustic imaging leading to a handheld real-time clinical photoacoustic imaging system. Methylene blue was used for sentinel lymph node imaging at 675 nm wavelength. Additionally, needle guidance with dual modal ultrasound and photoacoustic imaging was shown using the imaging system. Depth imaging of up to 1.5 cm was demonstrated with a 10 Hz laser at a photoacoustic imaging frame rate of 5 frames per second.

A clinical handheld photoacoustic imaging system will be demonstrated for real-time non-invasive small animal imaging.

Translation of photoacoustic imaging into the clinic is a major challenge. Handheld real-time clinical photoacoustic imaging systems are very rare. Here, ...

A Protocol for Real-time 3D Single Particle Tracking

Real-time three-dimensional single particle tracking (RT-3D-SPT) has the potential to shed light on fast, 3D processes in cellular systems. Although various RT-3D-SPT methods have been put forward in recent years, tracking high speed 3D diffusing particles at low photon count rates remains a challenge. Moreover, RT-3D-SPT setups are generally complex and difficult to implement, limiting their widespread application to biological problems. This protocol presents a RT-3D-SPT system named 3D Dynamic Photon Localization Tracking (3D-DyPLoT), which can track particles with high diffusive speed (up to 20 &#181;m2/s) at low photon count rates (down to 10 kHz). 3D-DyPLoT employs a 2D electro-optic deflector (2D-EOD) and a tunable acoustic gradient (TAG) lens to drive a single focused laser spot dynamically in 3D. Combined with an optimized position estimation algorithm, 3D-DyPLoT can lock onto single particles with high tracking speed and high localization precision. Owing to the single excitation and single detection path layout, 3D-DyPLoT is robust and easy to set up. This protocol discusses how to build 3D-DyPLoT step by step. First, the optical layout is described. Next, the system is calibrated and optimized by raster scanning a 190 nm fluorescent bead with the piezoelectric nanopositioner. Finally, to demonstrate real-time 3D tracking ability, 110 nm fluorescent beads are tracked in water.

This protocol details the construction and operation of a real-time 3D single particle tracking microscope capable of tracking nanoscale fluorescent probes at high diffusive speeds and low photon count rates.

Real-time three-dimensional single particle tracking (RT-3D-SPT) has the potential to shed light on fast, 3D processes in cellular systems. Although ...

In Situ Microscopy for Real-time Determination of Single-cell Morphology in Bioprocesses

In situ monitoring in microbial bioprocesses is mostly restricted to chemical and physical properties of the medium (e.g.,pH value and the dissolved oxygen concentration). Nevertheless, the morphology of cells can be a suitable indicator for optimal conditions, since it changes with dependence on the growth state, product accumulation and cell stress. Furthermore, the single-cell size distribution provides not only information about the cultivation conditions, but also about the population heterogeneity. To gain such information, a photo-optical in situ microscopy device1 was developed to enable the monitoring of the single-cell size distribution directly in the cell suspension in bioreactors. An automated image analysis is coupled to the microscopy based on a neural network model, which is trained with user-annotated images. Several parameters, which are gained from the captures of the microscope, are correlated to process relevant features of the cells, like their metabolic activity. Until now, the presented in situ microscopy probe series was applied to measure the pellet size in filamentous fungi suspensions. It was used to distinguish the single-cell size in microalgae cultivation and relate it to lipid accumulation. The shape of cellular particles was related to budding in yeast cultures. The microscopy analysis can be generally split into three steps: (i) image acquisition, (ii) particle identification, and (iii) data analysis, respectively. All steps have to be adapted to the organism, and therefore specific annotated information is required in order to achieve reliable results. The ability to monitor changes in cell morphology directly in line or on line (in a by-pass) enables real-time values for monitoring and control, in process development as well as in production scale. If the off line data correlates with the real-time data, the current tedious off line measurements with unknown influences on the cell size become needless.

In Situ Microscopy for Real-time Determination of Single-cell Morphology in Bioprocesses

A photo-optical in situ microscopy device was developed to monitor the size of single cells directly in the cell suspension. The real-time measurement is conducted by coupling the photo-optical sterilizable probe to an automated image analysis. Morphological changes appear with dependence on the growth state and cultivation conditions.

In situ monitoring in microbial bioprocesses is mostly restricted to chemical and physical properties of the medium (e.g.,pH value and the dissolved ...

In Vivo Two-Color 2-Photon Imaging of Genetically-Tagged Reporter Cells in the Skin

Fibroblasts are mesenchymal cells that change their morphology upon activation, ultimately influencing the microenvironment of the tissue they are located in. Although traditional imaging techniques are useful in identifying protein interactions and morphology in fixed tissue, they are not able to give insight as to how quickly cells are able to bind and uptake proteins, and once activated how their morphology changes in vivo. In the present study, we ask 2 major questions: 1) what is the time-course of fibroblast activation via toll-like receptor-4 (TLR4) and lipopolysaccharide (LPS) interaction and 2) how do these cells behave once activated? Using 2-photon imaging, we have developed a novel technique to assess the ability of LPS-FITC to bind to its cognate receptor, TLR4, expressed on peripheral fibroblasts in the genetic reporter mouse line; FSP1cre; tdTomatolox-stop-lox&#160;in vivo. This unique approach allows researchers to create in-depth, time-lapse videos and/or pictures of proteins interacting with live cells that allows one to have an increased level of granularity in understanding how proteins can alter cellular behavior.

Morphological changes occur in immune responsive fibroblast cells following activation and promote alterations in cellular recruitment. Utilizing 2-photon imaging in conjunction with a genetically engineered Fibroblast-specific protein 1 (FSP1)-cre; tdTomato floxed-stop-floxed (TB/TB) mouse line and green fluorescently tagged lipopolysaccharide-FITC, we can illustrate highly specific uptake of lipopolysaccharide in dermal fibroblasts and morphological changes in vivo.

Fibroblasts are mesenchymal cells that change their morphology upon activation, ultimately influencing the microenvironment of the tissue they are located ...

Real-Time Intravital Multiphoton Microscopy to Visualize Focused Ultrasound and Microbubble Treatments to Increase Blood-Brain Barrier Permeability

The blood-brain barrier (BBB) is a key challenge for the successful delivery of drugs to the brain. Ultrasound exposure in the presence of microbubbles has emerged as an effective method to transiently and locally increase the permeability of the BBB, facilitating para- and transcellular transport of drugs across the BBB. Imaging the vasculature during ultrasound-microbubble treatment will provide valuable and novel insights on the mechanisms and dynamics of ultrasound-microbubble treatments in the brain.
Here, we present an experimental procedure for intravital multiphoton microscopy using a cranial window aligned with a ring transducer and a 20x objective lens. This set-up enables high spatial and temporal resolution imaging of the brain during ultrasound-microbubble treatments. Optical access to the brain is obtained via an open-skull cranial window. Briefly, a 3-4 mm diameter piece of the skull is removed, and the exposed area of the brain is sealed with a glass coverslip. A 0.82 MHz ring transducer, which is attached to a second glass coverslip, is mounted on top. Agarose (1% w/v) is used between the coverslip of the transducer and the coverslip covering the cranial window to prevent air bubbles, which impede ultrasound propagation. When sterile surgery procedures and anti-inflammatory measures are taken, ultrasound-microbubble treatments and imaging sessions can be performed repeatedly over several weeks. Fluorescent dextran conjugates are injected intravenously to visualize the vasculature and quantify ultrasound-microbubble induced effects (e.g., leakage kinetics, vascular changes). This paper describes the cranial window placement, ring transducer placement, imaging procedure, common troubleshooting steps, as well as advantages and limitations of the method.

This protocol describes the surgical and technical procedures that enable real-time in vivo multiphoton fluorescence imaging of the rodent brain during focused ultrasound and microbubble treatments to increase blood-brain barrier permeability.

The blood-brain barrier (BBB) is a key challenge for the successful delivery of drugs to the brain. Ultrasound exposure in the presence of microbubbles ...

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 ...