This work was supported by the Hong Kong Research Grants Council through grants 16103215, 16148816, 16102518, 16102920, T13-607/12R, T13-706/11-1, T13-605/18W, C6002-17GF, C6001-19E, N_HKUST603/19, the Innovation and Technology Commission (ITCPD/17-9), the Area of Excellence Scheme of the University Grants Committee (AoE/M-604/16, AOE/M-09/12), and the Hong Kong University of Science &#38; Technology (HKUST) through grant RPC10EG33.

All animal procedures performed in this work are conducted according to the guidelines of the Laboratory Animal Facility of the Hong Kong University of Science and Technology (HKUST) and have been approved by the Animal Ethics Committee of HKUST. Safety training for laser handling is required to set up and operate the TPEF-SRS microscope. Always wear laser safety goggles with appropriate wavelength range when dealing with laser.
1. Setup of the TPEF-SRS microscope (for setup schematic see Figure 1)
<ol>
	<li>Use an integrated optical parametric oscillator (OPO) connected with a mode-locked Ytterbium fiber laser as the ps&#160;laser source for SRS imaging. 
	NOTE: OPO outputs a Stokes beam (1031 nm) and a pump beam (tunable from 780 nm to 960 nm) with 2 ps pulse duration and 80 MHz repetition rate. The Stokes beam is modulated at 20 MHz by a built-in electro-optical modulator (EOM) for&#160;high-frequency&#160;phase-sensitive SRS detection at MHz level.</li>
	<li>Use an fs titanium (Ti): sapphire laser as the laser source for TPEF imaging.</li>
	<li>Use a pair of lenses (L1 and L2) to collimate and adjust the size of the fs beam to 3 mm.</li>
	<li>Use a pair of lenses (L3 and L4) to collimate the ps laser beam and expand its diameter to 3 mm.</li>
	<li>Change the polarization of the fs laser beam from p polarization to s polarization using a half-wave plate.</li>
	<li>Combine the two laser beams with a polarizing beam splitter (PBS).</li>
	<li>Add a pair of 3 mm XY-scan galvanometer mirrors behind the PBS for beam scanning.</li>
	<li>Use a telecentric scan lens (L5) and an infinity-corrected tube lens (L6) to conjugate the scanning mirror and the rear pupil of the 25x objective lens. Expand the laser beam by the scan lens and tube lens to fill the back aperture of the objective.</li>
	<li>Place a PBS or dichroic mirror (D2) between the tube lens and objective lens for SRS or fluorescence signal collection. Use a motorized flipper to switch between PBS and D2. 
	&#8203;NOTE: Part of the back-scattered SRS signal is lost when passing through the PBS as a result of its randomly shifted polarization.</li>
	<li>Use a pair of lenses (L7 and L9) to narrow the detection beam and conjugate the rear pupil of the 25x objective lens with a photodiode sensor.</li>
	<li>Use a pair of lenses (L8 and L10) to narrow the detection beam and conjugate the rear pupil of the 25x objective with the detection surface of the photomultiplier (PMT).</li>
	<li>Use a dichroic mirror (D3) to separate the detection path of fluorescence and SRS signals.</li>
	<li>Place a filter set (Fs1) in front of the photodiode detector to block the Stokes laser and pass the pump beam only.</li>
	<li>Place a filter set (Fs2) in front of the PMT detector to pass the target fluorescence signal only.</li>
	<li>Connect the PMT to a current amplifier for signal amplification.</li>
	<li>Connect the photodiode to a lock-in amplifier.</li>
	<li>Connect the sync signal from the built-in EOM output to the reference input of the lock-in amplifier for SRS signal demodulation.</li>
	<li>Connect the PMT amplifier and lock-in amplifier outputs to the data acquisition module.</li>
</ol>
2. TPEF-SRS microscope system calibration
<ol>
	<li>Startup of the lasers
	<ol>
		<li>Switch the key switch from Standby to On position to turn on the Ti: sapphire laser and wait for 30 min for the laser to warm up.</li>
		<li>Switch on the OPO by clicking on the Start button on the OPO control panel and wait for 20 min for the laser to warm up.</li>
		<li>After the ps laser warms up, use a high-speed photodetector to check the modulation depth of the Stokes beam. Open the laser shutter for the Stokes beam. Click on the Set OPO Power box&#160;and enter 20. Place the high-speed photodetector at the OPO output to detect the beam. Connect the output port of the photodetector to the input port of an oscilloscope using a coaxial cable with Bayonet Neill-Concelman (BNC) connector to monitor the laser pulse.</li>
		<li>Open the EOM Control window in the OPO control software. Adjust the EOM power and phase according to the pulse intensity diagram shown on the oscilloscope to achieve maximal modulation depth at 20 MHz. 
		NOTE: EOM performance is usually stable and only needs to be checked when the SRS signal is found significantly decreased.</li>
	</ol>
	</li>
	<li>Optical alignment of the combined TPEF-SRS microscope
	<ol>
		<li>Perform optical alignment for colocalization of the ps and fs beams as described in steps 2.2.2 to 2.2.13.</li>
		<li>Open the pump laser shutter while stopping the Stokes output in the OPO control software. Use OPO control software to set the wavelength of the pump beam to 796 nm by clicking the Set Signal box and entering the wavelength value 796. Click on the Set OPO Power&#160;box and enter 20 to set its power to the minimum (~20 mW) for optical alignment.</li>
		<li>Switch on the microscope computer and all associated electronic components, including scanners, objective actuators, photodiodes, PMTs, current amplifiers, lock-in amplifiers, and motorized flippers. Start the microscope control software. 
		NOTE: The microscope control software is a homemade interface.</li>
		<li>Place two alignment plates (P1 and P2 in Figure 1) on the optical path. Place P1 behind PBS at a distance of about 10 cm and place P2 behind P1 at a distance of about 30 cm. 
		NOTE: The ps and fs beams are combined using a PBS (Figure 1). The two alignment plates are used to check the alignment as well as the colocalization of the two laser beams.</li>
		<li>Open the microscope shutter for the ps laser beam.</li>
		<li>Adjust the mirror M1 to locate the ps laser beam center at the through-hole of P1. Use an infrared (IR) scope to observe the position of the beam spot at P1 when adjusting the mirror M1.</li>
		<li>Adjust the mirror M2 to locate the ps laser beam center at the through-hole of P2. Use the IR scope to observe the position of the beam spot at P2 when adjusting the mirror M2.</li>
		<li>Repeat steps 2.2.6 and 2.2.7 until the ps beam center locates at the through-hole of both alignment plates. Close the shutter of the ps beam in the microscope control software.</li>
		<li>Set the wavelength of fs Ti: sapphire laser at 740 nm and open the laser shutter. Set the laser power to 5 mW (measured at the input port of the microscope system) for optical alignment.</li>
		<li>Open the microscope shutter for the fs laser beam.</li>
		<li>Adjust the mirror M3 to locate the fs laser beam spot center at the through-hole of P1.</li>
		<li>Adjust the mirror M4 to locate the laser beam spot center at the through-hole of P2.</li>
		<li>Repeat steps 2.2.11 and 2.2.12 until the fs laser beam center locates at the through-holes of two alignment plates. Close the microscope shutter for the fs beam.</li>
		<li>Perform spatial overlapping of the pump and Stokes beams as described in steps 2.2.15 to 2.2.18. 
		&#8203;NOTE: Although the pump and Stokes beams are roughly overlapped inside the ps laser, fine-tuning of the positions of the two laser beams is required to achieve optimal SRS performance. Since the pump beam is firstly aligned as previously described, next the Stokes beam is adjusted to colocalize with the pump beam.</li>
		<li>Place a camera at the position of the objective to visualize the location of two beam spots. Mark the position of the pump beam on the camera screen as a reference.</li>
		<li>Place an alignment plate P0 before L3 (Figure 1). Use a hex key to adjust the optical mirror 1 (OM1) so that the Stokes beam center passes the through-hole of the alignment plate and colocalizes with the pump beam at the laser output port. Use the IR scope to confirm the position of the beam spot at P0 during adjustment. 
		NOTE: OM1 and OM2 are two mirrors in the OPO head to adjust the position of the 1031 nm Stokes beam.</li>
		<li>Remove the alignment plate P0 and use the hex key to adjust the OM2 so that the center of the Stokes beam colocalizes with the reference mark of the pump beam on the camera.</li>
		<li>Repeat steps 2.2.16 and 2.2.17 until the Stokes beam strictly overlaps with the pump beam at both P0 and camera planes. 
		NOTE: When visualizing the beam spots on the camera, all the alignment plates should be removed from the optical path.</li>
	</ol>
	</li>
	<li>Optimize imaging conditions
	<ol>
		<li>Perform phase adjustment of the lock-in amplifier as described in steps 2.3.2. to 2.3.7. 
		NOTE: SRS detection is based on a high-frequency phase-sensitive scheme. For SRL detection, the intensity of the Stokes beam is modulated at 20 MHz and a lock-in amplifier is used to demodulate the signal. The phase and offset of the lock-in amplifier need to be adjusted to achieve the optimal image contrast.</li>
		<li>Open the lock-in amplifier control software.</li>
		<li>Set the wavelength of the pump laser to 796 nm and the power of the pump and Stokes beam to be 15 mW and 25 mW on the sample, respectively. 
		NOTE: Here use olive oil for SRS imaging optimization and calibration. The Raman peak of olive oil at the carbon-hydrogen region is at 2863.5 cm-1, corresponding to the pump beam wavelength at 796 nm.</li>
		<li>Seal the olive oil in a slide and attach a folded tissue paper to the bottom of the slide to enhance the signal backscattering for epi-SRS detection. Place the olive oil sample on the stage and adjust the focus of the 25x objective onto the sample.</li>
		<li>Use the microscope control software to set the imaging parameters as follows: 512 x 512 pixels for 500 &#956;m x 500 &#956;m field of view (FOV), 6.4 &#956;s pixel dwell time. Use the lock-in amplifier control software to set the time constant value to be 10 &#956;s, which is close to the pixel dwell time.</li>
		<li>Scan the laser beam over the sample. Use the lock-in amplifier control software to tune the phase (0-180&#176;) with a step size of 22.5&#176; until the SRS signal intensity reaches the maximum. 
		NOTE: In this all-analog lock-in amplifier, the signal output is the in-phase component, which is dependent on the phase of the reference signal. The phase can be adjusted by the lock-in amplifier control software with 11&#176; resolution, allowing to maximize the detected signal to within ~2%12. The lock-in amplifier gets out of phase once the sync signal is disrupted. The phase needs to be readjusted every time the sync signal is reestablished.</li>
		<li>Scan the sample with laser shutter closed. Use the lock-in amplifier control software to tune the offset value with a step size of 1 mV until the average SRS signal is close to zero. 
		NOTE: The average SRS signal is estimated as the average intensity of all pixels in the SRS image, which is calculated automatically by the microscope control software. Offsets are useful for canceling unwanted phase-coherent signals.</li>
		<li>Perform temporal synchronization optimization as described in steps 2.3.9 to 2.3.14.</li>
		<li>Open the Delay Manager dialogue (Figure 2) in the OPO control software.</li>
		<li>Scan the olive oil and tune the delay stage until the olive oil SRS signal reaches its maximum. 
		NOTE: For the first time of SRS imaging, when the phase of the lock-in amplifier has not been optimized, temporal synchronization is first roughly adjusted to visualize the SRS signal of the sample before optimizing the phase of the lock-in amplifier.</li>
		<li>Stop scanning and click on the Add Data button in the delay manager dialogue to record the current delay data.</li>
		<li>Repeat steps 2.3. 10 and 2.3.11 for different chemical samples at different wavelengths. 
		NOTE: Polystyrene beads, heavy water, 5-Ethynyl-2&#8242;-deoxyuridine, glycerol are used to measure the delay data at different vibrational bands.</li>
		<li>Select the Data Fit and Order 5 on the delay manager dialogue to fit the current data points with the fifth-order polynomic function. Apply the fitted data by clicking on the Use As Custom button and checking the check box. The delay stage will be auto-adjusted at different wavelengths according to the fitted delay curve.</li>
		<li>Save all the delay data in a text file, which can be loaded for future use.</li>
	</ol>
	</li>
</ol>
3. Surgical preparation of mouse for in vivo fluorescence and SRS imaging
<ol>
	<li>Sterilize all required tools, including scalpels, spring scissors, forceps, coverslips, and gauze pads.</li>
	<li>Disinfect all the surfaces, which would be touched during surgery with 70% ethanol. Cover the working area of the benchtop with sterile drapes. Put a heating pad under the drape.</li>
	<li>Use Thy1-YFP-H (Tg(Thy1-YFP)HJrs/J)37 transgenic mice that express enhanced yellow fluorescence protein (EYFP) in dorsal root ganglion afferent neurons for in vivo spinal cord imaging. Weigh the mouse and induce anesthetization by intraperitoneal (i.p.) injection of the ketamine-xylazine mixture (87.5 mg kg-1 and 12.5 mg kg-1).</li>
	<li>Pinch the toe of the mouse to ensure deep anesthesia. Supplement with half the original dose of the anesthetics if needed. Apply ointment to the mouse eyes to prevent corneal dryness.</li>
	<li>Shave the hair on the back above the thoracic spine, and then completely remove the hair using depilating cream. Disinfect the shaved area with iodine solution.</li>
	<li>Make a small midline incision (~1.5 cm) of the skin over the T11-T13 vertebra with a scalpel.</li>
	<li>Incise&#160;muscles and tendons both on top and on sides of the T11-T13 vertebra with spring scissors and forceps. Expose the three adjacent vertebras. Use sterile gauze pads and sterile saline to control bleeding and&#160;clean the surgical site.</li>
	<li>Stabilize the spine with the help of the two stainless steel sidebars on a custom-designed stabilization stage (Figure 2B).</li>
	<li>Use a #2 forcep to perform a laminectomy at T12. Carefully use the forcep to break the lamina piece by piece until the whole lamina of the T12 vertebra is removed.</li>
	<li>Wash away the blood overlying the spinal cord with sterile saline and use the gauze pad to absorb excessive fluid. Apply pressure on the bleeding site with a piece of gauze pad to control bleeding.</li>
	<li>Place a coverslip (22 x 22 mm) on the clamping bar and fill the interspace between the coverslip and the spinal cord with saline.</li>
</ol>
4. In vivo TPEF-SRS imaging of mouse spinal cord
<ol>
	<li>Mount the stabilization stage on a five-axis stage beneath the TPEF-SRS microscope. 
	NOTE: The five-axis stage allows three-axis translation and &#177;5&#176; pitch and roll flexure motion.</li>
	<li>Secure the mouse head with two head bars and lower the holding plate to offer enough space for chest movement during breathing, alleviating motion artifacts.</li>
	<li>Insert a heating pad under the mouse to keep the mouse warm during imaging.</li>
	<li>Adjust the z translational stage to tune the focus until the bright-field image of spinal cord vasculature can be observed under a 10x objective.</li>
	<li>Locate the spinal cord dorsal vein at the center of the FOV by tuning the two-axis translational stage of the five-axis stage.</li>
	<li>Tune the roll and pitch angles of the five-axis stage to adjust the spinal cord dorsal surface perpendicular to the objective axis based on the bright-field image.</li>
	<li>Replace the 10x with a 25x water immersion objective for TPEF-SRS imaging.</li>
	<li>Set the wavelength of the fs beam to 920 nm. Tune the fs beam power to be 10 mW on the sample.</li>
	<li>Set the wavelength of the pump beam to 796 nm. Tune the power of the pump and Stokes beam to be 60 mW and 75 mW on the sample, respectively. 
	NOTE: Spinal cord SRS imaging is performed at the wavenumber of 2863.5 cm-1, which corresponds to the Raman peak of myelin sheaths at the carbon-hydrogen region based on the measured SRS spectra7,9. The laser power is determined to ensure high-resolution TPEF-SRS imaging of the spinal cord. Tissue damage is not observed under the current imaging conditions.</li>
	<li>For SRS imaging, select PBS above the objective using a motorized flipper by pressing the Switch button connected to the motorized flipper.</li>
	<li>Set the imaging parameters as follows: 512 x 512 pixels for 150 &#956;m x 150 &#956;m FOV, 3.2 &#956;s pixel dwell time, 2 &#956;s time constant.</li>
	<li>Start scanning the sample and set the focus on the dorsal surface of the spinal cord.</li>
	<li>Finely tune the delay stage by the OPO control software until maximum spinal cord SRS signal is achieved. 
	NOTE: Biological samples can induce extra chromatic dispersion, the delay stage may require to be adjusted to optimize the temporal synchronization. However, when SRS imaging is performed near the tissue top surface, the temporal difference of the two laser pulses introduced by the biological tissue and the imaging window is usually small (less than several hundred fs).</li>
	<li>For TPEF imaging, select dichroic mirror D2 above the objective using a motorized flipper by pressing the Switch button connected to the motorized flipper.</li>
	<li>Set the imaging parameters and start scanning the sample. To capture the TPEF-SRS image stack, acquire the TPEF and SRS images sequentially with a 1 s interval at the same depth before going to the next depth. The imaging parameters for TPEF imaging are 512 x 512 pixels, 150 &#956;m x 150 &#956;m FOV, 3.2 &#956;s pixel dwell time.</li>
	<li>Remove the animal from the stabilization stage after all images are collected.</li>
	<li>Clean the exposed tissue by flushing saline and absorb the excess fluid using a gauze pad. Apply silicone gel on the exposed spinal cord and wait for ~5 min until it gets cured.</li>
	<li>Suture the skin with #6-0 surgical suture to close the wound. Apply burn cream on the skin of the surgical site to prevent infection. Administer analgesic treatment subcutaneously (0.1 mg/kg buprenorphine).</li>
	<li>Place the animal in a clean cage and place the cage on a heating pad until the mouse fully recovers from anesthesia.</li>
	<li>Repeat analgesic administration every 12 hours for 3 days after surgery.</li>
	<li>Close the shutter of OPO and fs laser. Set the power of the pump and Stokes beam to the maximum. 
	NOTE: Setting maximal power before turning off the laser is beneficial for laser maintenance.</li>
	<li>Set the wavelength of the pump beam and fs laser to 800 nm.</li>
	<li>Set the two lasers on standby, close all the electronics control software and turn off all the associated equipment.</li>
</ol>

The authors have nothing to disclose and have no competing financial interests.

In this protocol, the basic setup of the TPEF-SRS microscope is described in detail. For SRS imaging, the pump and Stokes beams are temporally and spatially overlapped inside the OPO.&#160;However, this overlapping can be disrupted after passing through the microscope system. Therefore, both spatial and temporal optimization of the colocalization of the pump and Stokes beams is necessary and critical to achieving optimal SRS imaging. The temporal delay between the pump and Stokes beam is related to the optical path diffe...

Raman microscopy1,2 is emerging as a powerful label-free method to image biological tissues based on the characteristic frequencies of various chemical bonds in biomolecules. Owing to its non-invasive and well-adaptive imaging capability, Raman microscopy has been widely used for imaging lipid-enriched components in biological tissues like myelin sheath3,4,5, adipocytes6,7, and lipid droplets8,9,10. Stimulated Raman scattering (SRS) signal acquired as stimulated Raman gain (SRG) or stimulated Raman loss (SRL) is background-free, showing perfect spectral resemblance to spontaneous Raman scattering11,12. In addition, SRL and SRG are linearly dependent on the analyte concentration, allowing for quantitative analysis of biochemical components9,11,13. Two-photon excited fluorescence microscopy (TPEF) has been widely used for in vivo biological imaging owing to its inherent optical sectioning capability, deep penetration depth, and low phototoxicity14,15,16. However, the performance of TPEF imaging depends on the characteristics of fluorescent tags, and the number of resolvable colors is limited due to the broadband fluorescence spectra8,17,18,19. Label-free SRS imaging and fluorescence-based TPEF imaging are two complementary imaging modalities, and their combination can provide abundant biophysical and biochemical information of tissues. These two imaging modalities are both based on the nonlinear optical (NLO) processes, which allows simple integration in one microscope system. The combination of the SRS and TPEF imaging, the so-called dual-modal imaging, enables high-dimensional imaging and profiling of cells and tissues, facilitating a comprehensive understanding of complex biological systems. Specifically, picosecond (ps) SRS microscopy can achieve chemical-bond imaging with high spectral resolution compared with femtosecond (fs) SRS technique11, allowing to differentiate multiple biochemical components in biological tissue, especially in the crowded fingerprint region20,21. In addition, compared with another commonly used dual-modal NLO microscope system with integration of coherent anti-Stokes scattering (CARS) microscope, SRS shows superior performance to CARS in terms of spectral and image interpretation as well as detection sensitivity11. The SRS-TPEF microscope has been used as a powerful tool to study various biological systems, such as Caenorhabditis elegans9,22, Xenopus laevis tadpole brain5, mouse brain23,24, spinal cord25,26, peripheral nerve27, and adipose tissue7, etc.
The spinal cord together with the brain makes up the central nervous system (CNS). Visualizing cellular activities in the CNS in vivo under physiological and pathological conditions is critical for understanding the mechanisms of CNS disorders28,29,30 and for developing corresponding therapies31,32,33. Myelin sheath, which wraps and insulates axons for high-speed action potential conduction, plays a significant role in the development of the CNS. Demyelination is thought of as a hallmark in white matter disorders, such as multiple sclerosis34. In addition, after spinal cord injury35, myelin debris can modulate macrophage activation, contributing to chronic inflammation and secondary injury36. Therefore, in vivo imaging of myelin sheath together with neurons and glial cells in living mouse models is of great help to understand the dynamic processes in CNS disorders.
In this protocol, the fundamental setup procedures of a home-built TPEF-SRS microscope are described and the dual-modal in vivo imaging methods for mouse spinal cord are introduced.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>#2 Forceps</td>
			<td>Dumont</td>
			<td>11223-20</td>
			<td>For laminectomy</td>
		</tr>
		<tr>
			<td>10X objective</td>
			<td>Nikon</td>
			<td>CFI Plan Apo Lambda 10X</td>
			<td></td>
		</tr>
		<tr>
			<td>25X objective</td>
			<td>Olympus</td>
			<td>XLPLN25XSVMP2</td>
			<td></td>
		</tr>
		<tr>
			<td>Burn cream</td>
			<td>Betadine</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Camera</td>
			<td>Sony</td>
			<td>&#945;6300</td>
			<td></td>
		</tr>
		<tr>
			<td>Current amplifier</td>
			<td>Stanford research</td>
			<td>SR570</td>
			<td></td>
		</tr>
		<tr>
			<td>Current photomultiplier modules</td>
			<td>Hamamatsu</td>
			<td>H11461-01</td>
			<td></td>
		</tr>
		<tr>
			<td>D2 665 nm long-pass dichroic mirror</td>
			<td>Semrock</td>
			<td>FF665-Di02-25x36</td>
			<td>For directing epi-fluorescence signal to the detection module</td>
		</tr>
		<tr>
			<td>D3 700 nm short-pass dichroic mirror</td>
			<td>Edmund</td>
			<td>69-206</td>
			<td>For separating SRS from TPEF detection path</td>
		</tr>
		<tr>
			<td>Depilating cream</td>
			<td>Veet</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>FS1 975 nm short-pass filter</td>
			<td>Edmund</td>
			<td>86-108</td>
			<td>For blocking stokes beam</td>
		</tr>
		<tr>
			<td>FS1 Bandpass filter</td>
			<td>Semrock</td>
			<td>FF01-850/310</td>
			<td>For blocking stokes beam</td>
		</tr>
		<tr>
			<td>Fs2 Bandpass filter</td>
			<td>Semrock</td>
			<td>FF01-525/50</td>
			<td>For selecting YFP signal</td>
		</tr>
		<tr>
			<td>Fs2 Shortpass filter</td>
			<td>Semrock</td>
			<td>FF01-715/SP-25</td>
			<td>For blocking fs excitation laser beam</td>
		</tr>
		<tr>
			<td>Half-wave plate</td>
			<td>Thorlabs</td>
			<td>SAHWP05M-1700</td>
			<td></td>
		</tr>
		<tr>
			<td>High-speed photodetector</td>
			<td>MenloSystems</td>
			<td>FPD 310-F</td>
			<td>For checking Stokes beam modulation</td>
		</tr>
		<tr>
			<td>Iodine</td>
			<td>Betadine</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>IR Scope</td>
			<td>FJW</td>
			<td>FIND-R-SCOPE Infrared Viewer 2X Kit Model 84499C2X</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris</td>
			<td>Thorlabs</td>
			<td>CPA1</td>
			<td></td>
		</tr>
		<tr>
			<td>L1</td>
			<td>Thorlabs</td>
			<td>AC254-060-B-ML</td>
			<td></td>
		</tr>
		<tr>
			<td>L10</td>
			<td>Thorlabs</td>
			<td>LA4052-A</td>
			<td></td>
		</tr>
		<tr>
			<td>L2</td>
			<td>Thorlabs</td>
			<td>LA1422-B</td>
			<td></td>
		</tr>
		<tr>
			<td>L3</td>
			<td>Thorlabs</td>
			<td>AC254-050-B</td>
			<td></td>
		</tr>
		<tr>
			<td>L4</td>
			<td>Thorlabs</td>
			<td>AC254-060-B-ML</td>
			<td></td>
		</tr>
		<tr>
			<td>L7</td>
			<td></td>
			<td></td>
			<td>f=100 mm, AB coating</td>
		</tr>
		<tr>
			<td>L8</td>
			<td>Thorlabs</td>
			<td>LA4874-A</td>
			<td></td>
		</tr>
		<tr>
			<td>L9</td>
			<td>Thorlabs</td>
			<td>AC254-035-B-ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Lock-in amplifier</td>
			<td>APE</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mirror</td>
			<td>Thorlabs</td>
			<td>PF10-03-P01</td>
			<td></td>
		</tr>
		<tr>
			<td>Motorized flipper</td>
			<td>Thorlabs</td>
			<td>MFF101/M</td>
			<td></td>
		</tr>
		<tr>
			<td>multifunctional acquisition card</td>
			<td>National Instrument</td>
			<td>PCIe-6363</td>
			<td></td>
		</tr>
		<tr>
			<td>Oscilloscope</td>
			<td>Tektronix</td>
			<td>TDS2012C</td>
			<td></td>
		</tr>
		<tr>
			<td>Photodiode</td>
			<td>APE</td>
			<td></td>
			<td>For detecting SRS signal</td>
		</tr>
		<tr>
			<td>Picosecond laser source</td>
			<td>APE</td>
			<td>picoEmerald</td>
			<td></td>
		</tr>
		<tr>
			<td>Polarizing beam splitter</td>
			<td>Thorlabs</td>
			<td>CCM1-PBS252/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Power meter</td>
			<td>Newport</td>
			<td>843-R</td>
			<td></td>
		</tr>
		<tr>
			<td>Saline</td>
			<td>Braun</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scan lens L5</td>
			<td>Thorlabs</td>
			<td>SL50-CLS2</td>
			<td></td>
		</tr>
		<tr>
			<td>Scanning mirror</td>
			<td>Cambridge Technology</td>
			<td>6215H</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicone gel</td>
			<td>World Precision Inc.</td>
			<td>KWIK-SIL</td>
			<td></td>
		</tr>
		<tr>
			<td>Ti:sapphire fs laser</td>
			<td>Coherent</td>
			<td>Chameleon Ultra II</td>
			<td></td>
		</tr>
		<tr>
			<td>Tube lens L6</td>
			<td>Thorlabs</td>
			<td>TTL200-S8</td>
			<td></td>
		</tr>
	</tbody>
</table>

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 dual-modal imaging of spinal axons as well as myelin sheaths is conducted using the Thy1-YFPH transgenic mice, which express EYFP in dorsal root ganglion afferent neurons (Figure 3). These labeled afferent neurons relay the sensory information from the peripheral nerve to the spinal cord, with the central branch located in the spinal cord dorsal column. With the TPEF-SRS microscope, densely distributed myelin sheath can be clearly visualized using label-free SRS imaging, and...

Watch this Scientific Journal Video about In vivo Imaging of Biological Tissues with Combined Two-Photon Fluorescence and Stimulated Raman Scattering Microscopy at JoVE.com

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.

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 ...

in-vivo-imaging-biological-tissues-with-combined-two-photon

Research

JoVE Journal

Bioengineering

3.1K Views.  The Hong Kong University of Science and Technology. 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.

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

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.

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 ...

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Small and Wide Angle X-Ray Scattering Studies of Biological Macromolecules in Solution

In this paper, Small and Wide Angle X-ray Scattering (SWAXS) analysis of macromolecules is demonstrated through experimentation. SWAXS is a technique where X-rays are elastically scattered by an inhomogeneous sample in the nm-range at small angles (typically 0.1 - 5&deg;) and wide angles (typically &gt; 5&deg;). This technique provides information about the shape, size, and distribution of macromolecules, characteristic distances of partially ordered materials, pore sizes, and surface-to-volume ratio. Small Angle X-ray Scattering (SAXS) is capable of delivering structural information of macromolecules between 1 and 200 nm, whereas Wide Angle X-ray Scattering (WAXS) can resolve even smaller Bragg spacing of samples between 0.33 nm and 0.49 nm based on the specific system setup and detector. The spacing is determined from Bragg's law and is dependent on the wavelength and incident angle.
In a SWAXS experiment, the materials can be solid or liquid and may contain solid, liquid or gaseous domains (so-called particles) of the same or another material in any combination. SWAXS applications are very broad and include colloids of all types: metals, composites, cement, oil, polymers, plastics, proteins, foods, and pharmaceuticals. For solid samples, the thickness is limited to approximately 5 mm.
Usage of a lab-based SWAXS instrument is detailed in this paper. With the available software (e.g., GNOM-ATSAS 2.3 package by D. Svergun EMBL-Hamburg and EasySWAXS software) for the SWAXS system, an experiment can be conducted to determine certain parameters of interest for the given sample. One example of a biological macromolecule experiment is the analysis of 2 wt% lysozyme in a water-based aqueous buffer which can be chosen and prepared through numerous methods. The preparation of the sample follows the guidelines below in the Preparation of the Sample section. Through SWAXS experimentation, important structural parameters of lysozyme, e.g. the radius of gyration, can be analyzed.

The demonstration of the small and wide angle X-ray scattering (SWAXS) procedure has become instrumental in the study of biological macromolecules. Through the use of the instrumentation and procedures of specific angle methods and preparation, the experimental data from the SWAXS displays the atomic and nano-scale characterization of macromolecules.

In this paper, Small and Wide Angle X-ray Scattering (SWAXS) analysis of macromolecules is demonstrated through experimentation. SWAXS is a technique ...

Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as growth factors. Here, we outline a method to create opposing gradients of adhesive and soluble cues in a microfluidic chamber, which is compatible with live cell imaging. A copolymer of poly-L-lysine and polyethylene glycol (PLL-PEG) is employed to passivate glass coverslips and prevent non-specific adsorption of biomolecules and cells. Next, microcontact printing or dip pen lithography are used to create tracks of streptavidin on the passivated surfaces to serve as anchoring points for the biotinylated peptide arginine-glycine-aspartic acid (RGD) as the adhesive cue. A microfluidic device is placed onto the modified surface and used to create the gradient of adhesive cues (100% RGD to 0% RGD) on the streptavidin tracks. Finally, the same microfluidic device is used to create a gradient of a chemoattractant such as fetal bovine serum (FBS), as the soluble cue in the opposite direction of the gradient of adhesive cues.

A method for the assembly of adhesive and soluble gradients in a microscopy chamber for live cell migration studies is described. The engineered environment combines antifouling surfaces and adhesive tracks with solution gradients and therefore allows one to determine the relative importance of guidance cues.

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as ...

Imaging of Biological Tissues by Desorption Electrospray Ionization Mass Spectrometry

Mass spectrometry imaging (MSI) provides untargeted molecular information with the highest specificity and spatial resolution for investigating biological tissues at the hundreds to tens of microns scale. When performed under ambient conditions, sample pre-treatment becomes unnecessary, thus simplifying the protocol while maintaining the high quality of information obtained. Desorption electrospray ionization (DESI) is a spray-based ambient MSI technique that allows for the direct sampling of surfaces in the open air, even in vivo. When used with a software-controlled sample stage, the sample is rastered underneath the DESI ionization probe, and through the time domain, m/z information is correlated with the chemical species' spatial distribution. The fidelity of the DESI-MSI output depends on the source orientation and positioning with respect to the sample surface and mass spectrometer inlet. Herein, we review how to prepare tissue sections for DESI imaging and additional experimental conditions that directly affect image quality. Specifically, we describe the protocol for the imaging of rat brain tissue sections by DESI-MSI.

Desorption electrospray ionization mass spectrometry (DESI-MS) is an ambient method by which samples, including biological tissues, can be imaged with minimal sample preparation. By rastering the sample below the ionization probe, this spray-based technique provides sufficient spatial resolution to discern molecular features of interest within tissue sections.

Mass spectrometry imaging (MSI) provides untargeted molecular information with the highest specificity and spatial resolution for investigating biological ...

Long-term Intravital Immunofluorescence Imaging of Tissue Matrix Components with Epifluorescence and Two-photon Microscopy

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue function during development and organ homeostasis. It does so by acting via biochemical, biomechanical, and biophysical signaling pathways, such as through the release of bioactive ECM protein fragments, regulating tissue tension, and providing pathways for cell migration. The extracellular matrix of the tumor microenvironment undergoes substantial remodeling, characterized by the degradation, deposition and organization of fibrillar and non-fibrillar matrix proteins. Stromal stiffening of the tumor microenvironment can promote tumor growth and invasion, and cause remodeling of blood and lymphatic vessels. Live imaging of matrix proteins, however, to this point is limited to fibrillar collagens that can be detected by second harmonic generation using multi-photon microscopy, leaving the majority of matrix components largely invisible. Here we describe procedures for tumor inoculation in the thin dorsal ear skin, immunolabeling of extracellular matrix proteins and intravital imaging of the exposed tissue in live mice using epifluorescence and two-photon microscopy. Our intravital imaging method allows for the direct detection of both fibrillar and non-fibrillar matrix proteins in the context of a growing dermal tumor. We show examples of vessel remodeling caused by local matrix contraction. We also found that fibrillar matrix of the tumor detected with the second harmonic generation is spatially distinct from newly deposited matrix components such as tenascin C. We also showed long-term (12 hours) imaging of T-cell interaction with tumor cells and tumor cells migration along the collagen IV of basement membrane. Taken together, this method uniquely allows for the simultaneous detection of tumor cells, their physical microenvironment and the endogenous tissue immune response over time, which may provide important insights into the mechanisms underlying tumor progression and ultimate success or resistance to therapy.

The extracellular matrix undergoes substantial remodeling during wound healing, inflammation and tumorigenesis. We present a novel intravital immunofluorescence microscopy approach to visualize the dynamics of fibrillar as well as mesh-like matrix components with high spatial and temporal resolution using epifluorescence or two-photon microscopy.

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

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 ...

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to visualize these highly dynamic events, two complementary light microscopy techniques that allow fast imaging of live samples were combined: spinning disk microscopy (SD) for fast and high-resolution volume recording and total internal reflection fluorescence (TIRF) microscopy for precise localization and visualization of the plasma membrane. A comprehensive and complete imaging protocol will be shown for guiding through sample preparation, microscope calibration, image formation and acquisition, resulting in multi-color SD-TIRF live imaging series with high spatio-temporal resolution. All necessary image post-processing steps to generate multi-dimensional live imaging datasets, i.e. registration and combination of the individual channels, are provided in a self-written macro for the open source software ImageJ. The imaging of fluorescent proteins during initiation and maturation of adhesion complexes, as well as the formation of the actin cytoskeletal network, was used as a proof of principle for this novel approach. The combination of high resolution 3D microscopy and TIRF provided a detailed description of these complex processes within the cellular environment and, at the same time, precise localization of the membrane-associated molecules detected with a high signal-to-background ratio.

An advanced microscope that permit fast and high-resolution imaging of both, the isolated plasma membrane and the surrounding intracellular volume, will be presented. The integration of spinning disk and total internal reflection fluorescence microscopy in one setup allows live imaging experiments at high acquisition rates up to 3.5 s per image stack.

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to ...