We thank Ruth A. Singer and Richard K.P. Benninger for providing mouse pancreas tissues. W.M. acknowledges support from NIH R01 (GM128214), R01 (GM132860), R01 (EB029523) and US Army (W911NF-19-1-0214).

The protocol was conducted in accordance with the animal experimental protocol (AC-AABD1552) approved by the Institutional Animal Care and Use Committee at Columbia University.
1. Preparation of Raman-dye-conjugated antibodies
<ol>
	<li>Prepare the conjugation buffer as ~0.1 M NaHCO3 in PBS buffer, pH = 8.3, store at 4 &#176;C.</li>
	<li>Prepare N-hydroxysuccinimidy (NHS) ester-functionated MARS probe (Supplementary Material) solution as 3 mM in anhydrous DMSO. Synthesis of MARS probes can be referred to prior reports13,17,18. 
	NOTE: For storage purposes, the dye NHS ester solution must be protected from light and kept at -20 &#176;C.</li>
	<li>Dissolve antibody solids in the conjugation buffer to a concentration of 2 mg/mL. For antibodies that are dissolved in other buffers, exchange them into the conjugation buffer to a concentration of 1-2 mg/mL with centrifugal filters. 
	NOTE: Highly cross-adsorbed secondary antibodies are preferred for multiplex staining to minimize cross-species reactivity. The secondary antibodies used are listed in Table 1 and Table of Materials.</li>
	<li>Perform the conjugation reaction.
	<ol>
		<li>For secondary antibodies, add 15-fold molar excess of dye solution to the antibody solution in a glass vial slowly with stirring. For example, in 0.5 mL 2 mg/mL antibody solution, add 35 &#181;L 3 mM dye solution.</li>
		<li>Incubate the reaction mixture at room temperature (RT) with stirring for 1 h. Protect the reaction from light.</li>
	</ol>
	</li>
	<li>Purification.
	<ol>
		<li>Prepare the slurry of gel filtration resin (Table of Materials) in PBS buffer, following steps 1.5.2-1.5.4.</li>
		<li>Add 10 mL of gel filtration resin powder into 40 mL of PBS buffer inside a 50-mL tube.</li>
		<li>Keep the solution in a 90 &#176;C water bath for 1 h.</li>
		<li>Decant the supernatant and re-add PBS to 40 mL. Store the slurry at 4 &#176;C.</li>
		<li>Pack the size exclusion column (1-cm diameter, gravity-flow columns) with the slurry solution to the height of 10-15 cm.</li>
		<li>Rinse and wash the column with ~10 mL of PBS buffer to further pack the resin.</li>
		<li>Pipet the conjugation reaction mixture (~0.5 mL) to the column. Immediately add 1 mL of PBS buffer as the elution buffer when all reaction mixture is loaded. Constantly refill the elution buffer (PBS) to the column.</li>
		<li>Collect the eluate of the conjugate solution by looking at the color on the column (MARS dyes have light green to blue colors) or measuring the absorbance at 280 nm (A280).</li>
	</ol>
	</li>
	<li>Concentrate the collected solution to 1-2 mg/mL with a centrifugal filter.</li>
	<li>Determine the concentration and the average degree of labeling (DOL, dye-to-protein ratio) by measuring the ultraviolet-visible (UV-Vis) spectrum of the conjugate solution with a Nano plate reader. 
	&#8203;NOTE: Supplementary Material provides properties of MARS-dye NHS-esters for calculation. The normal DOL for the secondary antibody is around 3.</li>
</ol>
2. Tissue sample preparation
<ol>
	<li>Paraformaldehyde fixed mouse brain tissues.
	<ol>
		<li>Anesthetize the mice (C57BL/6J, Female, 25 d postnatal) with isoflurane. Assess the proper anesthetization with a toe pinch test.</li>
		<li>Kill the mice by cervical displacement. Perfuse the mice immediately with 4% paraformaldehyde (PFA) in PBS transcardially.</li>
		<li>Collect the mouse brain, following steps 2.1.4-2.1.5</li>
		<li>Cut upward from the brain stem along the sagittal suture. Peel the two halves of the skull away to the side and scoop out the brain with a tweezer.</li>
		<li>Fix the collected brain in 4% PFA in PBS at 4 &#176;C for 24 h. Then, wash the brain in PBS buffer at 4 &#176;C for 24 h to remove excess PFA.</li>
		<li>Place solid agarose into water to a final concentration of 7% (w/v) in a beaker, with a loose lid. Stir the solution with a glass stirring rod. Heat the slurry in microwave until the solution is clear.</li>
		<li>Allow the agarose to cool to 45-55 &#176;C.</li>
		<li>Pour the liquid agarose into a small chamber, then transfer the brain from PBS to liquid agarose and orient it with a spatula to embed the brain. Wait for the tissue-agarose block to harden.</li>
		<li>Section the tissue-agrose into 40-&#181;m-thick coronal slices using a vibratome.</li>
		<li>Transfer the tissue to a 4-well plate for the following staining. Remove the agarose with a tweezer. Wash the tissue with 1 mL of PBS, three times.</li>
	</ol>
	</li>
	<li>Fixed frozen mouse pancreas tissues.
	<ol>
		<li>Fix the mouse pancreata in 4% PFA in PBS at 4 &#176;C with rocking for 16-20 h.</li>
		<li>Wash the sample in 1 mL of PBS (4 &#176;C) three times to remove PFA.</li>
		<li>Embed the sample (~0.3-0.5 cm in size) in optimal cutting temperature (OCT) compound blocks. Put 2 drops of OCT into a plastic cryomold. Place the tissue in correct orientation and pour OCT on top of tissues until none of the tissue remains exposed.</li>
		<li>Section the pancreata to 8-&#181;m-thick slices and immobilize them onto a tissue binding glass slide, store them in -80 &#176;C.</li>
		<li>Before staining, equilibrate the specimen to RT. Wash the tissue with PBS to remove OCT blocks.</li>
	</ol>
	</li>
	<li>FFPE samples.
	<ol>
		<li>Bake the FFPE tissue slide at 60 &#176;C for 10 min.</li>
		<li>Deparaffinization and rehydration: Place samples sequentially in the following solutions in a 50-mL tube at RT for 3 min each time with mild shaking: 
		xylene two times, 
		ethanol two times, 
		95%&#160;(vol/vol) ethanol in deionized water two times, 
		70%&#160;(vol/vol) ethanol in deionized water two times, 
		50%&#160;(vol/vol) ethanol in deionized water one time, 
		deionized water one time.</li>
		<li>Transfer the sample into 20 mM sodium citrate (pH 8.0) at 100 &#176;C in a glass jar. Make sure the tissues are immersed in the solution.</li>
		<li>Put the jar in a 60 &#176;C water bath for 45 min.</li>
		<li>Wash the sample with deionized water at RT for 5 min.</li>
	</ol>
	</li>
</ol>
3. Tissue immuno-eprSRS staining
<ol>
	<li>Use a hydrophobic pen to draw a boundary around the tissue sections on the slide. 
	NOTE: A slide staining jar is used for following incubation steps of tissue on the slide. Floating tissues (40-&#181;m-thick mouse brain sections) are stained in well plates.</li>
	<li>Incubate the tissues with 0.3-0.5% PBST (Triton X-100 in PBS) for 10 min.</li>
	<li>Incubate the tissues with blocking buffer (5% donkey serum, 0.5% Triton X-100 in PBS) for 30 min.</li>
	<li>Prepare the primary staining solution: add all primary antibodies to 200-500 &#181;L of staining buffer (2% donkey serum, 0.5% Triton X-100 in PBS) at desired concentrations. Centrifuge the primary staining solution at 13,000 x g for 5 min. Only use the supernatant if precipitates form.</li>
	<li>Incubate the tissue in the primary antibody solution at 4 &#176;C for 1-2 days. 
	NOTE: For staining tissue sections on the slide, put the sample in a staining box with a wet wipe to maintain humidity.</li>
	<li>Wash the slides three times with 0.3-0.5% PBST at RT for 5 min each. Use 1 mL PBST for floating tissues. For tissues on the slide, wash the slides in a slide staining jar and make sure tissues are all immersed in the solution.</li>
	<li>Incubate the tissue in 200-500 &#181;L of blocking buffer for 30 min.</li>
	<li>Prepare the secondary staining solution: add all secondary antibodies (and lectins if required) to 200-500 &#181;L of staining buffer with desired concentrations (normally 10 &#181;g/mL). Centrifuge the secondary staining solution at 13,000 x g for 5 min. Only use the supernatant if precipitates form.</li>
	<li>Incubate the tissues in 200-500 &#181;L of secondary antibody solution at 4 &#176;C for 1-2 days.</li>
	<li>Wash the slides twice with 0.3-0.5% PBST at RT for 5 min each.</li>
	<li>Incubate with 200-500 &#181;L of DAPI solution for 30 min.</li>
	<li>Wash the slides three times with PBS at RT for 5 min each.</li>
	<li>For floating tissue sections, transfer them to glass slides with a glass-dropping pipette. Spread the tissue with a tissue brush and clean the surrounding with wipes if needed.</li>
	<li>Mount the tissue in a drop of antifade reagents with a glass coverslip and secure it with nail polish.</li>
</ol>
4. SRS microscope assembly
NOTE: A commercial confocal fluorescence system is used in tandem SRS-fluorescence imaging. More descriptions can be found in a prior report17. This protocol will focus on the SRS imaging side using narrowband excitation.
<ol>
	<li>Prepare a vibration-isolated optical table in a room with temperature control.</li>
	<li>Place a synchronized dual-laser system (pump and Stokes) on the optical table (Figure 2A) with a chiller connected. 
	NOTE: The fundamental laser in the dual-laser system provides an output pulse train at 1064 nm with 6-ps pulse width and 80-MHz repetition rate. The Stokes beam is from the fundamental laser. The intensity of the Stokes beam was modulated sinusoidally by a built-in electro-optic amplitude modulator (EOM) at 8 MHz with a modulation depth of more than 90%. The other portion of the fundamental laser is frequency-doubled to 532 nm, which is further used to synchronously seed a picosecond optical parametric oscillator (OPO) to produce a mode-locked pulse train with 5-6 ps pulse width (the idler beam of the OPO is blocked with an interferometric filter). The output wavelength of the OPO is tunable from 720-950 nm, which serves as the pump beam.</li>
	<li>Mount the mirrors (wavelength range: 750-1100 nm), dichroic beam splitters (DBS, 980 nm long-pass filter, rectangular), and lens (achromatic, AR coating for 650-1050 nm) to their respective mounts. Use very stablekinematic mirror mounts for the mirrors and dichroic beam splitters.</li>
	<li>Measure the height of the laser output and the beam sizes of the pump and the Stokes beams. Adjust the height of mirrors and lenses to ensure the light will hit the center of all the optical elements.</li>
	<li>Place mirror M1 on the optical table and make it ~45&#176; to the laser output (Figure 2B). Use the knobs on the kinematic mount to make tip and tilt adjustments. Ensure the light travels at the same height along the length of the table and a straight line with respect to the table.</li>
	<li>Place and align the dichroic beam splitters (980 nm long-pass filters) and mirrors to split the pump and Stokes beams (Figure 2A-B).</li>
	<li>Place and align lens pairs (L1, L2, and L3, L4) on each of the beam paths to collimate the beams and expand the beam diameters to match the back pupil of the objective (Figure 2A-B).</li>
	<li>Use M7 and M8 mirrors to align combined laser beams into the microscope (Figure 2C). Align one beam into the microscope first and use the mirror pairs on the other beam to ensure the spatial overlap of the two beams.</li>
	<li>Set up the detection part.
	<ol>
		<li>Put on an infrared-coated oil condenser (1.4-NA) to collect the forward-going pump and Stokes beams after passing through the specimens (Figure 2C).</li>
		<li>Mount a large-area Si photodiode onto a shielded box with BNC connectors (Figure 2E). Add a 64-V DC power supply to the mounted photodiode to increase its saturation threshold and response bandwidth.</li>
		<li>Reflect the forward-going light with a 2-inch mirror. Refocus the light onto the photodiode after an optical filter to block the modulating Stokes beam (Figure 2D).</li>
		<li>Send the output current of the photodiode to a fast lock-in amplifier terminated with 50 for signal demodulation. Send an 8-MHz trigger to the lock-in amplifier as the reference signal.</li>
		<li>Send the in-phase X-component of the lock-in amplifier into the analog interface box of the microscope.</li>
	</ol>
	</li>
	<li>Optimize the temporal overlap with the built-in motorized delay stage by measuring the SRS signal of pure D2O liquid at the microscope.</li>
</ol>
5. Image acquisition and analysis
<ol>
	<li>Perform multi-channel epr-SRS imaging with sequential pump wavelength tuning.
	<ol>
		<li>Set laser power to Ppump = 10-40 mW and PStokes = 40-80 mW on the laser control panel.</li>
		<li>Set pixel dwell time to 2-4 &#181;s and use multiple frames averaging typically 10-20 frames on the microscopy software. 
		NOTE: Avoid a combinatorial use of high laser power (Ppump&#62; 40 mW, PStokes&#62; 80 mW) and small pixel size (&#60;0.2 &#181;m), which likely cause &#39;bleaching effect&#39; of Raman dyes due to multiphoton excitation.</li>
		<li>Set the time constants of the lock-in amplifier to half of the pixel dwell time.</li>
	</ol>
	</li>
	<li>Linear spectral unmixing. 
	NOTE: epr-SRS follows a strict linear signal-to-concentration dependence over the entire concentration range; thus, linear spectral unmixing is effective to remove any potential cross-talks between channels. For N-channel epr-SRS measurement with N MARS probes, measured signals (S) can be expressed as S = MC, where C is the MARS probe concentrations, and M is an N x N matrix determined by Raman cross-sections of MARS probes.
	<ol>
		<li>Measure matrix M on single-color immuno-eprSRS samples labeled with different MARS probes.</li>
		<li>Use equation C = M&#8722;1&#183;S to determine the concentration matrix of the MARS probe with multiplex sample signal measurement S.</li>
	</ol>
	</li>
</ol>

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Coherent Raman Scattering Microscope Available from: <a target="_blank" href="https://www.leica-microsystems.com/products/confocal-microscopes/p/leica-tcs-sp8-cars/">https://www.leica-microsystems.com/products/confocal-microscopes/p/leica-tcs-sp8-cars/</a> (2022)</li></ol>

The authors have nothing to disclose.

Here, we present the immuno-eprSRS protocol which is broadly applicable to common tissue types, including freshly-preserved mouse tissues, FFPE human tissues, and frozen mouse tissues. Immuno-eprSRS has been validated for a panel of epitopes in cells and tissues, as listed in Table 1. This one-shot platform is particularly suitable for applications where cyclic strategies do not function well. For example, cyclic fluorescence is demanding for thick tissues as multiple rounds of 3D immunolabeling are unpr...

Complex tissue systems are composed of distinct cellular subpopulations whose spatial locations and interaction networks are deeply intertwined with their functions and dysfunctions1,2. To reveal the tissue architecture and interrogate its complexity, knowledge of the spatial locations of proteins at single-cell resolution is essential. Hence, highly multiplexed protein-imaging technologies have been increasingly appreciated and could become a cornerstone for studying tissue biology3,4,5. Current common multiplexed protein imaging methods can be classified into two main categories. One is serial immunofluorescence imaging relying on multiple rounds of tissue staining and imaging, and the other is imaging mass cytometry coupled with heavy metal tagged antibodies6,7,8,9,10,11,12.
Here, an alternative strategy for multiplexed antibody-based protein imaging is introduced. Unlike the prevalent fluorescence imaging modality, which can only visualize 4-5 channels simultaneously due to the broad excitation and emission spectra (full width at half maximum (FWHM) ~500 cm-1), Raman microscopy exhibits much narrower spectral linewidth (FWHM ~10 cm-1) and hence provides scalable multiplexity. Recently, by harnessing the narrow spectrum, a novel scheme of Raman microscopy named electronic pre-resonance stimulated Raman scattering (epr-SRS) microscopy has been developed, providing a powerful strategy for multiplexed imaging13. By probing the electronically coupled vibrational modes of Raman dyes, epr-SRS achieves a drastic enhancement effect of 1013-fold on Raman cross-sections and overcomes the sensitivity bottleneck of conventional Raman microscopy (Figure 1A)13,14,15. As a result, the detection limit of epr-SRS has been pushed to sub-&#181;M, which enables Raman detection of interesting molecular markers such as specific proteins and organelles inside cells13,16. In particular, utilizing Raman dye-conjugated antibodies, epr-SRS imaging of specific proteins in cells and tissues (called immuno-eprSRS) was demonstrated with comparable sensitivity to standard immunofluorescence (Figure 1B)13,17. By tuning the pump wavelength by only 2 nm, the epr-SRS signal will be completely off (Figure 1B), which showcases high vibrational contrast.
On the probe side, a set of rainbow-like Raman probes called Manhattan Raman scattering (MARS) dyes has been developed for antibody conjugation13,18,19,20. This unique Raman palette consists of novel dyes bearing &#960;-conjugated triple bonds (Supplementary Material), each displaying a single and narrow epr-SRS peak in the bioorthogonal Raman spectral range (Figure 1C). By modifying the structure of the core chromophore and isotopically editing both atoms of the triple bond (Supplementary Material), spectrally separated Raman probes have been developed. Leveraging the scalable multiplexity, epr-SRS microscopy coupled with the MARS dye palette offers an optical strategy for one-shot multiplex protein imaging in cells and tissues.
Immuno-eprSRS provides an alternative strategy to current multiplex protein imaging methods with unique strengths. Compared to fluorescence approaches with cyclic staining, imaging, and signal removal, this Raman-based platform ensures single-round staining and imaging. Therefore, it circumvents practical complexity in cyclic procedures and largely simplifies the protocol, hence opening new territories of multiplexed protein imaging. For instance, harnessing a Raman-dye-tailored tissue clearing protocol, immuno-eprSRS has been extended to three dimensions for highly multiplexed protein mapping in thick intact tissues17. Over 10 protein targets were visualized along millimeter-thick mouse brain tissues17. More recently, coupling immuno-eprSRS with an optimized biomolecule-retention expansion microscopy (ExM) protocol21, one-shot nanoscale imaging of multiple targets has also been demonstrated22. Compared to imaging mass spectroscopy4,9, epr-SRS is nondestructive and has intrinsically optical sectioning ability. Furthermore, epr-SRS is more time-efficient on tissue scanning. Typically, a tissue region of 0.25 mm2 with a pixel size of 0.5 &#181;m takes merely a few minutes to image for a single epr-SRS channel. For example, the total imaging time of four SRS channels plus four fluorescence channels in Figure 4 is about 10 min.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>16% Paraformaldehyde, EM Grade</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710</td>
			<td></td>
		</tr>
		<tr>
			<td>&#945;-tubulin</td>
			<td>Abcam</td>
			<td>ab18251</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>&#945;-tubulin</td>
			<td>BioLegend</td>
			<td>625902</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>&#946;-III-tubulin</td>
			<td>BioLegend</td>
			<td>657402</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>&#946;-III-tubulin</td>
			<td>Abcam</td>
			<td>ab41489</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>&#946;-tubulin</td>
			<td>Abcam</td>
			<td>ab131205</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Agarose, low gellling temperature</td>
			<td>Sigma Aldrich</td>
			<td>A9414</td>
			<td>For brain embedding</td>
		</tr>
		<tr>
			<td>Anti-a-tubulin antibody produced in rabbit (&#945;-tubulin)</td>
			<td>Abcam</td>
			<td>ab52866</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-Calbindin antibody produced in mouse (Calbindin)</td>
			<td>Abcam</td>
			<td>ab82812</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-GABA B receptor R2&#160; antibody produced in guinea pig (GABA B receptor R2)</td>
			<td>Millipore Sigma</td>
			<td>AB2255</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-GFAP antibody produced in goat (GFAP)</td>
			<td>Thermo Scientific</td>
			<td>PA5-18598</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-Glucagon&#160; antibody produced in mouse (Glucagon)</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-514592</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-insulin antibody produced in guinea pig (insulin)</td>
			<td>DAKO</td>
			<td>IR00261-2</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-MBP antibody produced in rat (MBP)</td>
			<td>Abcam</td>
			<td>ab7349</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-NeuN antibody produced in rabbit (NeuN)</td>
			<td>Thermo Scientific</td>
			<td>PA5-78639</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-Pancreatic polypeptide (PP) antibody produced in goat- Pancreatic polypeptide (PP)</td>
			<td>Sigma Aldrich</td>
			<td>SAB2500747</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-Pdx1 antibody produced in rabbit (Pdx1)</td>
			<td>Milipore</td>
			<td>06-1379</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-Somatostatin antibody produced in rat (Somatostatin)</td>
			<td>Abcam</td>
			<td>ab30788</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Anti-Vimentin antibody produced in chicken (Vimentin)</td>
			<td>Abcam</td>
			<td>ab24525</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Band-pass filter</td>
			<td>KR Electronics</td>
			<td>KR2724</td>
			<td>8 MHz</td>
		</tr>
		<tr>
			<td>BNC 50 Ohm Terminator</td>
			<td>Mini Circuits</td>
			<td>STRM-50</td>
			<td></td>
		</tr>
		<tr>
			<td>BNC cable</td>
			<td>Thorlabs</td>
			<td>2249-C</td>
			<td>Coaxial Cable, BNC Male / Male</td>
		</tr>
		<tr>
			<td>Broadband dielectric mirror</td>
			<td>Thorlabs</td>
			<td>BB1-E03</td>
			<td>750 - 1100 nm</td>
		</tr>
		<tr>
			<td>C57BL/6J mice</td>
			<td>Jackson Laboratory</td>
			<td>000664</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Condenser</td>
			<td>Olympus</td>
			<td></td>
			<td>oil immersion, 1.4 N.A.</td>
		</tr>
		<tr>
			<td>Cytokeratin 18</td>
			<td>Abcam</td>
			<td>ab7797</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Cytokeratin 18</td>
			<td>Abcam</td>
			<td>ab24561</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>DC power supply</td>
			<td>TopWard</td>
			<td>6302D</td>
			<td>Bias voltage is 64 V</td>
		</tr>
		<tr>
			<td>Dichroic mount</td>
			<td>Thorlabs</td>
			<td>KM100CL</td>
			<td>Kinematic Mount for up to 1.3&#34; (33 mm) Tall Rectangular Optics, Left Handed</td>
		</tr>
		<tr>
			<td>Donkey anti-Chicken IgY (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>703-005-155</td>
			<td>Secondary antibodies for MARS conjugation</td>
		</tr>
		<tr>
			<td>Donkey anti-Goat IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>705-005-147</td>
			<td>Secondary antibodies for MARS conjugation</td>
		</tr>
		<tr>
			<td>Donkey anti-Guinea Pig IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>706-005-148</td>
			<td>Secondary antibodies for MARS conjugation</td>
		</tr>
		<tr>
			<td>Donkey anti-Mouse IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>715-005-151</td>
			<td>Secondary antibodies for MARS conjugation</td>
		</tr>
		<tr>
			<td>Donkey anti-Rabbit IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>711-005-152</td>
			<td>Secondary antibodies for MARS conjugation</td>
		</tr>
		<tr>
			<td>Donkey anti-Rat IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>712-005-153</td>
			<td>Secondary antibodies for MARS conjugation</td>
		</tr>
		<tr>
			<td>Donkey anti-Sheep IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>713-005-147</td>
			<td>Secondary antibodies for MARS conjugation</td>
		</tr>
		<tr>
			<td>DPBS</td>
			<td>Fisher Scientific</td>
			<td>14-190-250</td>
			<td></td>
		</tr>
		<tr>
			<td>EpCAM</td>
			<td>Abcam</td>
			<td>ab71916</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sigma Aldrich</td>
			<td>443611</td>
			<td></td>
		</tr>
		<tr>
			<td>Fast-speed look-in amplifier</td>
			<td>Zurich Instruments</td>
			<td>HF2LI</td>
			<td>DC - 50 MHz</td>
		</tr>
		<tr>
			<td>FFPE Kidney Sample</td>
			<td>USBiomax</td>
			<td>HuFPT072</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibrillarin</td>
			<td>Abcam</td>
			<td>ab5821</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Giantin</td>
			<td>Abcam</td>
			<td>ab24586</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Glucagon</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-514592</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>H2B</td>
			<td>Abcam</td>
			<td>ab1790</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>HeLa</td>
			<td>ATCC</td>
			<td>ATCC CCL-2</td>
			<td></td>
		</tr>
		<tr>
			<td>High O.D. bandpass filter</td>
			<td>Chroma Technology</td>
			<td>ET890/220m</td>
			<td>Filter the Stokes beam and transmit the pump beam</td>
		</tr>
		<tr>
			<td>Hydrophobic pen</td>
			<td>Fisher Scientific</td>
			<td>NC1384846</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>ThermoFisher</td>
			<td>701265</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Integrated SRS laser system</td>
			<td>Applied Physics &#38; Electronics, Inc.</td>
			<td>picoEMERALD</td>
			<td>picoEMERALD provides an output pulse train at 1,064 nm with 6-ps pulse width and 80-MHz repetition rate, which serves as the Stokes beam. The frequency doubled beam at 532 nm is used to synchronously seed a picosecond optical parametric oscillator (OPO) to produce a mode-locked pulse train with five~6 ps pulse width (the idler beam of the OPO is blocked with an interferometric filter). The output wavelength of the OPO is tunable from 720&#8211;950 nm, which serves as the pump beam. The intensity of the 1,064-nm Stokes beam is modulated sinusoidally by a built-in EOM at 8 MHz with a modulation depth of more than 90%. The pump beam is spatially overlapped with the Stokes beam by using a dichroic mirror inside picoEMERALD. The temporal overlap between pump and Stokes pulse trains is achieved with a built-in delay stage and optimized by the SRS signal of pure D2O at the microscope.</td>
		</tr>
		<tr>
			<td>Inverted laser-scanning microscope</td>
			<td>Olympus</td>
			<td>FV1200MPE</td>
			<td></td>
		</tr>
		<tr>
			<td>Kinematic mirror mount</td>
			<td>Thorlabs</td>
			<td>POLARIS-K1-2AH</td>
			<td>2 Low-Profile Hex Adjusters</td>
		</tr>
		<tr>
			<td>Lectin from Triticum vulgaris (wheat)</td>
			<td>Sigma Aldrich</td>
			<td>L0636-5 mg</td>
			<td></td>
		</tr>
		<tr>
			<td>Long-pass dichroic beam splitter</td>
			<td>Semrock</td>
			<td>Di02-R980-25x36</td>
			<td>980 nm laser BrightLine&#160;single-edge laser-flat dichroic beamsplitter</td>
		</tr>
		<tr>
			<td>MAP2</td>
			<td>BioLegend</td>
			<td>801810</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Microscopy imaging software</td>
			<td>Olympus</td>
			<td>FluoView</td>
			<td></td>
		</tr>
		<tr>
			<td>NanoQuant Plate</td>
			<td>Tecan</td>
			<td></td>
			<td>For absorbance-based, small volume analyses in a plate reader.</td>
		</tr>
		<tr>
			<td>Normal donkey serum</td>
			<td>Jackson ImmunoResearch</td>
			<td>017-000-121</td>
			<td></td>
		</tr>
		<tr>
			<td>NucBlue Fixed Cell ReadyProbes Reagent (DAPI)</td>
			<td>Thermo Scientific</td>
			<td>R37606</td>
			<td></td>
		</tr>
		<tr>
			<td>Nunc 4-Well Dishes</td>
			<td>Fisher Scientific</td>
			<td>12-566-300</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective lens</td>
			<td>Olympus</td>
			<td>XLPlan N</td>
			<td>x25, 1.05-NA, MP, working distance&#8201;=&#8201;2&#8201;mm</td>
		</tr>
		<tr>
			<td>Paint brush</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Periscope assembly</td>
			<td>Thorlabs</td>
			<td>RS99</td>
			<td>includes the top and bottom units, &#216;1&#34; post, and clamping fork.</td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Plate reader</td>
			<td>Tecan</td>
			<td>Infinite 200 PRO</td>
			<td>An easy-to-use multimode plate reader. Absorbance measurement capabilities over a spectral range of 230&#8211;1000 nm.</td>
		</tr>
		<tr>
			<td>ProLong Gold antifade reagent</td>
			<td>Thermo Scientific</td>
			<td>P36930</td>
			<td></td>
		</tr>
		<tr>
			<td>PSD95</td>
			<td>Invitrogen</td>
			<td>51-6900</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Sephadex G-25 Medium</td>
			<td>GE Life Sciences</td>
			<td>17-0033-01</td>
			<td>gel filtration resin for desalting and buffer exchange</td>
		</tr>
		<tr>
			<td>Shielded box with BNC connectors</td>
			<td>Pomona Electronics</td>
			<td>2902</td>
			<td>Aluminum Box With Cover, BNC Female/Female</td>
		</tr>
		<tr>
			<td>Si photodiode</td>
			<td>Thorlabs</td>
			<td>FDS1010</td>
			<td>350&#8211;1100 nm, 10 mm x 10 mm Active Area</td>
		</tr>
		<tr>
			<td>Synapsin 2</td>
			<td>ThermoFisher</td>
			<td>OSS00073G</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Tissue Path Superfrost Plus Gold Slides</td>
			<td>Fisher Scientific</td>
			<td>22-035813</td>
			<td>Adhesive slide to attract and chemically bond fresh or formalin-fixed tissue sections firmly to the slide surface (tiisue bindling glass slides)</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher Scientific</td>
			<td>BP151-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica</td>
			<td>VT1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Vimentin</td>
			<td>Abcam</td>
			<td>ab8069</td>
			<td>Primary antibodies</td>
		</tr>
		<tr>
			<td>Xylenes</td>
			<td>Sigma Aldrich</td>
			<td>214736</td>
			<td></td>
		</tr>
	</tbody>
</table>

highly-multiplexed tissue imaging with raman dyes

Visualizing a vast scope of specific biomarkers in tissues plays a vital role in exploring the intricate organizations of complex biological systems. Hence, highly multiplexed imaging technologies have been increasingly appreciated. Here, we describe an emerging platform of highly-multiplexed vibrational imaging of specific proteins with comparable sensitivity to standard immunofluorescence via electronic pre-resonance stimulated Raman scattering (epr-SRS) imaging of rainbow-like Raman dyes. This method circumvents the limit of spectrally-resolvable channels in conventional immunofluorescence and provides a one-shot optical approach to interrogate multiple markers in tissues with subcellular resolution. It is generally compatible with standard tissue preparations, including paraformaldehyde-fixed tissues, frozen tissues, and formalin-fixed paraffin-embedded (FFPE) human tissues. We envisage this platform will provide a more comprehensive picture of protein interactions of biological specimens, particularly for thick intact tissues. This protocol provides the workflow from antibody preparation to tissue sample staining, to SRS microscope assembly, to epr-SRS tissue imaging.

Figure 3&#160;shows example images of epr-SRS in different samples, including fixed cells (Figure 3A), paraformaldehyde (PFA)-fixed mouse tissues (Figure 3B), and formalin-fixed paraffin-embedded (FFPE) human specimens (Figure 3C). The spatial resolution of SRS microscopy is diffraction-limited, the typical lateral resolution is ~300 nm, and the axial resolution is 1-2 &#181;m using near-infrared light ...

Watch this Scientific Journal Video about Highly-Multiplexed Tissue Imaging with Raman Dyes at JoVE.com

Highly-Multiplexed Tissue Imaging with Raman Dyes

Electronic pre-resonance stimulated Raman scattering (epr-SRS) imaging of rainbow-like Raman dyes is a new platform for highly multiplexed epitope-based protein imaging. Here, we present a practical guide including antibody preparation, tissue sample staining, SRS microscope assembly, and epr-SRS tissue imaging.

Visualizing a vast scope of specific biomarkers in tissues plays a vital role in exploring the intricate organizations of complex biological systems. ...

highly-multiplexed-tissue-imaging-with-raman-dyes

Research

JoVE Journal

Bioengineering

2.7K Views.  Columbia University. Electronic pre-resonance stimulated Raman scattering (epr-SRS) imaging of rainbow-like Raman dyes is a new platform for highly multiplexed epitope-based protein imaging. Here, we present a practical guide including antibody preparation, tissue sample staining, SRS microscope assembly, and epr-SRS tissue imaging.

Video: Highly-Multiplexed Tissue Imaging with Raman Dyes

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 ...

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, ...

Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of tri-leaflet polymer valve prostheses. However, identification of a protocol for initial assessment of polymer valve hydrodynamic functionality is paramount during the early stages of the design process. Traditional in vitro pulse duplicator systems are not configured to accommodate flexible tri-leaflet materials; in addition, assessment of polymer valve functionality needs to be made in a relative context to native and prosthetic heart valves under identical test conditions so that variability in measurements from different instruments can be avoided. Accordingly, we conducted hydrodynamic assessment of i) native (n = 4, mean diameter, D = 20 mm), ii) bi-leaflet mechanical (n= 2, D = 23 mm) and iii) polymer valves (n = 5, D = 22 mm) via the use of a commercially available pulse duplicator system (ViVitro Labs Inc, Victoria, BC) that was modified to accommodate tri-leaflet valve geometries. Tri-leaflet silicone valves developed at the University of Florida comprised the polymer valve group. A mixture in the ratio of 35:65 glycerin to water was used to mimic blood physical properties. Instantaneous flow rate was measured at the interface of the left ventricle and aortic units while pressure was recorded at the ventricular and aortic positions. Bi-leaflet and native valve data from the literature was used to validate flow and pressure readings. The following hydrodynamic metrics were reported: forward flow pressure drop, aortic root mean square forward flow rate, aortic closing, leakage&#160;and regurgitant volume, transaortic closing, leakage, and total energy losses. Representative results indicated that hydrodynamic metrics from the three valve groups could be successfully obtained by incorporating a custom-built assembly into a commercially available pulse duplicator system and subsequently, objectively compared to provide insights on functional aspects of polymer valve design.

There has been renewed interest in developing polymer valves. Here, the objectives are to demonstrate the feasibility of modifying a commercial pulse duplicator to accommodate tri-leaflet geometries and to define a protocol to present polymer valve hydrodynamic data in comparison to native and prosthetic valve data collected under near-identical conditions.

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of ...

Sensing of Barrier Tissue Disruption with an Organic Electrochemical Transistor

The gastrointestinal tract is an example of barrier tissue that provides a physical barrier against entry of pathogens and toxins, while allowing the passage of necessary ions and molecules. A breach in this barrier can be caused by a reduction in the extracellular calcium concentration. This reduction in calcium concentration causes a conformational change in proteins involved in the sealing of the barrier, leading to an increase of the paracellular flux. To mimic this effect the calcium chelator ethylene glycol-bis(beta-aminoethyl ether)-N,N,N&#39;,N&#39;-tetra acetic acid (EGTA) was used on a monolayer of cells known to be representative of the gastrointestinal tract. Different methods to detect the disruption of the barrier tissue already exist, such as immunofluorescence and&#160;permeability assays. However, these methods are time-consuming and costly and not suited to dynamic or high-throughput measurements. Electronic methods for measuring barrier tissue integrity also exist for measurement of the transepithelial resistance (TER), however these are often costly and complex. The development of rapid, cheap, and sensitive methods is urgently needed as the integrity of barrier tissue is a key parameter in drug discovery and pathogen/toxin diagnostics. The organic electrochemical transistor (OECT) integrated with barrier tissue forming cells has been shown as a new device capable of dynamically monitoring barrier tissue integrity. The device is able to measure minute variations in ionic flux with unprecedented temporal resolution and sensitivity, in real time, as an indicator of barrier tissue integrity. This new method is based on a simple device that can be compatible with high throughput screening applications and fabricated at low cost.

The Organic Electrochemical Transistor is integrated with live cells and used to monitor ion flux across the gastrointestinal epithelial barrier. In this study, an increase in ion flux, related to disruption of tight junctions, induced by the presence of the calcium chelator EGTA (ethylene glycol-bis(beta-aminoethyl ether)-N,N,N',N'-tetra acetic acid), is measured.

The gastrointestinal tract is an example of barrier tissue that provides a physical barrier against entry of pathogens and toxins, while allowing the ...

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 ...

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 ...

Conducting Multiple Imaging Modes with One Fluorescence Microscope

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to conventional epi-fluorescence microscopy, various imaging techniques have been developed to achieve specific experimental goals. Some of the widely used techniques include single-molecule fluorescence resonance energy transfer (smFRET), which can report conformational changes and molecular interactions with angstrom resolution, and single-molecule detection-based super-resolution (SR) imaging, which can enhance the spatial resolution approximately ten&#160;to twentyfold compared to diffraction-limited microscopy. Here we present a customer-designed integrated system, which merges multiple imaging methods in one microscope, including conventional epi-fluorescent imaging, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging. Different imaging methods can be achieved easily and reproducibly by switching optical elements. This set-up is easy to adopt by any research laboratory in biological sciences with a need for routine and diverse imaging experiments at a reduced cost and space relative to building separate microscopes for individual purposes.

Here we present a practical guide of building an integrated microscopy system, which merges conventional epi-fluorescent imaging, single-molecule detection-based super-resolution imaging, and multi-color single-molecule detection, including single-molecule fluorescence resonance energy transfer imaging, into one set-up in a cost-efficient way.

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to ...

Establishing Single-Cell Based Co-Cultures in a Deterministic Manner with a Microfluidic Chip

Cell co-culture assays have been widely used for studying cell-cell interactions between different cell types to better understand the biology of diseases including cancer. However, it is challenging to clarify the complex mechanism of intercellular interactions in highly heterogeneous cell populations using conventional co-culture systems because the heterogeneity of the cell subpopulation is obscured by the average values; the conventional co-culture systems can only be used to describe the population signal, but are incapable of tracking individual cells behavior. Furthermore, conventional single-cell experimental methods have low efficiency in cell manipulation because of the Poisson distribution. Microfabricated devices are an emerging technology for single-cell studies because they can accurately manipulate single cells at high-throughput and can reduce sample and reagent consumption. Here, we describe the concept and application of a microfluidic chip for multiple single-cell co-cultures. The chip can efficiently capture multiple types of single cells in a culture chamber (~46%) and has a sufficient culture space useful to study the cells&#39; behavior (e.g., migration, proliferation, etc.) under cell-cell interaction at the single-cell level. Lymphatic endothelial cells and oral squamous cell carcinoma were used to perform a single-cell co-culture experiment on the microfluidic platform for live multiple single-cell interaction studies.

This report describes a microfluidic chip-based method to set up a single cell culture experiment in which high-efficiency pairing and microscopic analysis of multiple single cells can be achieved.

Cell co-culture assays have been widely used for studying cell-cell interactions between different cell types to better understand the biology of diseases ...

Blood Flow Imaging with Ultrafast Doppler

The pulsed-Doppler effect is the main technique used in clinical echography to assess blood flow. Applied with conventional focused ultrasound Doppler modes, it has several limits. Firstly, a finely tuned signal filtering operation is needed to distinguish blood flows from surrounding moving tissues. Secondly, the operator must choose between localizing the blood flows or quantifying them. In the last two decades, ultrasound imaging has undergone a paradigm shift with the emergence of ultrafast ultrasound using unfocused waves. In addition to a hundredfold increase in framerate (up to 10000 Hz), this new technique also breaks the conventional quantification/localization trade-off, offering a complete blood flow mapping of the field of view and a simultaneous access to fine velocities measurements at the single-pixel level (down to 50 &#181;m). This data continuity in both spatial and temporal dimensions strongly improves the tissue/blood filtering process, which results in an increase sensitivity to small blood flow velocities (down to 1 mm/s). In this method paper, we aim to introduce the concept of ultrafast Doppler as well as its main parameters. Firstly, we summarize the physical principles of unfocused wave imaging. Then, we present the Doppler signal processing main steps. Particularly, we explain the practical implementation of the critical tissue/blood flow separation algorithms and on the extraction of velocities from these filtered data. This theoretical description is supplemented by in vitro experiences. A tissue phantom embedding a canal with flowing blood-mimicking fluid is imaged with a research programmable ultrasound system. A blood flow image is obtained and the flow characteristics are displayed for several pixels in the canal. Finally, a review of in vivo applications is proposed, showing examples in several organs such as carotids, kidney, thyroid, brain and heart.

This protocol shows how to apply ultrafast ultrasound Doppler imaging to quantify blood flows. After a 1 s long acquisition, the experimenter has access to a movie of the full field of view with axial velocity values for each pixel every &#8776;0.3 ms (depending on the ultrasound time of flight).

The pulsed-Doppler effect is the main technique used in clinical echography to assess blood flow. Applied with conventional focused ultrasound Doppler ...

Quantifying Fibrillar Collagen Organization with Curvelet Transform-Based Tools

Fibrillar collagens are prominent extracellular matrix (ECM) components, and their topology changes have been shown to be associated with the progression of a wide range of diseases including breast, ovarian, kidney, and pancreatic cancers. Freely available fiber quantification software tools are mainly focused on the calculation of fiber alignment or orientation, and they are subject to limitations such as the requirement of manual steps, inaccuracy in detection of the fiber edge in noisy background, or lack of localized feature characterization. The collagen fiber quantitation tool described in this protocol is characterized by using an optimal multiscale image representation enabled by curvelet transform (CT). This algorithmic approach allows for the removal of noise from fibrillar collagen images and the enhancement of fiber edges to provide location and orientation information directly from a fiber, rather than using the indirect pixel-wise or window-wise information obtained from other tools. This CT-based framework contains two separate, but linked, packages named &#8220;CT-FIRE&#8221; and &#8220;CurveAlign&#8221; that can quantify fiber organization on a global, region of interest (ROI), or individual fiber basis. This quantification framework has been developed for more than ten years and has now evolved into a comprehensive and user-driven collagen quantification platform. Using this platform, one can measure up to about thirty fiber features including individual fiber properties such as length, angle, width, and straightness, as well as bulk measurements such as density and alignment. Additionally, the user can measure fiber angle relative to manually or automatically segmented boundaries. This platform also provides several additional modules including ones for ROI analysis, automatic boundary creation, and post-processing. Using this platform does not require prior experience of programming or image processing, and it can handle large datasets including hundreds or thousands of images, enabling efficient quantification of collagen fiber organization for biological or biomedical applications.

Here, we present a protocol to use a curvelet transform-based, open-source MATLAB software tool for quantifying fibrillar collagen organization in the extracellular matrix of both normal and diseased tissues. This tool can be applied to images with collagen fibers or other types of line-like structures.

Fibrillar collagens are prominent extracellular matrix (ECM) components, and their topology changes have been shown to be associated with the progression ...