The authors thank Sanaz Vahidinia for introducing us to Dr. Jorgensen&#39;s colleagues and his work. This research was supported by grants from the Army Research Office (no. W911NF-10-0139), the Office of Naval Research (through MURI award no. N00014-09-1-1053), and NSF-INSPIRE award NSF-1343158.

This study is compliant with all the relevant ethical regulations of Yale University regarding vertebrate and invertebrate animal research.
1. Building the optical micro-probe
<ol>
	<li>Building the glass sleeve, material: Pasteur pipette, 5.75 in(see Table of Materials)
	<ol>
		<li>Using a mountable alligator clip (Table of Materials), mount the glass pipette by the wide end such that the tapered end is facing down toward the floor and the orientation of the pipette is perpendicular to the floor.</li>
		<li>Hang a 50 g plastic vice grip (Table of Materials) from the tapered end of the pipette. Place a pad of electrical tape between the pipette and the vice grip to help prevent slippage.</li>
		<li>Heat the tapered end of the pipette using a small butane torch (Table of Materials). Apply the flame 1 cm above the vice grip. Hold the torch such that the brightest part of the flame is 3 cm from the glass and the glass is at the tip of the flame. When the pipette&#39;s length has increased by about 5 in, immediately remove the flame.</li>
		<li>Use small scissors or a glass cutter to trim the pulled end of the pipette at the point where the new pulled diameter is roughly twice that of the optical fiber used in the next section (see step 1.2). File down any sharp areas of the trimmed end using carborundum paper (Table of Materials). Rinse away any resulting small glass shards or dust using a squirt bottle of isopropyl alcohol followed by compressed air.</li>
	</ol>
	</li>
	<li>Pulling the optical fiber (Figure 1D)
	<ol>
		<li>Use a razor blade to sever the optical fiber, removing one SMA connector of a 200 &#181;m diameter SMA-terminated optical fiber (Table of Materials). Cut close to one of the SMA terminations. Then, use the razor blade to remove the next 5 cm of plastic and fiberglass jacketing so that the bare optical fiber is exposed and protrudes from the rest of the intact assembly.</li>
		<li>Use the butane torch to burn off the polyimide polymer coating from the glass fiber. Rinse with isopropanol. Wipe the bare glass fiber clean using a lint-free wipe.</li>
		<li>The next step is to mount the bare fiber so it can also be pulled with a flame. Mount the bare end of the optical fiber directly in a plier clamp (Table of Materials) on a table or shelf edge, letting the SMA-terminated end of the fiber hang toward the floor. Add the weights for pulling in the form of two small clamps (total weight: 10 g) on the jacketed end of the fiber about 12 in down from where the bare optical fiber is held in the plier clamp.</li>
		<li>To pull the fiber, start with the butane torch flame on. With the flame on, hold the butane torch 1 cm from the bare fiber so the flame is perpendicular to the optical fiber and 3 cm down vertically from where the plier clamp holds the bare fiber. Allow the fiber to stretch, pull, separate, and drop to the floor.</li>
		<li>Examine the resulting pulled optical fiber end. Gently sand away any irregularities with carborundum paper, clean with isopropyl alcohol and a lint-free wipe, and dry with compressed air. 
		NOTE:&#160;At this point, one may use a microscope to characterize and document the size and shape of the pulled fiber.</li>
		<li>Use a film-opaquing pen to darken the sides of the fiber and prevent the entry of stray light (Table of Materials). Gently pull the fiber across the tip of the pen, leaving only a small area at the tip uncovered.</li>
	</ol>
	</li>
	<li>Mounting the pulled fiber inside the glass pipette sleeve (Figure 1A)
	<ol>
		<li>Working under a stereomicroscope, carefully insert the tapered optical fiber from step 1.2 into the wide end of the altered glass pipette from step 1.1, and push the fiber through until ~1 mm of bare fiber is sticking out of the tapered end of the pipette.</li>
		<li>Using electrical tape, secure the jacketed end of the optical fiber to the wide end of the pipette.</li>
		<li>Put a drop of cyanoacrylate adhesive onto a small-gauge needle (Table of Materials). Carefully touch the drop of adhesive to the cut edge of the pulled pipette, avoiding the bare end of the pulled fiber.</li>
		<li>Allow capillary action to wick the adhesive into the pipette, wetting the area between the wall of the pipette and the optical fiber. Allow the adhesive to cure for at least 15 min before moving the probe assembly.</li>
	</ol>
	</li>
	<li>Modifying the fiber tip with a scattering sphere (Figure 1C)
	<ol>
		<li>Create the raw material for the scattering ball tip by mixing a drop of UV-curable adhesive with titanium dioxide powder (Table of Materials) at ~1:1 v/v.</li>
		<li>Attach the probe to a micromanipulator with a mounting rod holder (Table of Materials) such that the probe is in a horizontal orientation. Prepare a working reservoir of the scattering material by dipping the tip of a wire or needle in the adhesive/TiO2 mixture prepared above such that a droplet of adhesive forms. Mount the wire or needle with the droplet near the tip of the horizontal probe. It is convenient to do this using an alligator clip mounted to the bench.</li>
		<li>Deposit a scattering sphere on the tip of the probe. Use the micromanipulator to slowly and carefully push the tip of the probe&#39;s optical fiber into the droplet of adhesive/TiO2. Then, quickly withdraw the tip from the glue. Repeat two to three times until a spherical droplet of adhesive of the desired size is deposited on the tip of the optical fiber. 
		NOTE: The scattering sphere will be stable at diameters from two to eight times greater than the diameter of the tapered optical fiber. The final optimal size depends on the final desired application.</li>
		<li>Cure the sphere by connecting the SMA-terminated end of the optical fiber to a fiber-coupled UV light source (Table of Materials). 
		&#8203;NOTE: The light source used in this study had a power of 5.3 mW. At this power, the recommended cure time is at least 12 h or overnight.&#160;After curing is a good time to characterize and measure the size of the final optical probe tip.&#160;</li>
	</ol>
	</li>
</ol>
2. Preparing and mounting the tissue for optical measurements (Figure 2 and Figure 3)
<ol>
	<li>Prepare the dishes to mount the samples for an experiment. Heat the large end of a Pasteur pipette using a butane torch to make a punch to melt a 0.5 cm diameter hole in the bottom of a 35 mm x 10 mm plastic Petri dish. Seal the hole on the bottom side of the dish with electrical tape or lab tape. Make several at a time for efficiency.</li>
	<li>Prepare liquid gelatin (Table of Materials) according to the instructions on the package. 
	NOTE: Commercial food-grade flavorless gelatin from the grocery store is better for this purpose than chemical-grade gelatin from chemical suppliers because the chemical-grade gelatin is less optically clear and less mechanically reliable than the food product.</li>
	<li>Perform the dissection and preparation of the tissue that will be used. 
	NOTE: From experience, any flat tissue up to 1 cm thick can be successfully measured in this experiment using any dissection technique appropriate for the system of interest. For giant clam tissue, an 8 mm biopsy punch was used to make a flat, regular circle of clam tissue for use in the experiment12. For this system, only one sample could effectively be prepared at a time given the length of time required to complete a measurement to ensure that the sample was still in lifelike condition at the end of the experiment.</li>
	<li>Fill two Petri dishes from step 2.1 a quarter of the way with the liquid gelatin, and let cool to room temperature (RT) just short of gelation. In one dish, place the biopsy in the cushion of viscous gelatin that forms in the hole in the bottom of the dish. Use a Pasteur pipette to gently add RT gelatin around the biopsy until the Petri dish is full (Figure 3). Fill the second dish with gelatin to make a blank sample.</li>
	<li>Place the sample dishes into a refrigerator for about 10-30 min until the gelatin is fully cured and slightly elastic to the touch. 
	NOTE: In a cool room, this may not be necessary; when working in the tropics, this step is required to fully cure the gelatin.</li>
	<li>Making a mount for the Petri dish on the micromanipulator (Figure 2)
	<ol>
		<li>Cut a 6 cm x 6 cm square of 1/4 in thick plexiglass (Table of Materials).</li>
		<li>Drill or melt a hole in the plastic square the same diameter as the Petri dish (~35 mm). Ensure that the Petri dish fits snugly in this hole and is held in place with friction. Add tape to the edges of the dish to slightly increase the size and frictional contact.</li>
		<li>Drill or melt a 1/4 in diameter hole in the corner of the plastic square. Use this hole to attach the square to the micromanipulator using a 1/4 in screw and bolt.</li>
		<li>Cover the plastic square with black electrical tape to reduce the stray light reaching the probe. Similarly, use tape to create a skirt around the sides of the square extending below the bottom of the inserted dish (&#62;10 mm) to reduce stray light (Figure 2B).</li>
		<li>Use a bolt and appropriate hardware to attach the Petri dish holder to a micromanipulator that allows for three-dimensional adjustments and well-resolved vertical movement over an extent greater than the thickness of the samples (part 1 and part 2 of the manipulator mentioned in the Table of Materials are good examples). Affix this micromanipulator to an optical breadboard.</li>
	</ol>
	</li>
</ol>
3. Measurement setup for tissue biopsies
<ol>
	<li>Mount a light source above the area where the measurement takes place such that the light is collimated when it reaches the plane where the sample will be placed (Figure 2A). For example, attach a 5 cm collimating lens (Table of Materials) to a 5 mm liquid light guide (Table of Materials), and elevate above the sample by &#62;0.6 m using a vice and frame attached to an optical breadboard (Table of Materials). Then, connect the light guide to a broadband light source, such as the plasma light source listed in the Table of Materials.</li>
	<li>Attach the probe to a vertically oriented mount such that it points toward the light source. Secure the probe with clamps, hose clamps, or tape to an optical table post. 
	NOTE: In the giant clam experiment, a specifically designed rod holder, such as the micromanipulator listed in the Table of Materials, is used. In this work, it was secured by magnets to an optical breadboard (Table of Materials). The additional degrees of freedom for alignment obtained by having both the probe and the sample on manipulators are convenient but not required. Be careful not to over-clamp to the post, as this can break the pipette housing.</li>
	<li>Position the sample manipulator near the vertically mounted probe. Use the sample manipulator to adjust the position of the sample stage such that the probe is centered in the 0.5 cm opening at the bottom of the Petri dish. Use extreme caution to avoid touching or bumping the tip of the mounted probe. 
	NOTE: A boom-mounted stereomicroscope positioned over the sample area can be useful. This way, the top-down alignment of the probe tip, stage, and sample can be viewed before starting an experiment (Figure 2A).</li>
	<li>Connect the SMA-terminated end of the probe&#39;s optical fiber to a USB fiber-optic spectrometer (Table of Materials), and connect the spectrometer to the computer using a USB cable.</li>
</ol>
4. Data collection
<ol>
	<li>Experimental setup
	<ol>
		<li>Have the probe and manipulators in the exact aligned position in which the data will be collected, as in step 3.4.</li>
		<li>Use a cotton swab or fine-gauge needle to carefully apply a small amount of silicone lubricant (Table of Materials) to the part of the probe that will be inserted into the tissue to lower the friction between the sides of the probe and the tissue sample. Reapply the silicone lubricant before each baseline or tissue measurement.</li>
		<li>Remove the tape covering the hole in the blank sample Petri dish, and place it in the sample holder (step 2.7), making sure it is held securely in place by friction.</li>
		<li>Use the micromanipulator to lower the sample onto the probe. Allow the probe to enter the gelatin via the hole on the underside of the petri dish. Continue until the probe is approximately 5 mm from the bottom of the gelatin layer.</li>
		<li>Turn on the light source. Using the spectrometer software, adjust the spectrometer integration time until the signal is as high as possible but not saturating. Set the number of averaged scans between two and five and the smoothing pixels value to six. Usable integration times at this stage can vary between 1-50 ms for this baseline.</li>
	</ol>
	</li>
	<li>Collecting the reference spectra
	<ol>
		<li>Collect a dark measurement.
		<ol>
			<li>Once the spectrometer parameters have been set with the blank sample, detach the probe fiber from the spectrometer, and cover the port with the opaque metal cover that came with the spectrometer.</li>
			<li>Obtain a dark measurement from the spectrometer (often using the dark lightbulb icon in the GUI) to characterize the noise of the spectrometer with no input light. Save this measurement according to the data acquisition strategy.</li>
		</ol>
		</li>
		<li>Collect a lamp measurement. Reconnect the probe fiber to the spectrometer, wait for the signal to stabilize, and acquire another spectrum. This measurement characterizes the light reaching the probe through the gelatin in the absence of tissue.</li>
	</ol>
	</li>
	<li>Collecting the tissue spectra
	<ol>
		<li>Remove the tape on the bottom of the sample Petri dish, and place it in the sample holder.</li>
		<li>Align the sample dish with the probe using three-dimensional manipulation so that the tissue biopsy is centered above the tip of the probe.</li>
		<li>Apply silicone gel to the probe (see 4.1.2), and slowly lower the sample onto the probe until the probe tip has passed through the gelatin supporting the tissue sample and has just entered the tissue sample. 
		NOTE: Some spectrometers and computer programs allow real-time monitoring of the detected light. Use this to determine when the probe enters the tissue. The spectrum intensity will abruptly decrease as soon as the probe tip is in direct contact with the tissue.</li>
		<li>Verify that the integration time is appropriate for the measurement by ensuring that the measured spectrum is neither too noisy nor saturated. Change the integration time if necessary.
		<ol>
			<li>Any time the integration time is adjusted, perform and store a new dark measurement (step 4.2.1). Do not change the number of averaged scans or the number of smoothed pixels.</li>
			<li>After finding suitable measurement parameters, take the measurements and save the spectrum.</li>
		</ol>
		</li>
		<li>Move the sample down a specified distance using the micromanipulator. For the manipulator used for this study, each small tick mark was 0.001 in or 25.4 &#181;m. The sample was moved down through five tick marks, or 127 &#181;m, for each measurement. Repeat step 4.3.4.</li>
		<li>Continue to save the spectra at each vertical position within the tissue. 
		NOTE: For the system described here, the required integration times varied between 1 ms to 5 s for the measurements within a single tissue sample. Eventually, the probe tip will exit the top of the tissue and be visible there (Figure 4A). This is the final measurement and characterizes the light incident at the top of the tissue.</li>
	</ol>
	</li>
	<li>Loading and processing the data
	<ol>
		<li>Use the spectrometer&#39;s software to convert all the collected spectra (dark spectra, lamp spectra, and tissue measurements) to delimited text files.</li>
		<li>Using Matlab or a coding language of choice, load all the measurement files for a sample. For each tissue measurement spectrum, subtract the dark spectrum with the matching integration time, and then divide it by the integration time. 
		NOTE: In this study, a Matlab script was used for loading and processing the data (Supplementary File 1).</li>
		<li>Calculate the percentage of incident light reaching each position in the tissue by dividing the spectra obtained with the probe inside the tissue by the baseline spectrum obtained in empty gelatin. 
		NOTE: The measurement at the top of the tissue (step 4.3.6) should be very similar to the baseline measurement in gelatin (step 4.1) and, when divided, should be close to 100%. Depending on the experimental context, either lamp spectrum (the spectrum from empty gelatin or that from the very top of the tissue) can be used as a reference. However, it is still important to collect a lamp spectrum before starting the experiment. This is because, if the probe breaks or the experiment otherwise fails before reaching the top of the tissue, the data obtained before the failure are still usable, which is not the case if there is no corresponding initial lamp reference spectrum.</li>
		<li>Visualize the data; contour plots can be helpful (Figure 4B).</li>
	</ol>
	</li>
</ol>

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There are no conflicts of interest.

This protocol describes a technique for systematically characterizing the optical environment through a large volume of living tissue with a spatial resolution approximately on the scale of single cells. This inexpensive, flexible, and field-tractable method could be useful to any researchers studying the propagation of light within living systems. From experience, compared to existing methods7, these probes require a little more practice and skill to build but result in less tissue damage and the...

Surprisingly, land animals and shallow ocean dwellers have enough light within their body for visual physiology and even photosynthesis. For example, the light levels in the center of a mouse&#39;s head (outside of the strong hemoglobin absorbance bands) are attenuated by three or four orders of magnitude relative to the outside world. This is roughly the difference between the light levels indoors and outside. So, the opacity of a tissue or material due to strong scattering is not the same as opacity due to strong light absorption. Light can keep propagating over long distances in a strongly forward-scattering system, similar to light propagating through aquatic systems with high concentrations of cells and particles1.&#160;This observation is particularly salient in light of the fact that opsin proteins are near-ubiquitously expressed in all tissues of all animals. Thus, it is important to understand how and where light is attenuated and scattered within living tissue. However, unlike aquatic systems, with living tissue, it is impossible to immerse an instrument in the water column and obtain radiance and irradiance measurements, and a new technique is necessary.
Other methods previously used to characterize the absorption and scattering properties of living tissue include measuring tissue reflectance probes and/or integrating spheres2,3, microscopic methods such as scanning confocal microscopy4, measuring the diffusion of laser light on the surface5, and modeling techniques such as Monte Carlo radiative transfer6. The experimental methods mentioned often require specific, large, and expensive equipment or detailed knowledge about tissue structure and are generally limited in their ability to characterize the spatial structure of light deep within the tissue.
There are also similar probe-based methods that use a hypodermic needle to insert an optical fiber through tissue7,8,9. In our experience, modified needles are effective at puncturing tissue but require considerable force and generally tear delicate tissues when passaging through densely packed cells. Therefore, these needles generally require a surgical procedure to insert more than a millimeter or so into a tissue layer. The method described here, using a lubricated, pulled glass support, is capable of sliding between cells with minimal wounding of the tissue and without additional surgery.
This manuscript presents a method inspired by the work of Jorgenson and colleagues on measuring light within algal mats10,11, using glass-supported optical microprobes and portable electronics that are amenable to probing deep into dense tissue and to construction and use in the field. These probes can be constructed to characterize scalar irradiance (light hitting a point from all directions) and downwelling radiance (light intersecting a horizontal plane) inside living tissue at high spatial resolutions. These probes were originally developed to measure radiative transfer within the tissue of photosymbiotic giant clams12. Standard measurements of absorption and transmission of the total tissue were not enough to characterize the photosynthetic performance of the tissue, since it makes a big difference whether all the incident light is absorbed by a few cells experiencing high intensity at the surface of the tissue or many cells experiencing low intensities throughout the volume of the tissue. In a second project, these probes were used to measure in vivo irradiance within a mouse&#39;s brain13,14, thereby characterizing the light environment of opsins expressed deep within the brain. These micro-probes are both small and sensitive enough to measure irradiance within mouse brain tissue with all the fur, skin, and bone intact and demonstrate that physiological light levels are easily high enough to stimulate deep-brain opsins.
This micro-optical probe and measuring setup could be useful to researchers needing to quantify and characterize the light internal to biological tissue, particularly for a more nuanced understanding of photosynthesis or the functions of visual pigments expressed outside of the eyes. This method can be used alone or in conjunction with other techniques to fully characterize the optical properties and light propagation within living tissue at a low cost, with small portable equipment built in-house and with task-dependent adjustable parameters.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1&#34; travel ball bearing center+D11+A2:D31+A2:A2:D31</td>
			<td>Edmond Optics</td>
			<td>37-935</td>
			<td>Part 2 of manipulator for lowering sample</td>
		</tr>
		<tr>
			<td>1/4&#34; thick acrylic sheet</td>
			<td>McMaster-Carr</td>
			<td>8505K754</td>
			<td>For making Petri dish holder</td>
		</tr>
		<tr>
			<td>3/4&#34; mini spring clamp</td>
			<td>Anvil</td>
			<td>99693</td>
			<td>Use as weight for pulling optical fiber</td>
		</tr>
		<tr>
			<td>8 mm biopsy punch</td>
			<td>Fisher Scientific</td>
			<td>NC9324386</td>
			<td>For tissue sample</td>
		</tr>
		<tr>
			<td>Butane Torch</td>
			<td>McMaster-Carr</td>
			<td>MT-51</td>
			<td>Heat source for pulling fiber and pipette</td>
		</tr>
		<tr>
			<td>Collimating lens</td>
			<td>Thorlabs</td>
			<td>LLG5A1-A</td>
			<td>To collimate light source through liquid light guide</td>
		</tr>
		<tr>
			<td>Compressed air</td>
			<td>McMaster-Carr</td>
			<td>7437K35</td>
			<td>For drying pulled fiber and pipette</td>
		</tr>
		<tr>
			<td>Cyanoacrylate glue - liquid</td>
			<td>McMaster-Carr</td>
			<td>66635A31</td>
			<td>For securing tapered fiber end at top of pulled pipette</td>
		</tr>
		<tr>
			<td>Electrical tape</td>
			<td>McMaster-Carr</td>
			<td>76455A21</td>
			<td>For securing fiber in pipette and for adding grip to clamps</td>
		</tr>
		<tr>
			<td>Fine grade carborundum paper</td>
			<td>McMaster-Carr</td>
			<td>4649A24</td>
			<td>Small triangle on exacto knife holder works well</td>
		</tr>
		<tr>
			<td>Gelatin</td>
			<td>Knox</td>
			<td>10043000048679</td>
			<td>For securing the tissue biopsy in the petri dish</td>
		</tr>
		<tr>
			<td>Glass Pasteur Pipete</td>
			<td>Fisher Scientific</td>
			<td>13-678-20B</td>
			<td>Disposable glass pipette 5.75&#34; in length</td>
		</tr>
		<tr>
			<td>Insulin syringes, 31G needle</td>
			<td>BD</td>
			<td>320440</td>
			<td>For applying glue</td>
		</tr>
		<tr>
			<td>Isopropanol</td>
			<td>McMaster-Carr</td>
			<td>54845T42</td>
			<td>For cleaning pulled fiber and pipette</td>
		</tr>
		<tr>
			<td>Kimwipe</td>
			<td>Cole-Parmer</td>
			<td>SKU 33670-04</td>
			<td>For wiping optical fiber and glass pipette clean</td>
		</tr>
		<tr>
			<td>LED driver</td>
			<td>Thorlabs</td>
			<td>LEDD1B</td>
			<td>For powering the UV LEDs</td>
		</tr>
		<tr>
			<td>Light source for measurements</td>
			<td>Cole-Parmer</td>
			<td>UX-78905-05</td>
			<td>Low heat white light source for measurements</td>
		</tr>
		<tr>
			<td>Linear metric X-Y-Z axis rack and pinion stage</td>
			<td>Edmond Optics</td>
			<td>55-023</td>
			<td>Part 1 of manipulator for lowering sample</td>
		</tr>
		<tr>
			<td>Liquid light guide</td>
			<td>Thorlabs</td>
			<td>LLG5-4T</td>
			<td>For light source in measurements</td>
		</tr>
		<tr>
			<td>Magnetic feet</td>
			<td>Siskiyou</td>
			<td>MGB 8-32</td>
			<td>For use with magnetic strips</td>
		</tr>
		<tr>
			<td>Magnetic strips</td>
			<td>Siskiyou</td>
			<td>MS-6.0</td>
			<td>For mounting magnetically to breadboard</td>
		</tr>
		<tr>
			<td>Manipulator #1</td>
			<td>Siskiyou</td>
			<td>MX10R</td>
			<td>4-axis manipulator with pipette holder</td>
		</tr>
		<tr>
			<td>Opaquer pen, small</td>
			<td>WindowTint</td>
			<td>TOP01</td>
			<td>For opaquing side of optical fiber to prevent stray light from enter probe</td>
		</tr>
		<tr>
			<td>Optical breadboard</td>
			<td>Edmond Optics</td>
			<td>03-640</td>
			<td>For stable affixation of probe holder, sample, microscope and light source</td>
		</tr>
		<tr>
			<td>Optical fibers</td>
			<td>Ocean Optics</td>
			<td>P-100-2-UV-VIS</td>
			<td>About 4 fibers are good to have</td>
		</tr>
		<tr>
			<td>Plasma light source</td>
			<td>Thorlabs</td>
			<td>HPLS345</td>
			<td>For tissue radiometry measurements</td>
		</tr>
		<tr>
			<td>Plastic plier clamp</td>
			<td>McMaster-Carr</td>
			<td>5070A11</td>
			<td>Plier clamp used for weight in pulling pipette</td>
		</tr>
		<tr>
			<td>Polystyrene Petri dishes</td>
			<td>Thomas scientific</td>
			<td>3488N10</td>
			<td>Sample holders, enough volume to hold sample thickness plus ~10 mm of gelatin on top</td>
		</tr>
		<tr>
			<td>Razor blades</td>
			<td>McMaster-Carr</td>
			<td>3962A3</td>
			<td>For stripping jacketing from optical fiber</td>
		</tr>
		<tr>
			<td>Silicone oil lubricant</td>
			<td>Thomas scientific</td>
			<td>1232E30</td>
			<td>For reducing friction between probe and tissue</td>
		</tr>
		<tr>
			<td>Software for analyzing data</td>
			<td>Matlab</td>
			<td></td>
			<td>Chosen software for data analysis</td>
		</tr>
		<tr>
			<td>Spectrometer + spectrometer software</td>
			<td>Avantes</td>
			<td>AvaSpec-2048L</td>
			<td>Spectrometer can be any brand, this one is compatible with sma-terminated optical fibers and comes with its own software for running the spectrometer</td>
		</tr>
		<tr>
			<td>Titanium dioxide powder</td>
			<td>Sigma Aldrich</td>
			<td>718467-100G</td>
			<td>For making scattering sphere</td>
		</tr>
		<tr>
			<td>Toolour tabletop clip</td>
			<td>Toolour</td>
			<td>Toolour0004</td>
			<td>For holding pipette while pulling and for holding finished probes</td>
		</tr>
		<tr>
			<td>Trigger-action bar clamps</td>
			<td>mcMaster-Carr</td>
			<td>51755A2</td>
			<td>Good for holding optical fibers while pulling or curing</td>
		</tr>
		<tr>
			<td>UV curable adhesive</td>
			<td>Delo Photobond</td>
			<td>GB368</td>
			<td>For making scattering sphere</td>
		</tr>
		<tr>
			<td>UV light source</td>
			<td>Thorlabs</td>
			<td>M365FP1</td>
			<td>Light source for curing adhesive in scattering ball, this one is sma-fiber compatible, higher intensity = less cure time</td>
		</tr>
		<tr>
			<td>White LED light source</td>
			<td>Thorlabs</td>
			<td>MCWHF2</td>
			<td>For characterizing pulled fiber and scattering sphere</td>
		</tr>
	</tbody>
</table>

an intra-tissue radiometry microprobe for measuring radiance in situ in living tissue

Organisms appear opaque largely because their outer tissue layers are strongly scattering to incident light; strongly absorbing pigments, such as blood, typically have narrow absorbances, such that the mean free path of light outside the absorbance peaks can be quite long. As people cannot see through tissue, they generally imagine that tissues like the brain, fat, and bone contain little or no light. However, photoresponsive opsin proteins are expressed within many of these tissues, and their functions are poorly understood. Radiance internal to tissue is also important for understanding photosynthesis. For example, giant clams are strongly absorbing yet maintain a dense population of algae deep in the tissue. Light propagation through systems like sediments and biofilms can be complex, and these communities can be major contributors to ecosystem productivity. Therefore, a method for constructing optical micro-probes for measuring scalar irradiance (photon flux intersecting a point) and downwelling irradiance (photon flux crossing a plane perpendicularly) to better understand these phenomena inside living tissue has been developed. This technique is also tractable in field laboratories. These micro-probes are made from heat-pulled optical fibers that are then secured in pulled glass pipettes. To change the angular acceptance of the probe, a 10-100 &#181;m sized sphere of UV-curable epoxy mixed with titanium dioxide is then secured to the end of a pulled, trimmed fiber. The probe is inserted into living tissue, and its position is controlled using a micromanipulator. These probes are capable of measuring in situ tissue radiance at spatial resolutions of 10-100 &#181;m or on the scale of single cells. These probes were used to characterize the light reaching the adipose and brain cells 4 mm below the skin of a living mouse and to characterize the light reaching similar depths within living algae-rich giant clam tissue.

This protocol describes the procedure for constructing a micro-optical probe that can be used to measure the downwelling irradiance (the light reaching a point from one direction) or, with the addition of a light-scattering spherical tip, to measure the scalar irradiance (the light reaching a point from all directions). These probes can measure irradiance at spatial resolutions approaching the length scales of single cells inside living tissue. This protocol also describes a representative method for preparing a tissue s...

Watch this Scientific Journal Video about An Intra-Tissue Radiometry Microprobe for Measuring Radiance In Situ in Living Tissue at JoVE.com

An Intra-Tissue Radiometry Microprobe for Measuring Radiance In Situ in Living Tissue

In this paper, a method for measuring radiance in situ in living tissue is described. This work includes details of the construction of micro-scale probes for different measurements of radiance and irradiance, provides guidance for mounting tissue for the characterization of radiance, and outlines computational methods for analyzing the resulting data.

An Intra-Tissue Radiometry Microprobe for Measuring Radiance In Situ in Living Tissue

Organisms appear opaque largely because their outer tissue layers are strongly scattering to incident light; strongly absorbing pigments, such as blood, ...

an-intra-tissue-radiometry-microprobe-for-measuring-radiance-situ

Research

JoVE Journal

Bioengineering

522 Views.  Yale University. In this paper, a method for measuring radiance in situ in living tissue is described. This work includes details of the construction of micro-scale probes for different measurements of radiance and irradiance, provides guidance for mounting tissue for the characterization of radiance, and outlines computational methods for analyzing the resulting data. 

Video: An Intra-Tissue Radiometry Microprobe for Measuring Radiance In Situ in Living Tissue

Quantitatively Measuring In situ Flows using a Self-Contained Underwater Velocimetry Apparatus (SCUVA)

The ability to directly measure velocity fields in a fluid environment is necessary to provide empirical data for studies in fields as diverse as oceanography, ecology, biology, and fluid mechanics. Field measurements introduce practical challenges such as environmental conditions, animal availability, and the need for field-compatible measurement techniques. To avoid these challenges, scientists typically use controlled laboratory environments to study animal-fluid interactions. However, it is reasonable to question whether one can extrapolate natural behavior (i.e., that which occurs in the field) from laboratory measurements. Therefore, in situ quantitative flow measurements are needed to accurately describe animal swimming in their natural environment.
We designed a self-contained, portable device that operates independent of any connection to the surface, and can provide quantitative measurements of the flow field surrounding an animal. This apparatus, a self-contained underwater velocimetry apparatus (SCUVA), can be operated by a single scuba diver in depths up to 40 m. Due to the added complexity inherent of field conditions, additional considerations and preparation are required when compared to laboratory measurements. These considerations include, but are not limited to, operator motion, predicting position of swimming targets, available natural suspended particulate, and orientation of SCUVA relative to the flow of interest. The following protocol is intended to address these common field challenges and to maximize measurement success.

Quantitatively Measuring In situ Flows using a Self-Contained Underwater Velocimetry Apparatus SCUVA

This protocol provides instructions on how to use a self-contained underwater velocimetry apparatus (SCUVA), which is designed for quantification of in situ animal-generated flows. In addition, this protocol addresses challenges posed by field conditions, and includes operator motion, predicting position of animals, and orientation of SCUVA.

The ability to directly measure velocity fields in a fluid environment is necessary to provide empirical data for studies in fields as diverse as ...

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Generation of a Three-dimensional Full Thickness Skin Equivalent and Automated Wounding

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models reflect the human wound situation better than animal models. Although two-dimensional models are widely used to investigate processes such as cellular migration and proliferation, models that are more complex are required to gain a deeper knowledge about wound healing. Besides a suitable model system, the generation of precise and reproducible wounds is crucial to ensure comparable results between different test runs. In this study, the generation of a three-dimensional full thickness skin equivalent to study wound healing is shown. The dermal part of the models is comprised of human dermal fibroblast embedded in a rat-tail collagen type I hydrogel. Following the inoculation with human epidermal keratinocytes and consequent culture at the air-liquid interface, a multilayered epidermis is formed on top of the models. To study the wound healing process, we additionally developed an automated wounding device, which generates standardized wounds in a sterile atmosphere.

The goal of this protocol is to build up a three-dimensional full thickness skin equivalent, which resembles natural skin. With a specifically constructed automated wounding device, precise and reproducible wounds can be generated under maintenance of sterility.

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models ...

Eye Irritation Test (EIT) for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial (RhCE) Tissue Model

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and accurate assessment of ocular toxicity in man are needed. To address this need, we have developed an eye irritation test (EIT) which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model that is based on normal human cells. The EIT is able to separate ocular&#160;irritants and corrosives (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category). The test utilizes two separate protocols, one designed for liquid chemicals and a second, similar protocol for solid test articles. The EIT prediction model uses a single exposure period (30 min for liquids, 6 hr for solids) and a single tissue viability cut-off (60.0% as determined by the MTT assay). Based on the results for 83 chemicals (44 liquids and 39 solids) EIT achieved 95.5/68.2/ and 81.8% sensitivity/specificity and accuracy (SS&#38;A) for liquids, 100.0/68.4/ and 84.6% SS&#38;A for solids, and 97.6/68.3/ and 83.1% for overall SS&#38;A. The EIT will contribute significantly to classifying the ocular irritation potential of a wide range of liquid and solid chemicals without the use of animals to meet regulatory testing requirements. The EpiOcular EIT method was implemented in 2015 into the OECD Test Guidelines as TG 492.

Eye Irritation Test EIT for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial RhCE Tissue Model

We have developed an eye irritation test which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model. The test is able to discriminate between ocular irritant and corrosive materials (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category).

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and ...

Sandwich-like Microenvironments to Harness Cell/Material Interactions

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. However in vivo, most of the cells are completely surrounded by the extracellular matrix (ECM), resulting in a three-dimensional (3D) distribution of receptors. This may trigger differences in the outside-in signaling pathways and thus in cell behavior.
This article shows that stimulating the dorsal receptors of cells already adhered to a 2D substrate by overlaying a film of a new material (a sandwich-like culture) triggers important changes with respect to standard 2D cultures. Furthermore, the simultaneous excitation of ventral and dorsal receptors shifts cell behavior closer to that found in 3D environments. Additionally, due to the nature of the system, a sandwich-like culture is a versatile tool that allows the study of different parameters in cell/material interactions, e.g., topography, stiffness and different protein coatings at both the ventral and dorsal sides. Finally, since sandwich-like cultures are based on 2D substrates, several analysis procedures already developed for standard 2D cultures can be used normally, overcoming more complex procedures needed for 3D systems.

The following protocol describes the procedure to assemble sandwich-like cultures to be used as an intermediate stage between bi-dimensional (2D) and three-dimensional (3D) cellular environments. The engineered systems can have applications in microscopy, biomechanics, biochemistry and cell biology assays.

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. ...

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen Presenting Cells (APCs). This is despite T-cell antigen receptors (TCRs) exerting usually a moderate affinity (&#181;M range) to antigen when binding is measured in vitro1. In view of the molecular and cellular parameters contributing to T-cell antigen sensitivity, a microscopy-based methodology has been developed as a means to monitor TCR-pMHC binding in situ, as it occurs within the synapse of a live T-cell and an artificial and functionalized glass-supported planar lipid bilayer (SLB), which mimics the cell membrane of an Antigen presenting Cell (APC) 2. Measurements are based on F&#246;rster Resonance Energy Transfer (FRET) between a blue- and red-shifted fluorescent dye attached to the TCR and the pMHC. Because the efficiency of FRET is inversely proportional to the sixth power of the inter-dye distance, one can employ FRET signals to visualize synaptic TCR-pMHC binding. The sensitive of the microscopy approach supports detection of single molecule FRET events. This allows to determine the affinity and off-rate of synaptic TCR-pMHC interactions and in turn to interpolate the on-rate of binding. Analogous assays could be applied to measure other receptor-ligand interactions in their native environment.

Measuring TCR-pMHC Binding In Situ using a FRET-based Microscopy Assay

This manuscript describes how to conduct (single molecule) F&#246;rster Resonance Energy Transfer (FRET)- based assays to measure the binding dynamics between T-cell antigen receptor (TCR) and antigenic peptide-loaded MHC molecules as they occur within the immunological synapse of a T-cell in contact with a functionalized planar supported lipid bilayer.

T-cells are remarkably specific and effective when recognizing antigens in the form of peptides embedded in MHC molecules (pMHC) on the surface of Antigen ...

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and trade one defect for another. Tissue engineering holds the promise to address this increasing demand. However, most tissue engineering strategies fail to generate stable and functional tissue substitutes because of poor vascularization. This paper focuses on an in vivo tissue engineering chamber model of intrinsic vascularization where a perfused artery and a vein either as an arteriovenous loop or a flow-through pedicle configuration is directed inside a protected hollow chamber. In this chamber-based system angiogenic sprouting occurs from the arteriovenous vessels and this system attracts ischemic and inflammatory driven endogenous cell migration which gradually fills the chamber space with fibro-vascular tissue. Exogenous cell/matrix implantation at the time of chamber construction enhances cell survival and determines specificity of the engineered tissues which develop. Our studies have shown that this chamber model can successfully generate different tissues such as fat, cardiac muscle, liver and others. However, modifications and refinements are required to ensure target tissue formation is consistent and reproducible. This article describes a standardized protocol for the fabrication of two different vascularized tissue engineering chamber models in vivo.

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

This is a guideline for constructing in vivo vascularized tissue using a microsurgical arteriovenous loop or a flow-through pedicle configuration inside a tissue engineering chamber. The vascularized tissues generated can be employed for organ regeneration and replacement of tissue defects, as well as for drug testing and disease modeling.

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Melt Electrospinning Writing of Three-dimensional Poly(&#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology with great potential for biomedical applications. The technique facilitates the direct deposition of biocompatible polymer fibers to fabricate well-ordered scaffolds in the sub-micron to micro scale range. The establishment of a stable, viscoelastic, polymer jet between a spinneret and a collector is achieved using an applied voltage and can be direct-written. A significant benefit of a typical porous scaffold is a high surface-to-volume ratio which provides increased effective adhesion sites for cell attachment and growth. Controlling the printing process by fine-tuning the system parameters enables high reproducibility in the quality of the printed scaffolds. It also provides a flexible manufacturing platform for users to tailor the morphological structures of the scaffolds to their specific requirements. For this purpose, we present a protocol to obtain different fiber diameters using melt electrospinning writing (MEW) with a guided amendment of the parameters, including flow rate, voltage and collection speed. Furthermore, we demonstrate how to optimize the jet, discuss often experienced technical challenges, explain troubleshooting techniques and showcase a wide range of printable scaffold architectures.

Melt Electrospinning Writing of Three-dimensional Poly(&amp;#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This protocol serves as a comprehensive guideline to fabricate scaffolds via electrospinning with polymer melts in a direct writing mode. We systematically outline the process and define the appropriate parameter settings for achieving targeted scaffold architectures.

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology ...

A Novel Surgical Technique As a Foundation for In Vivo Partial Liver Engineering in Rat

Organ engineering is a novel strategy to generate liver organ substitutes that can potentially be used in transplantation. Recently, in vivo liver engineering, including in vivo organ decellularization followed by repopulation, has emerged as a promising approach over ex vivo liver engineering. However, postoperative survival was not achieved. The aim of this study is to develop a novel surgical technique of in vivo selective liver lobe perfusion in rats as a prerequisite for in vivo liver engineering. We generate a circuit bypass only through the left lateral lobe. Then, the left lateral lobe is perfused with heparinized saline. The experiment is performed with 4 groups (n = 3 rats per group) based on different perfusion times of 20 min, 2 h, 3 h, and 4 h. Survival, as well as the macroscopically visible change of color and the histologically determined absence of blood cells in the portal triad and the sinusoids, is taken as an indicator for a successful model establishment. After selective perfusion of the left lateral lobe, we observe that the left lateral lobe, indeed, turned from red to faint yellow. In a histological assessment, no blood cells are visible in the branch of the portal vein, the central vein, and the sinusoids. The left lateral lobe turns red after reopening the blocked vessels. 12/12 rats survived the procedure for more than one week. We are the first to report a surgical model for in vivo single liver lobe perfusion with a long survival period of more than one week. In contrast to the previously published report, the most important advantage of the technique presented here is that perfusion of 70% of the liver is maintained throughout the whole procedure. The establishment of this technique provides a foundation for in vivo partial liver engineering in rats, including decellularization and recellularization.

A Novel Surgical Technique As a Foundation for In Vivo Partial Liver Engineering in Rat

We establish a novel surgical technique for an in vivo single liver lobe perfusion model in rat as a prerequisite for further studying in vivo partial liver engineering in the future.