This work was supported by NIH K08EY027006, R01EY030564, UH3NS100614, Research Grants from Carl Zeiss Meditec Inc (Dublin, CA) and Unrestricted Department Funding from Research to Prevent Blindness (New York, NY).

This study was approved by the University of Southern California Institutional Review Board and adhered to the tenets of the Declaration of Helsinki.
1. Setup of Gas Non-rebreathing Apparatus
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60948/60948fig1v2.jpg" /> 
Figure 1: Diagram of the non-rebreathing apparatus.&#160;The full setup has been broken into three separate units according to their function and the frequency with which they are dealt with independently. These include: the Air-Control Unit, the Non-rebreathing Unit, and the Subject/Imaging Device Unit <a href="https://www.jove.com/files/ftp_upload/60948/60948fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Apparatus assembly
	<ol>
		<li>Connect the Douglas bag (Figure 1, #1) to the three-way valve (#3) at a selective inlet port via the 35 mm inner-diameter tube (#2; see Table of Materials) with adapter (#2*). This combination will be called the &#8220;Air Control Unit&#8221; as shown in Figure 1.</li>
		<li>Connect the two-way non-rebreathing valve (#6) to the elbow joint connector (#7) at the non-rebreathing valve&#8217;s mouth port. Form the connection using a rubber tube (#5) fitted with an adapter (#4).</li>
		<li>Connect the elbow joint to the gas delivery tubing (#8). This setup, including the non-rebreathing valve (#6), in-house tubing (#5), adapters (#4), elbow joint (#7), and gas delivery tubing (#8) will be called the &#8220;Non-rebreathing Unit&#8221;. 
		NOTE: Minimize the amount of dead space between the subject&#8217;s mouth and the diaphragm of the two-way non-rebreathing valve (#6).</li>
		<li>Connect the Air Control Unit at the outlet port of the three-way valve (#3) to the Non-rebreathing Unit at the inlet port of the two-way non-rebreathing valve (#6). Make the connection using additional rubber tubing (#5) and adapters (#4) as those described earlier that allow the pieces to be inserted into one another.</li>
		<li>Seal all&#160;loose connections by wrapping the joints with sealing tape to ensure a hermetic fit.</li>
		<li>Connect the gas delivery tubing (#8) at its open end to a mouthpiece (#9) as shown in the Subject/Imaging Device Unit of Figure 1. 
		NOTE: This step (1.1.6) can be deferred until the subject testing is ready to begin (Step 3.5).</li>
	</ol>
	</li>
	<li>Preparation of the Air Control Unit for gas non-rebreathing
	<ol>
		<li>Isolate the Air Control Unit by disconnecting it from any in-house tubing (#5) or adapters (#4) if it is not already separated.</li>
		<li>Ensure the Douglas bag (#1) is empty or empty the Douglas bag (#1) of any air by systematically rolling-up the bag from the distal end towards the bag&#8217;s inlet port with the three-way valve (#3) set to Configuration 1 as shown in Figure 1.</li>
		<li>Fill the Douglas bag (#1) with the appropriate gas mixture.
		<ol>
			<li>If only room-air non-rebreathing is intended, set the three-way valve to Configuration 2 (shown in Figure 1) and do not fill the Douglas bag (#1). Otherwise continue with the steps that comprise Step 1.2.3.</li>
			<li>Connect the Air Control Unit (shown in Figure 1) at the outlet port of the three-way valve (#3) to a gas-cylinder (containing the desired air-mixture) using the appropriate adapters and tubing. Use a cuff adapter to mount a 1/8&#8221; gas filling tube to the outer diameter of the three-way valve (#3).</li>
			<li>Set the three-way valve assembly to Configuration 1 (as shown in Figure 1) to allow the intended gas to flow from the storage cylinder into the Douglas bag (#1). Open the gas cylinder.</li>
			<li>Once the Douglas bag (#1) is filled to the intended volume (usually half-filled), close the gas cylinder outlet and set the three-way valve to Configuration 2, which isolates the gas within the Douglas bag (#1). Disconnect the Air Control Unit from any tubing used to fill the Douglas bag (#1).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Preparing the Subject for Imaging
<ol>
	<li>After the subject consents to participate in the study, sit the subject behind the OCTA imaging device. Explain the testing procedures to the subject.</li>
	<li>Confirm the subject&#8217;s medical history to ensure that the subject has no existing medical conditions that increase the risk of participating in the study. 
	NOTE: Pre-existing cardiovascular or pulmonary diseases are risk factors for which subjects may be excluded from participating. It is essential that the subject understand that they can stop the procedure at any time for any reason such as feeling lightheaded or some additional unexpected discomfort.</li>
	<li>Determine the eye to be assessed as per the testing protocol. One eye only may be imaged to limit the testing time and minimize the potential discomforts from the gas non-rebreathing.</li>
	<li>Consider eye dilation if the subject has a pupil size of about 2.5 mm or less. Although dilation is not mandatory, it enhances the chances of acquiring good quality images. To dilate, instill one drop each of 0.5% proparacaine hydrochloride ophthalmic solution, 1% tropicamide ophthalmic solution and 2.5% phenylephrine hydrochloride ophthalmic solution. Full dilation should occur within 10&#8211;15 min.</li>
</ol>
3. Gas Provocation Experiment and Image Acquisition
<ol>
	<li>Create a profile for the patient in the OCTA machine.</li>
	<li>Wear gloves.</li>
	<li>Wipe down the OCTA head and chin rest with an alcohol swab to disinfect the setup.</li>
	<li>Free the mouthpiece (#9) from its sterile packaging. 
	NOTE: Refrain from touching the mouthpiece as much as possible as this component makes direct contact with the mucus lining of the mouth of the subject</li>
	<li>Connect the mouthpiece (#9) to gas delivery tubing (#8)</li>
	<li>Place a pulse oximeter on the subjects&#8217; finger and begin monitoring oxygen saturation levels and pulse. 
	NOTE: Once the subject begins breathing the desired air mixture, the pulse oximeter should be continuously monitored by the examiner. If the oxygen saturation of the subject drops below 94%, the experiment should be stopped, as a safety precaution, and the subject observed until they return to baseline.</li>
	<li>Adjust the height of the OCTA setup so that the subject can easily rest their chin on the chinrest (#11) without overextending or flexing their neck.</li>
	<li>Loop the gas delivery tubing (#8) with mouthpiece (#9) attachment through the head and chin rest with the mouthpiece (#9) facing the patient. Have the tubing loop through the machine oppposite the side of the eye that the subject is having imaged.</li>
	<li>Insert the mouthpiece into the patient&#8217;s mouth. Encourage the subject to practice breathing through the non-rebreathing setup to create familiarity with the apparatus. Ensure the subject takes deep breathes to facilitate gas exchange.</li>
	<li>Place the nose clip (#10) on the subject to ensure they are breathing through the mouthpiece.</li>
	<li>Keep the three-way valve on Configuration 2 or change it to Configuration 1 depending on whether images are being acquired for exposure to room air or a specific gas mixture, respectively. For future reference, note the time as the start of gas inhalation.</li>
	<li>Have the subject place their chin on the right or left section of the chinrest (#11) according to the eye selected for imaging.</li>
	<li>Ensure they move their head forward until their forehead is in firm contact with the headrest (#11).</li>
	<li>Capture the OCTA scan of interest as determined by the testing protocol. In this study, three 3 mm x 3 mm images centered on the fovea were captured after 1 min of gas breathing.
	<ol>
		<li>Have the subject keep their head facing forward and still while fixating on the target in the center of their view</li>
		<li>In the live image seen in the iris view, center the scan.</li>
		<li>Bring the iris into focus by moving the chinrest in or out using the left-right arrows.</li>
		<li>Make sure the foveal dip is centered in the OCT scan, which should occur by default.</li>
		<li>Take an image. Scanning will usually last several seconds on an OCTA machine.</li>
		<li>View the OCTA image after the completion of the scan and ensure it is of adequate quality. Signal strength should be a 7 or better on a 10-point scale provided by the OCTA manufacturer.</li>
		<li>Select save or rescan the eye.</li>
		<li>Repeat steps 3.14.1&#8211;3.14.7 for as many scans are desired.</li>
		<li>Allow the subject to sit back from the machine. Remove the nose clip (#10) and the mouthpiece (#9) when no more scans of the eye with this gas mixture are needed.</li>
	</ol>
	</li>
	<li>Allow subjects a 2 min break before starting CO2 gas provocation experiments.</li>
	<li>Fill the Douglas bag with the first desired air mixture (consisting of 5% CO2, 21% oxygen and 74% nitrogen) as specified in step 1.2. The three-way valve will be in Configuration 2 after this step.</li>
	<li>Complete gas non-rebreathing apparatus setup by connecting the Air Control Unit to the Non-rebreathing Unit as shown in Figure 1 and described in step 1.1.4. Make sure all joints are airtight with sealing tape.</li>
	<li>Repeat steps 3.9&#8211;3.14, but now set the three-way valve to Configuration 1 when directed in step 3.11.</li>
	<li>Give subjects a 10 min break after the CO2 gas provocation to allow a return to baseline.</li>
	<li>While the subject is on break, fill the Douglas bag with 100% O2 according to step 1.2.</li>
	<li>Repeat steps 3.17&#8211;3.18 to perform the experiment under 100% O2 gas provocation conditions.</li>
</ol>
4. Experimental Clean Up
<ol>
	<li>Discard the disposable elements of the setup: the subject&#8217;s mouthpiece (#9) and nose clip (#10).</li>
	<li>Clean the head and chin rest (#11) using an alcohol swab. Wipe the subject chair, OCTA table and OCTA handles with a disinfectant wipe to remove any errant saliva.</li>
	<li>Disconnect the setup into its base components&#8212;the Air Control Unit and Non-rebreathing Unit&#8212;at the three-way valve (#3).</li>
	<li>As no air exhaled from the subject should have reached the elements of the Air Control Unit, empty the Douglas bag according to step 1.2.2 and place in a location for future retrieval. Disconnect the clean-bor tube (#2) with adapter (#2*) and three-way valve (#3) from the Douglas bag if desired for easier storage. This completes the Air Control Unit clean up.</li>
	<li>Remove the gas delivery tubing (#8) from the Non-rebreathing Unit by disconnecting it from the elbow joint (#7). Disconnect the in-house rubber tubing (#5) and tubing adapters (#4), from the two-way non-rebreathing valve (#6). Then do the same from the elbow joint (#7) by removing the sealing tape and detaching the parts by pulling them apart. 
	NOTE: More extensive cleaning of the two-way non-rebreathing valve may be facilitated by disassembling it to remove the internal diaphragms for additional care.</li>
	<li>Prepare a disinfectant bath for cleanup of the reusable components
	<ol>
		<li>Fill a container large enough to submerge the gas delivery tubing (#8) with an appropriately diluted and well mixed detergent disinfectant. In this case, dilute the detergent with water to a ratio of 1:6425.</li>
	</ol>
	</li>
	<li>Soak the gas delivery tubing (#8), two-way non-rebreathing valve (#6), elbow joint (#7), in-house rubber tubing (#5) and tubing adapters (#4) in the prepared disinfectant bath for at least 10 min.</li>
	<li>Remove all parts after the bath is over and rinse them thoroughly with water.</li>
	<li>Place them on a paper towel on a clean countertop to be air-dried.</li>
	<li>Once air drying has completed, dispose of the paper towel and place all components away for storage.</li>
</ol>
5. OCTA Data Export and Analysis
<ol>
	<li>OCTA data export
	<ol>
		<li>Export OCTA data by inserting a removable media device of choice into the OCTA computer. Find the subject and scan of interest.</li>
		<li>Select Export to create a zip folder containing the subject of interest&#8217;s data in a .bmp format on the removable media device.</li>
	</ol>
	</li>
	<li>OCTA data analysis
	<ol>
		<li>Organize the OCTA data on a laboratory computer with the ability to perform additional image analysis and processing.</li>
		<li>Use a custom script to suppress noise with a global thresholding technique and perform additional feature extraction. Binarize and skeletonize the OCTA images.</li>
		<li>On the post-processed images, calculate the vessel skeleton density (VSD)19,26, a dimensionless measure of the total linear length of vessels in an image calculated by the following equation performed on a binarized skeletonized image of the OCTA: 
		<img alt="Equation 1" src="/files/ftp_upload/60948/60948equ01.jpg" /> 
		where i and j refer to pixel coordinate (i,j), L(i,j) refers to white pixels representing decorrelation, X(i,j) refers to all pixels, and n refers to the dimensions of the pixel array, which can be assumed to be n x n pixels19,26. The denominator of this equation represents the total number of pixels which is calculated as written from the skeletonized image, but can be thought of as representing the physical area of the entire image.</li>
	</ol>
	</li>
</ol>

Carl Zeiss Meditec has provided grant funding, equipment and financial support to AHK related to the topic of this article.

The methodology just described is the complete protocol for a gas breathing provocation experiment that allows for the measurement of a subject&#8217;s RVR in a controlled environment at specific timepoints with no modifications to the OCTA imaging device and minimal discomfort or risk to the subject. This setup is described in a way that allows for easy modifications to fit the needs of the researcher. It can accommodate additional tubing to fit different clinic rooms and certain elements such as the in-house tubing or ...

The metabolic demand of the retina is dependent on an adequate and constant supply of oxygen provided by a well-regulated system of arterioles, capillaries and venules1. Several studies have demonstrated that the function of larger caliber human retinal vessels can be assessed in vivo with various physiologic2,3,4,5 and pharmacologic6,7 stimuli. In addition, abnormal function of this vascular system is common in retinal vascular diseases such as diabetic retinopathy where retinal vascular reactivity (RVR) has been shown to be attenuated even in its earliest stages8,9 through both gas provocation9 and flickering light experiments5,10,11. Retinal vascular risk factors such as smoking have also been correlated with impaired RVR12 and retinal blood flow13. These findings are important since the clinical symptoms of retinal vascular disease occur relatively late in the disease process and proven early clinical markers of disease are lacking14. Thus, assessing RVR can provide useful measures of vascular integrity for the early assessment of abnormalities that can initiate or exacerbate retinal degenerative diseases.
Previous RVR experiments have usually relied upon devices such as a laser blood flowmeter9 or fundus cameras equipped with special filters15 for retinal image acquisition. However, these technologies are optimized for larger diameter vessels such as arterioles16 and venules15, which are not where gas, micronutrient and molecular exchange occur. A more recent study was able to quantify the RVR of capillaries using adaptive optics imaging17, but despite the improved spatial resolution, these images have a smaller field size and are not FDA approved for clinical use18.
The recent advent of optical coherence tomography angiography (OCTA) has provided an FDA approved, noninvasive and dyeless angiographic method of assessing capillary level changes in human patients and subjects in vivo. OCTA is widely accepted in clinical practice as an effective tool for assessing impairment in capillary perfusion in retinal vascular diseases such as diabetic retinopathy19, retinal venous occlusions20, vasculitis21 and many others22. OCTA therefore provides an excellent opportunity for the evaluation of capillary level changes, which can have significant spatial and temporal heterogeneity23 as well as pathologic changes, in a clinical setting. Our group recently demonstrated that OCTA can be used to quantify the responsiveness of retinal vessels at the capillary level2 to physiologic changes in inspired oxygen, which is a retinal vasoconstrictive stimulus16,24, and carbon dioxide, which is a retinal vasodilatory stimulus3,5.
The goal of this article is to describe a protocol that will allow the reader to assess the retinal vascular reactivity of the smaller arterioles and capillary bed using OCTA. The methods are adapted from those presented in Lu et al.25 who described the measurement of cerebrovascular reactivity with magnetic resonance imaging. Although the present methods were developed and used during OCTA imaging2, they are applicable to other retinal imaging devices with relatively simple and obvious modifications.

<table>
	<tbody>
		<tr>
			<td>5% CO2 gas [5% CO2, 21% O2, 74% N2] (Compressed)</td>
			<td>Institution Dependent (Praxair)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bacdown Disinfectant Detergent</td>
			<td>Decon Labs</td>
			<td>8001</td>
			<td>https://deconlabs.com/products/disinfectant-bdd/</td>
		</tr>
		<tr>
			<td>Clean-Bor Tubes (35 mm Inner Diameter)</td>
			<td>Vacumed</td>
			<td>1011-108</td>
			<td>http://www.vacumed.com/zcom/product/Product.do?compid=27&#38;skuid=1197</td>
		</tr>
		<tr>
			<td>Cuff adapter for Douglas bag filling</td>
			<td>Vacumed</td>
			<td>22254</td>
			<td>http://www.vacumed.com/zcom/product/Product.do?compid=27&#38;prodid=343</td>
		</tr>
		<tr>
			<td>Douglas bag (200-liter capacity)</td>
			<td>Harvard Apparatus</td>
			<td>500942</td>
			<td>https://www.harvardapparatus.com/douglas-bag.html</td>
		</tr>
		<tr>
			<td>Elbow Joint (Inner Diameter 19mm/ Outer Diameter 22 mm), Modified in House</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fingertip Pulse Oximeter (Pro-Series)</td>
			<td>CMS</td>
			<td>CMS 500DL</td>
			<td>https://www.walmart.com/ip/Pro-Series-CMS-500DL-Fingertip-Pulse-Oximeter-Blood-Oxygen-Saturation-Monitor-with-silicon-cover-batteries-and-lanyard/479049154</td>
		</tr>
		<tr>
			<td>Gas Delivery Tube (22 mm Inner Diameter) Modified in House</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gas filling tube (1/8&#34; for compressed gas)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrogen Peroxide Cleaner Disinfectant Wipes</td>
			<td>Clorox Healthcare</td>
			<td>30824</td>
			<td>https://www.cloroxpro.com/products/clorox-healthcare/hydrogen-peroxide-cleaner-disinfectants/?gclid=EAIaIQobChMIk-KG4vi15QIVcRh9Ch0NNwLPEAAYASAAEgJIa_D_BwE&#38;gclsrc=aw.ds</td>
		</tr>
		<tr>
			<td>Lubricant Eye Drops</td>
			<td>Refresh</td>
			<td>Refresh Plus</td>
			<td>https://www.refreshbrand.com/Products/refresh-plus</td>
		</tr>
		<tr>
			<td>Manual Directional Control Valves: Three-Way T-Shape Stopcock Type (Inner Diameter 28.6 mm, Outer Diameter 35 mm)</td>
			<td>Hans Rudolph</td>
			<td>2100C Series</td>
			<td>www.rudolphkc.com</td>
		</tr>
		<tr>
			<td>Medical O2 (Compressed)</td>
			<td>Institution Dependent</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mouth piece (Silicone, Model #9061)</td>
			<td>Hans Rudolph</td>
			<td>602076</td>
			<td>www.rudolphkc.com</td>
		</tr>
		<tr>
			<td>OCTA Imaging Device (PLEX Elite 9000)</td>
			<td>Carl Zeiss Meditec, Dublin, CA, USA</td>
			<td></td>
			<td>https://www.zeiss.com/meditec/int/product-portfolio/optical-coherence-tomography-devices/plex-elite-9000-swept-source-oct.html</td>
		</tr>
		<tr>
			<td>Phenylephrine Hydrochloride Ophthalmic Solution, USP 2.5%</td>
			<td>Paragon Bioteck, Inc</td>
			<td>NDC 42702-102-15</td>
			<td>https://paragonbioteck.com/products/diagnostics/phenylephrine-hydrochloride-ophthalmic-solution-usp-2-5/</td>
		</tr>
		<tr>
			<td>Plastic Nose Clip Sterile Foam CS100</td>
			<td>Sklar Sterile</td>
			<td>96-2951</td>
			<td>https://www.sklarcorp.com/disposables/plastic/plastic-nose-clip-sterile-foam-box-of-100.html</td>
		</tr>
		<tr>
			<td>Proparacaine Hydrochloride Ophthalmic Solution, USP .5%</td>
			<td>Bausch + Lomb</td>
			<td>NDC 24208-730-06</td>
			<td>https://www.bausch.com/ecp/our-products/rx-pharmaceuticals/generics</td>
		</tr>
		<tr>
			<td>Regulator (tank dependent- 5% CO2: Fisherbrand Mulitstage Gas Cylinder Regulators)</td>
			<td>Genstar Technologies Company</td>
			<td>10575150</td>
			<td>https://www.fishersci.com/shop/products/fisherbrand-multistage-cylinder-regulators-22/10575150?keyword=true</td>
		</tr>
		<tr>
			<td>Regulator (tank dependent- Oxygen: Fisherbrand Multistage Gas Cylinder Regulators)</td>
			<td>Genstar Technologies Company</td>
			<td>10575145</td>
			<td>https://www.fishersci.com/shop/products/fisherbrand-multistage-cylinder-regulators-22/10575145?keyword=true</td>
		</tr>
		<tr>
			<td>Rubber Tubing (Inner diameter 19 mm, Outer diameter 27 mm), Made in House</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sealing tape- Parafilm Wrap (2&#34; Wide)</td>
			<td>Cole Parmer</td>
			<td>PM992</td>
			<td>https://www.coleparmer.com/i/parafilm-pm992-wrap-2-wide-250-ft-roll/0672050?PubID=VV&#38;persist=True&#38;ip=no&#38;gclid=EAIaIQobChMInY3vqomz5QIVfyCtBh1VSg64EAAYASAAEgJ9n_D_BwE</td>
		</tr>
		<tr>
			<td>Sterile Alcohol Prep Pads</td>
			<td>Medline</td>
			<td>MDS090670</td>
			<td>https://www.medline.com/product/Sterile-Alcohol-Prep-Pads/Swab-Pads/Z05-PF03816</td>
		</tr>
		<tr>
			<td>Tropicamide Ophthalmic Solution, USP 1%</td>
			<td>Akorn</td>
			<td>NDC 17478-102-12</td>
			<td>http://www.akorn.com/prod_detail.php?ndc=17478-102-12</td>
		</tr>
		<tr>
			<td>Tubing Adapter, Made in House</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Two-way non-rebreathing valve (2600 Series- Inner Diameter 28.6 mm, Outer Diameter 35 mm)</td>
			<td>Hans Rudolph</td>
			<td>2600 Series, UM-112078</td>
			<td>www.rudolphkc.com</td>
		</tr>
	</tbody>
</table>

retinal vascular reactivity as assessed by optical coherence tomography angiography

The vascular supply to the retina has been shown to dynamically adapt through vasoconstriction and vasodilation to accommodate the metabolic demands of the retina. This process, referred to as retinal vascular reactivity (RVR), is mediated by neurovascular coupling, which is impaired very early in retinal vascular diseases such as diabetic retinopathy. Therefore, a clinically feasible method of assessing vascular function may be of significant interest in both research and clinical settings. Recently, in vivo imaging of the retinal vasculature at the capillary level has been made possible by the FDA approval of optical coherence tomography angiography (OCTA), a noninvasive, minimal risk and dyeless angiography method with capillary level resolution. Concurrently, physiological and pathological changes in RVR have been shown by several investigators. The method shown in this manuscript is designed to investigate RVR using OCTA with no need for alterations to the clinical imaging procedures or device. It demonstrates real time imaging of the retina and retinal vasculature during exposure to hypercapnic or hyperoxic conditions. The exam is easily performed with two personnel in under 30 min with minimal subject discomfort or risk. This method is adaptable to other ophthalmic imaging devices and the applications may vary based on the composition of the gas mixture and patient population. A strength of this method is that it allows for an investigation of retinal vascular function at the capillary level in human subjects in vivo. Limitations of this method are largely those of OCTA and other retinal imaging methods including imaging artifacts and a restricted dynamic range. The results obtained from the method are OCT and OCTA images of the retina. These images are amenable to any analysis that is possible on commercially available OCT or OCTA devices. The general method, however, can be adapted to any form of ophthalmic imaging.

The output from this experiment consists of the manual readings taken from the pulse oximeter, the timing noted for gas exposure or OCTA scanning and the raw OCTA imaging data. An OCTA image consists of the OCT B-scans and the decorrelation signal associated with each B-scan. The data parameters are given by the specifications of the device. A swept source laser platform OCTA machine with a central wavelength of 1040&#8211;1060 nm was used. The images provide a transverse resolution of 20 &#181;m and optical axial resolu...

Watch this Scientific Journal Video about Retinal Vascular Reactivity as Assessed by Optical Coherence Tomography Angiography at JoVE.com

Retinal Vascular Reactivity as Assessed by Optical Coherence Tomography Angiography

This article describes a method for measuring retinal vasculature reactivity in vivo with human subjects using a gas breathing provocation technique to deliver vasoactive stimuli while acquiring retinal images.

The vascular supply to the retina has been shown to dynamically adapt through vasoconstriction and vasodilation to accommodate the metabolic demands of ...

retinal-vascular-reactivity-as-assessed-optical-coherence-tomography

Research

JoVE Journal

Neuroscience

7.0K Views.  USC Roski Eye Institute. This article describes a method for measuring retinal vasculature reactivity in vivo with human subjects using a gas breathing provocation technique to deliver vasoactive stimuli while acquiring retinal images.

Video: Retinal Vascular Reactivity as Assessed by Optical Coherence Tomography Angiography

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1. This information is then segregated into what are known as the ON and OFF pathways. The photoreceptors signal light information to the bipolar cells (BCs), which depolarize in response to increases (On BCs) or decreases (Off BCs) in light intensity. This segregation of light information is maintained at the level of the retinal ganglion cells (RGCs), which have dendrites stratifying in either the Off sublamina of the inner plexiform layer (IPL), where they receive direct excitatory input from Off BCs, or stratifying in the On sublamina of the IPL, where they receive direct excitatory input from On BCs. This segregation of information regarding increases or decreases in illumination (the On and Off pathways) is conserved and signaled to the brain in parallel.
The RGCs are the output cells of the retina, and are thus an important cell to study in order to understand how light information is signaled to visual nuclei in the brain. Advances in mouse genetics over recent decades have resulted in a variety of fluorescent reporter mouse lines where specific RGC populations are labeled with a fluorescent protein to allow for identification of RGC subtypes 2 3 4 and specific targeting for electrophysiological recording. Here, we present a method for recording light responses from fluorescently labeled ganglion cells in an intact, isolated retinal preparation. This isolated retinal preparation allows for recordings from RGCs where the dendritic arbor is intact and the inputs across the entire RGC dendritic arbor are preserved. This method is applicable across a variety of ganglion cell subtypes and is amenable to a wide variety of single-cell physiological techniques.

This article provides a description of how to dissect and record from the isolated retinal preparation in mouse. In particular, we describe how to record light responses from a fluorescently labeled ganglion cell population and subsequently identify and analyze its morphology.

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1.  This information is then ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

Optical coherence tomography (OCT) is a biomedical imaging technique with high spatial-temporal resolution. With its minimally invasive approach OCT has been used extensively in ophthalmology, dermatology, and gastroenterology1-3. Using a thinned-skull cortical window (TSCW), we employ spectral-domain OCT (SD-OCT) modality as a tool to image the cortex in vivo. Commonly, an opened-skull has been used for neuro-imaging as it provides more versatility, however, a TSCW approach is less invasive and is an effective mean for long term imaging in neuropathology studies. Here, we present a method of creating a TSCW in a mouse model for in vivo OCT imaging of the cerebral cortex.

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

We present a method of creating a thinned-skull cortical window (TSCW) in a mouse model for in vivo OCT imaging of the cerebral cortex.

Optical coherence tomography (OCT) is a biomedical imaging technique  with high spatial-temporal resolution. With its minimally invasive approach OCT  has ...

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

A Protocol for Measuring Cue Reactivity in a Rat Model of Cocaine Use Disorder

Cocaine use disorder (CUD) follows a trajectory of repetitive self-administration during which previously neutral stimuli gain incentive value. Cue reactivity, the sensitivity to cues previously linked with the drug-taking experience, plays a prominent role in human craving during abstinence. Cue reactivity can be assessed as the attentional orientation toward drug-associated cues that is measurable as appetitive approach behavior in both preclinical and human studies. Herein describes an assessment of cue reactivity in rats trained to self-administer cocaine. Cocaine self-administration is paired with the presentation of discrete cues that act as conditioned reinforcers (i.e., house light, stimulus light, infusion pump sounds). Following a period of abstinence, lever presses in the cocaine self-administration context accompanied by the discrete cues previously paired with cocaine infusion are measured as cue reactivity. This model is useful to explore neurobiological mechanisms underlying cue reactivity processes as well as to assess pharmacotherapies to suppress cue reactivity and therefore, modify relapse vulnerability. Advantages of the model include its translational relevance, and its face and predictive validities.&#160;The primary limitation of the model is that the cue reactivity task can only be performed infrequently and must only be used in short duration (e.g., 1 hour), otherwise rats will begin to extinguish the pairing of the discrete cues with the cocaine stimulus. The model is extendable to any positively reinforcing stimulus paired with discrete cues; though particularly applicable to drugs of abuse, this model may hold future applications in fields such as obesity, where palatable food rewards can act as positively reinforcing stimuli.

Cue reactivity is conceptualized as sensitivity to cues linked with drug-taking experiences that contribute to craving and relapse in abstinent humans. Cue reactivity is modeled in rats by measuring attentional orientation toward drug-associated cues that results in appetitive approach behavior in a cue reactivity test following self-administration and forced abstinence.

Cocaine use disorder (CUD) follows a trajectory of repetitive self-administration during which previously neutral stimuli gain incentive value. Cue ...

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in vivo&#8212;rapidly, repetitively, and at a high resolution. This protocol describes OCT imaging in the mouse retina as a powerful tool to study optic neuropathies (OPN). The OCT system is an interferometry-based, non-invasive alternative to common post mortem histological assays. It provides a fast and accurate assessment of retinal thickness, allowing the possibility to track changes, such as retinal thinning or thickening. We present the imaging process and analysis with the example of the Opa1delTTAG mouse line. Three types of scans are proposed, with two quantification methods: standard and homemade calipers. The latter is best for use on the peripapillary retina during radial scans; being more precise, is preferable for analyzing thinner structures. All approaches described here are designed for retinal ganglion cells (RGC) but are easily adaptable to other cell populations. In conclusion, OCT is efficient in mouse model phenotyping and has the potential to be used for the reliable evaluation of therapeutic interventions.

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

This manuscript describes a protocol for the in vivo imaging of the mouse retina with high-resolution spectral domain optical coherence tomography (SD-OCT). It focuses on retinal ganglion cells (RGC) in the peripapillary region, with several scanning and quantifying approaches described.

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in ...

Using Optical Coherence Tomography and Optokinetic Response As Structural and Functional Visual System Readouts in Mice and Rats

Optical coherence tomography (OCT) is a fast, non-invasive, interferometric technique allowing high-resolution retinal imaging. It is an ideal tool for the investigation of processes of neurodegeneration, neuroprotection and neuro-repair involving the visual system, as these often correlate well with retinal changes. As a functional readout, visually evoked compensatory eye and head movements are commonly used in experimental models involving the visual function. Combining both techniques allows a quantitative in vivo investigation of structure and function, which can be used to investigate the pathological conditions or to evaluate the potential of novel therapeutics. A great benefit of the presented techniques is the possibility to perform longitudinal analyses allowing the investigation of dynamic processes, reducing variability and cuts down the number of animals needed for the experiments. The protocol described aims to provide a manual for acquisition and analysis of high quality retinal scans of mice and rats using a low cost customized holder with an option to deliver inhalational anesthesia. Additionally, the proposed guide is intended as an instructional manual for researchers using optokinetic response (OKR) analysis in rodents, which can be adapted to their specific needs and interests.

A detailed protocol for the assessment of structural and visual readouts in rodents by optical coherence tomography and optokinetic response is presented. The results provide valuable insights for ophthalmologic as well as neurologic research.

Optical coherence tomography (OCT) is a fast, non-invasive, interferometric technique allowing high-resolution retinal imaging. It is an ideal tool for ...

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for determining the mechanisms underlying retinal diseases. Maintaining structural integrity throughout the tissue preparation is vital for obtaining reproducible retinal IHC data but can be challenging due to the fragility and complexity of retinal cytoarchitecture. Strong fixatives like 10% formalin or Bouin's solution optimally preserve the retinal structure, they often impede IHC analysis by enhancing the background fluorescence and/or diminishing antibody-epitope interactions, a process known as epitope masking. Milder fixatives, on the other hand, like 4% paraformaldehyde, reduces background fluorescence and epitope-masking, meticulous dissection techniques must be utilized to preserve the retinal structure. In this article, we present a comprehensive method to prepare mouse ocular posterior cups for IHC that is sufficient to preserve most antibody-epitope interactions without loss of retinal structural integrity. We include representative IHC with antibodies to various retinal cell type markers to illustrate tissue preservation and orientation under optimal and sub-optimal conditions. Our goal is to optimize IHC studies of the retina by providing a complete protocol from ocular posterior cup dissection to IHC.

This report describes comprehensive methods for preparing frozen mouse retina sections for immunohistochemistry (IHC). Methods described include dissection of the ocular posterior cup, paraformaldehyde fixation, embedding in Optimal Cutting Temperature (OCT) media and tissue orientation, sectioning and immunostaining.

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for ...

Coculture of Axotomized Rat Retinal Ganglion Neurons with Olfactory Ensheathing Glia, as an In Vitro Model of Adult Axonal Regeneration

Olfactory ensheathing glia (OEG) cells are localized all the way from the olfactory mucosa to and into the olfactory nerve layer (ONL) of the olfactory bulb. Throughout adult life, they are key for axonal growing of newly generated olfactory neurons, from the lamina propria to the ONL. Due to their pro-regenerative properties, these cells have been used to foster axonal regeneration in spinal cord or optic nerve injury models.
We present an in vitro model to assay and measure OEG neuroregenerative capacity after neural injury. In this model, reversibly immortalized human OEG (ihOEG) is cultured as a monolayer, retinas are extracted from adult rats and retinal ganglion neurons (RGN) are cocultured onto the OEG monolayer. After 96 h, axonal and somatodendritic markers in RGNs are analyzed by immunofluorescence and the number of RGNs with axon and the mean axonal length/neuron are quantified.
This protocol has the advantage over other in vitro assays that rely on embryonic or postnatal neurons, that it evaluates OEG neuroregenerative properties in adult tissue. Also, it is not only useful for assessing the neuroregenerative potential of ihOEG but can be extended to different sources of OEG or other glial cells.

We present an in vitro model to assess olfactory ensheathing glia (OEG) neuroregenerative capacity, after neural injury. It is based on a coculture of axotomized adult retinal ganglion neurons (RGN) on OEG monolayers and subsequent study of axonal regeneration, by analyzing RGN axonal and somatodendritic markers.

Olfactory ensheathing glia (OEG) cells are localized all the way from the olfactory mucosa to and into the olfactory nerve layer (ONL) of the olfactory ...

Quantification of Vascular Parameters in Whole Mount Retinas of Mice with Non-Proliferative and Proliferative Retinopathies

Retinopathies are a heterogeneous group of diseases that affect the neurosensory tissue of the eye. They are characterized by neurodegeneration, gliosis&#160;and a progressive change in vascular function and structure. Although the onset of the retinopathies is characterized by subtle disturbances in visual perception, the modifications in the vascular plexus are the first signs detected by clinicians. The absence or presence of neovascularization determines whether the retinopathy is classified as either non-proliferative (NPDR) or proliferative (PDR). In this sense, several animal models tried to mimic specific vascular features of each stage to determine the underlying mechanisms involved in endothelium alterations, neuronal death and other events taking place in the retina. In this article, we will provide a complete description of the procedures required for the measurement of retinal vascular parameters in adults and early birth mice at postnatal day (P)17. We will detail the protocols to carry out retinal vascular staining with Isolectin GSA-IB4 in whole mounts for later microscopic visualization. Key steps for image processing with Image J Fiji software are also provided, therefore, the readers will be able to measure vessel density, diameter and tortuosity, vascular branching, as well as avascular and neovascular areas. These tools are highly helpful to evaluate and quantify vascular alterations in both non-proliferative and proliferative retinopathies.

This article describes a well-established and reproducible lectin stain assay for the whole mount retinal preparations and the protocols required for the quantitative measurement of vascular parameters frequently altered in proliferative and non-proliferative retinopathies.