Funding for this project was through discretionary funds of Dr. Colleen M. McDowell. This work was supported in part by an unrestricted grant from Research to Prevent Blindness, Inc. to the UW Madison Department of Ophthalmology and Visual Sciences. We thank Drs. Abbot F. Clark and Weiming Mao for their technical assistance with the perfusion organ culture model. We thank the Lions Eye Institute for Transplant and Research (Tampa, FL) for providing the human donor eyes.

Eyes were obtained according to the provisions of the Declaration of Helsinki for research involving human tissue.
NOTE: Eyes from reputable eye banks (e.g., Lions Eye Institute for Transplant, Research, Tampa FL) were harvested within 6&#8211;12 h of death and donor serum was tested for hepatitis B, hepatitis C, and human immunodeficiency virus 1 and 2. Once they were received, the eyes were dissected and set up in the TAS model within 24 h. Exclusion criteria included any ocular pathology. Eyes were not excluded based on age, race, or gender. To ensure the viability of the retina upon receipt, retinal explants were harvested from the tissue donors and cultured for 7 and 14 days (Supplemental Figure 1). These retinas were also dissociated and grew healthy RGCs in culture for 7 days with positive staining for RGC marker, RNA-binding protein with multiple splicing (RBPMS), as well as positive neurofilament light chain (NEFL) staining in their neurofilaments (Supplemental Figure 2).&#160;.
1. Preparation and sterilization of equipment and supplies
<ol>
	<li>Refer to the Table of Materials for a complete list of supplies required as well as supplier and catalogue numbers.</li>
	<li>Prior to use, sterilize all equipment and instruments by autoclaving or using ethylene oxide ampules.</li>
</ol>
2. Preparation of perfusion medium
<ol>
	<li>Add 1% penicillin streptomycin (10,000 U/mL penicillin, 10,000 &#956;g/mL streptomycin in 0.85% NaCl) and 1% L-glutamine (200 mM) to 1,000 mL high glucose Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM).</li>
	<li>Sterilize the perfusion medium by passing through a 0.22 &#956;m filter.</li>
</ol>
3. Translaminar autonomous system (TAS) setup
<ol>
	<li>Set up inflow syringes (IOP and ICP reservoirs).
	<ol>
		<li>Add 30 mL of the perfusion medium (section 2) to a 30 mL syringe. Attach a 3-way stopcock to the 30 mL syringe. Attach a 0.22 &#956;m hydrophilic filter to the 3-way stopcock. Attach a 15 G Luer stub adapter to the 0.22 &#956;m hydrophilic filter.</li>
		<li>Remove air bubbles from the syringe setup. Attach tubing to the 15 G Luer stub adapter. Close the side port of the stopcock with an unvented universal lock cap. Repeat for a total of two setups.</li>
		<li>Label one syringe as channel 1 intracranial pressure (CH1 ICP) and the other syringe as channel 2 intraocular pressure (CH2 IOP).</li>
	</ol>
	</li>
	<li>Set up outflow syringes (IOP and ICP reservoirs).
	<ol>
		<li>Attach a 3-way stopcock to a 30 mL syringe. Attach a 15 G Luer stub adapter to the 3-way stopcock. Attach tubing to the 15 G Luer stub adapter.</li>
		<li>Close the side port of the stopcock with an unvented universal lock cap. Repeat for a total of two setups. Label one syringe as CH1 ICP and the other syringe as CH2 IOP.</li>
	</ol>
	</li>
</ol>
4. Preparation of human whole eye globe
NOTE: If whole eyes are received, follow the procedure below to separate the anterior segment from the posterior segment of the eye. If the eyes are received bisected, start at step 4.4.
<ol>
	<li>Place a whole eye into povidone-iodine solution for 2 min.</li>
	<li>Rinse the eye in sterile phosphate buffered solution (PBS) to rinse off the povidone-iodine. Repeat 2 times.</li>
	<li>Remove the adnexa from the whole eye globe using&#160;forceps and scissors. Bisect the eye at the equator to separate the anterior and posterior segments of the eye.</li>
	<li>Remove the optic nerve sheath. Remove the vitreous humor from the posterior segment.</li>
	<li>Trim additional sclera from posterior segment, if needed, to ensure a good fit on the round dome of the IOP (bottom) chamber. Using forceps, ensure that the retina is spread evenly over the posterior of the segment.</li>
	<li>IOP (bottom) chamber setup
	<ol>
		<li>Place the human posterior segment into the IOP (bottom) chamber of the TAS over the round dome with the optic nerve facing the top.</li>
		<li>Seal the posterior segment using the epoxy resin O-ring with four screws, ensuring a tight seal.</li>
		<li>Insert the tubing into the IN and OUT ports of the IOP (bottom) chamber. The IOP inflow syringe with tubing containing medium is inserted into the IN port and the empty IOP outflow syringe with tubing is inserted into the OUT port.</li>
		<li>Use the push/pull method to slowly infuse the perfusion medium into the inflow port to fill the posterior eye cup while simultaneously slowly pulling the perfusion medium out through the outflow syringe to remove any air bubbles from the lines. Stop infusing medium once both the IN and OUT tubes are void of air bubbles.</li>
		<li>Lock the stopcocks in the off position. Remove the 30 mL syringe from the IOP IN port filter assembly and refill with a total of 30 mL of medium. Replace the 30 mL syringe onto the filter assembly.</li>
	</ol>
	</li>
	<li>ICP (top) chamber setup
	<ol>
		<li>Place the ICP (top) chamber/lid over the back of the posterior segment. Make certain that the optic nerve is within the top chamber. Seal the top chamber with four screws.</li>
		<li>Insert the tubing into the IN and OUT ports of the ICP (top) chamber. The ICP inflow syringe with tubing containing medium is inserted into the IN port and the empty ICP outflow syringe with tubing is inserted into the OUT port.</li>
		<li>Gently and slowly infuse the medium into the IN port to fill the ICP chamber and remove air bubbles from the lines using the push/pull method. Stop infusing medium once the ICP chamber as well as both the IN and OUT tubing are void of air bubbles.</li>
		<li>Lock the stopcocks in the off position. Remove the 30 mL syringe from the ICP in port filter assembly and refill with a total of 30 mL of medium. Replace the 30 mL syringe onto the filter assembly.</li>
	</ol>
	</li>
</ol>
5. Data recording system setup
NOTE: The data recording system is comprised of an 8-channel power source, multichannel bridge amplifier, hydrostatic pressure transducers, and a computer with data acquisition software (see Table of Materials). The following describes how to set up and calibrate the system.
<ol>
	<li>Connect the power cord into the back of the 8-channel power source and plug into a battery back-up device.</li>
	<li>Connect the USB cable from the 8-channel power source into the back of the computer.</li>
	<li>Connect the 8-channel power source to the multichannel bridge amplifier using the supplied I2C cord.</li>
	<li>Connect the Bayonet Neill-Concelman (BNC) cables into the channel inputs in front of the 8-channel power source and the end of the cables into the corresponding channels in the back of the multichannel amplifier.</li>
	<li>Connect the transducer cables to the front of the multichannel amplifier.</li>
	<li>Install the data acquisition software on the computer.
	<ol>
		<li>Run the data acquisition software setup installer from the supplied software CD.</li>
		<li>Follow the instructions on the computer screen.</li>
		<li>When the installation is complete, select Finish.</li>
	</ol>
	</li>
	<li>Turn on the 8-channel power source.</li>
	<li>Turn on the computer and initiate the data acquisition software.
	<ol>
		<li>Select File | New.</li>
		<li>Select Setup | Channel Settings. Select three channels (bottom left of screen). In the Channel Title column rename the channels as follows: CH1 ICP; CH2 IOP; CH3 TLPG (IOP-ICP).</li>
		<li>Select 2 mV for the Range on all channels. In the Calculation column select No Calculation for channels 1 and 2.</li>
		<li>In the Calculation column select Arithmetic for channel 3. In the Formula section: Select channels/CH2; Select arithmetic &#34;-&#34;; Select channels/CH1. In the Output section select mmHg. Select OK. Select OK again.</li>
	</ol>
	</li>
	<li>Set up and calibrate the hydrostatic pressure transducers. 
	NOTE: Hydrostatic pressure transducers must be calibrated prior to experiments using the following method.
	<ol>
		<li>Connect the hydrostatic pressure transducers to the transducer lines attached to the multichannel bridge amplifier.</li>
		<li>Attach a 30 mL syringe filled with air to the side port of the CH1 (ICP) pressure transducer. Attach the sphygmomanometer to the bottom of the CH1 (ICP) pressure transducer.</li>
		<li>On the chart, view the page of the data acquisition software, set the sampling speed by left clicking the arrow next to the sampling time and select 100. Then right click in the CH1 (ICP) area of the page.</li>
		<li>Select Bridge Amp. Select Mains Filter. Select Zero and wait for the system to zero out, taking care to not move the pressure transducer.</li>
		<li>Pinch the white tabs of the pressure transducer and push air through the transducer until 40 mmHg is obtained on the sphygmomanometer. Release the white tabs and remove the syringe and sphygmomanometer.</li>
		<li>On the Units Conversion page, select &#8216;minus (-)&#8217; sign. Highlight the highest plateau to indicate 40 mmHg. Click the Arrow for point 1 and enter 40.</li>
		<li>Highlight the lowest plateau to indicate 0 mmHg. Click the Arrow for point 2 and enter 0. Select mmHg for the units. Select OK.</li>
		<li>Select OK (Bridge Amp page). Repeat steps 9.1&#8211;9.7 for CH2 (IOP) using 100 mmHg for the highest plateau and 0 for the lowest plateau.</li>
	</ol>
	</li>
	<li>Connect the TAS/posterior segment unit onto the data acquisition system.
	<ol>
		<li>Place the TAS/posterior segment unit into an incubator (37 &#176;C, 5% CO2). Attach the ICP tubing from the OUT port to the CH1 (ICP) pressure transducer.</li>
		<li>Attach the IOP tubing from the OUT port to the CH2 (IOP) pressure transducer.</li>
		<li>Attach the syringe setups (ICP and IOP) with medium from the IN ports to the ring stand.</li>
		<li>On the Chart view page select Start Sampling. Set the sampling speed by left clicking the arrow next to the sampling time and select Slow and 1 min.</li>
		<li>Adjust the syringes on the ring stand up or down to regulate ICP and IOP pressures to protocol requirements.</li>
		<li>Perform systemic replenishment of medium in the system every 48&#8211;72 h through the push and pull method.</li>
	</ol>
	</li>
</ol>
6. Data retrieval and analysis
<ol>
	<li>Open the data file in the data acquisition software.</li>
	<li>In the Data Pad section, click on the Multiple Add to Data Pad icon. A new window will appear.
	<ol>
		<li>In the Find Using section select Time from the drop-down menu.</li>
		<li>In the Select section select 1 hr every 1 hr from the drop-down menus.</li>
		<li>In the Step Through section select Whole File then click Add.</li>
	</ol>
	</li>
	<li>In the Data Pad section click on the Data Pad View icon. Highlight all the data and copy/paste into a spreadsheet.</li>
	<li>Calculate the mean and standard deviations for IOP, ICP, and TLPG for every 24 h. Collate the data using the pivot table option in a spreadsheet program and graph.</li>
</ol>
7. Immunohistochemistry and hematoxylin and eosin staining of posterior segments
<ol>
	<li>Remove the posterior eye segments following various timepoints from the TAS model and fix in formalin prior to paraffinizing.</li>
	<li>Section the eyes to produce sagittal tissue planes.</li>
	<li>Deparaffinize the paraffin-embedded segments with a 100% xylene, 95% ethanol, 50% ethanol solution.</li>
	<li>Wash the slides with PBS for 10 min and block with a blocking buffer at room temperature for 1 h.</li>
	<li>Label sections with primary antibodies: anti-collagen IV (Extracellular Matrix (ECM) marker, NB120-6586, 1:100) and anti-laminin (ECM marker, NB300-144, 1:100, anti-RBPMS (RGC marker), GTX118619, 1:50).</li>
	<li>Detect the primary antibodies using Alexa Fluor secondary antibodies (Alexa Fluor 488 goat anti-rabbit, A11008, 1:500).</li>
	<li>Counterstain the cell nuclei using DAPI anti-fade solution.</li>
	<li>Capture images of the stained sections and phase images with 4x and 10x objective lenses using a fluorescence microscope (see Table of Materials).</li>
	<li>For the hematoxylin and eosin (H&#38;E) staining, process the sections in an automated staining system (see Table of Materials) for deparaffinization using a 100%, xylene 95% ethanol, 50% ethanol solution and stain with H&#38;E.</li>
	<li>Capture images with the 4x and 10x objective lenses using a microscope with a bright field light source.</li>
</ol>

The authors of the manuscript have no potential conflicts of interest to disclose.

Human postmortem tissues are an especially valuable resource for studying human neurodegenerative diseases because identification of potential drugs developed in&#160;animal models need to be translatable to humans47. The effects of human IOP elevation are well-established, but minimal research has been performed on abnormal ONH translaminar pressure changes. Even though multiple animal models and finite modeling of human ONH exist, there is no ex vivo human model to study translaminar pressure ch...

Global estimates in recent surveys suggest that 285 million people live with visual impairment, including 39 million who are blind1. In 2010, the World Health Organization documented that three of the nine listed leading causes of blindness occur in the posterior segment of the eye1. Posterior segment eye diseases involve the retina, choroid, and optic nerve2. The retina and optic nerve are central nervous system (CNS) extensions of the brain. The retinal ganglion cell (RGC) axons are vulnerable to damage because they exit the eye through the optic nerve head (ONH) to form the optic nerve3. The ONH remains the most vulnerable point for the RGC&#160;axons because of the 3D meshwork of connective tissue beams called the lamina cribrosa (LC)4. The ONH is the initial site of insult to RGC axons in glaucoma5,6,7, and gene expression changes within the ONH have been studied in ocular hypertension and glaucoma models8,9,10. The RGC axons are susceptible at the ONH due to pressure differentials between the intraocular compartment, called the intraocular pressure (IOP), and within the external perioptic subarachnoid space, called the intracranial pressure (ICP)11. The LC region separates both areas, maintaining normal pressure differentials, with IOP ranging from 10&#8211;21 mmHg and ICP from 5&#8211;15 mmHg12. The pressure difference through the lamina between the two chambers is called the translaminar pressure gradient (TLPG)13. A major risk factor of glaucoma is elevated IOP14.
Increased IOP increases the strain within and across the laminar region6,15,16. Experimental observations in humans and animal models present the ONH as being the initial site of axonal damage17,18. The biomechanical paradigm of IOP-related stress and strain causing glaucomatous damage at the ONH also influences the pathophysiology of glaucoma19,20,21. Even though in humans pressure-induced changes mechanically damage RGC axons22, rodents lacking collagenous plates within the lamina can also develop glaucoma7,23. In addition, elevated IOP remains the most prominent risk factor in primary open angle glaucoma patients, while normal tension glaucoma patients develop glaucomatous optic neuropathy even without elevated IOP. Furthermore, there are also a subset of ocular hypertensive patients that show no optic nerve damage. It has also been suggested that cerebrospinal fluid pressure (CSFp) may play a role in glaucoma pathogenesis. Evidence indicates that ICP is lowered to ~5 mmHg in glaucoma patients compared to normal individuals, thereby causing increased translaminar pressure and playing a crucial role in disease24,25. Previously, it was demonstrated in a canine model, that by controlling IOP and CSFp changes, there can be large displacements of the optic disc26. Elevating CSFp in porcine eyes has also shown increased principal strain within the LC region and retrolaminar neural tissue. Increased strain on the RGCs and the LC region contributes to axonal transport blockage and loss of RGCs27. Progressive degeneration of RGCs has been associated with loss of trophic support28,29, stimulation of inflammatory processes/immune regulation30,31, and apoptotic effectors29,32,33,34,35. Additionally, axonal injury (Figure 3) causes detrimental effects on the RGCs, triggering regenerative failure36,37,38,39. Even though the effects of IOP have been well studied, minimal research has been performed on abnormal translaminar pressure changes. Most treatments for glaucoma focus on stabilizing IOP. However, even though lowering of IOP slows the progression of the disease, it does not reverse visual field loss and prevent complete loss of RGCs. Understanding pressure-related neurodegenerative changes in glaucoma will be crucial to preventing RGC death.
Current evidence indicates that translaminar pressure modulations due to various mechanical, biological, or physiological changes in patients suffering from traumatic or neurodegenerative visual impairments can cause significant vision loss. Currently, no true preclinical human posterior segment model exists that can allow the study of glaucomatous biomechanical damage within the ex vivo human ONH. Observation and treatment of the posterior segment of the eye is a huge challenge in ophthalmology27. There are physical and biological barriers to target the posterior eye, including high elimination rates, blood-retinal barrier, and potential immunological responses40. Most efficacy and safety tests for novel drug targets are accomplished utilizing in vitro cellular and in vivo animal models41. Ocular anatomy is complex, and in vitro studies do not accurately mimic the anatomical and physiological barriers presented by tissue model systems. Even though animal models are a necessity for pharmacokinetic studies, the ocular physiology of the human posterior eye may vary between various animal species, including cellular anatomy of the retina, vasculature, and ONH41,42.
The use of living animals requires intensive and detailed ethical regulations, high financial commitment, and effective reproducibility43. Recently,&#160;multiple other guidelines have ensued for the ethical use of animals in experimental research44,45,46. An alternative to animal testing is the use of ex vivo human eye models to investigate disease pathogenesis and potential analysis of drugs for protecting ONH damage. Human postmortem tissue is a valuable resource for studying human disease paradigms, especially in the case of human neurodegenerative diseases, because identification of potential drugs developed in&#160;animal models require the need to be translatable to humans47. The ex vivo human donor tissue has been extensively utilized for the study of human disorders47,48,49, and human anterior segment perfusion organ culture systems have previously provided a unique ex vivo model to study the pathophysiology of elevated IOP50,51,52.
To study translaminar pressure related to IOP and ICP in human eyes, we successfully designed and developed a two-chamber translaminar autonomous system (TAS) that can independently regulate IOP and ICP using posterior segments from human donor eyes. It is the first ex vivo human model to study translaminar pressure and exploit the biomechanical effects of TLPG on the ONH.
This ex vivo human TAS model can be used to discover and classify cellular and functional modifications that occur due to chronic elevation of IOP or ICP. In this report, we detail the step-by-step protocol of dissecting, setting up, and monitoring the TAS human posterior segment model. The protocol will allow other researchers to effectively reproduce this novel ex vivo pressurized human posterior segment model to study biomechanical disease pathogenesis.

<table>
	<tbody>
		<tr>
			<td>#122, 1-1/8&#34; Inside x 1-5/16&#34; Outside Diam, Viton O-Ring, 3/32&#34; Thick, 
			755 Durometer 50 Pack</td>
			<td>Amazon</td>
			<td>B07DRGPPZJ</td>
			<td></td>
		</tr>
		<tr>
			<td>114 Buna-N O-Ring, 70A Durometer, Black, 5/8&#34; ID, 13/16&#34; OD, 3/32&#34; Width (Pack of 100)</td>
			<td>Amazon</td>
			<td>B000FMYRHK</td>
			<td></td>
		</tr>
		<tr>
			<td>30 mL Syringes without Needle</td>
			<td>Vitality Medical</td>
			<td>302832</td>
			<td></td>
		</tr>
		<tr>
			<td>3-Way Stopcock, 2 Female Luer Locks, Swivel Male Luer Lock, Vented Cap</td>
			<td>QOSINA</td>
			<td>2C6201</td>
			<td></td>
		</tr>
		<tr>
			<td>4-40 X 1/2 PH PAN MS SS/CHROME &#38; appropriate sized phillips screwdriver</td>
			<td>Brikksen Stainless Steel Fastners</td>
			<td>PPMSSSCH4C.5</td>
			<td>&#160;</td>
		</tr>
		<tr>
			<td>ANPROLENE 16 LARGE AMPULE</td>
			<td>Fisher Scientific</td>
			<td>NC9085343</td>
			<td>&#160;</td>
		</tr>
		<tr>
			<td>Betadine</td>
			<td>Purdue</td>
			<td>PUR1815001EACH</td>
			<td>&#160;</td>
		</tr>
		<tr>
			<td>Corning 100 x 20mm tissue-culture treated culture dishes</td>
			<td>Sigma-Aldrich</td>
			<td>CLS430167-100EA</td>
			<td>&#160;</td>
		</tr>
		<tr>
			<td>Corning&#160;L-glutamine Solution</td>
			<td>Fisher Scientific</td>
			<td>MT25005CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Covidien 3033 Curity Gauze Sponge, 4&#34; x 4&#34;, 12-Ply, Sterile, 1200/CS</td>
			<td>Med Plus Medical Supply</td>
			<td>COV-3033-CS</td>
			<td></td>
		</tr>
		<tr>
			<td>Dressing Forceps Delicate Curved (serrated)</td>
			<td>Katena</td>
			<td>K5-4010</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 - Fine Forceps</td>
			<td>F.S.T.</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Eye Scissors Standard Curved</td>
			<td>Katena</td>
			<td>K4-7410</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon 150 x 15mm Plain Sterile Disposable Petri Dishes</td>
			<td>Capitol Scientific</td>
			<td>351058</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand 4 oz. Specimen Containers</td>
			<td>Fisher Scientific</td>
			<td>16-320-730</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Instant Sealing Sterilization Pouches</td>
			<td>Fisher Scientific</td>
			<td>01-812-54</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Instant Sealing Sterilization Pouches</td>
			<td>Fisher Scientific</td>
			<td>01-812-55</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Instant Sealing Sterilization Pouches</td>
			<td>Fisher Scientific</td>
			<td>01-812-58</td>
			<td></td>
		</tr>
		<tr>
			<td>HyClone Dulbecco&#39;s Modified Eagles Medium</td>
			<td>Fisher Scientific</td>
			<td>SH3024302</td>
			<td></td>
		</tr>
		<tr>
			<td>HyClone Penicillin Streptomycin 100X Solution</td>
			<td>Fisher Scientific</td>
			<td>SV30010</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrophilic Filter with Female Luer Lock Inlet, Male Luer Slip Outlet, Blue and Clear</td>
			<td>Qosina</td>
			<td>28217</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrostatic pressure transducers, DELTRAN &#174; II, Catalog # DPT-200 with a 3CC/HR flow rate</td>
			<td>AD instruments</td>
			<td>DPT-200</td>
			<td></td>
		</tr>
		<tr>
			<td>JG15-0.5HPX 15 Gauge 0.5&#34; NT Premium Series Dispensing Tip 50/Box</td>
			<td>Jenson Global</td>
			<td>JG15-0.5HPX 15</td>
			<td></td>
		</tr>
		<tr>
			<td>Keyence B2&#8208;X710 microscope</td>
			<td>Keyence</td>
			<td>B2-X710</td>
			<td></td>
		</tr>
		<tr>
			<td>LabChart 8</td>
			<td>AD instruments</td>
			<td>LabChart 8</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica ST5020 Multi-stainer</td>
			<td>Leica</td>
			<td>ST5020</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-Vented Universal Luer Lock Cap, White</td>
			<td>QOSINA</td>
			<td>65811</td>
			<td></td>
		</tr>
		<tr>
			<td>Octal Bridge Amp (Model # FE228)</td>
			<td>AD instruments</td>
			<td>FE228</td>
			<td></td>
		</tr>
		<tr>
			<td>Pharmco Products ETHYL ALCOHOL, 200 PROOF</td>
			<td>Fisher Scientific</td>
			<td>NC1675398</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Solution (PBS)</td>
			<td>Sigma-Aldrich</td>
			<td>D8537-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>PowerLab 8/35 (Model # PL3508)</td>
			<td>AD instruments</td>
			<td>PL3508</td>
			<td></td>
		</tr>
		<tr>
			<td>ProLong Gold Antifade Mountant with DAPI</td>
			<td>ThermoFisher</td>
			<td>P36935</td>
			<td></td>
		</tr>
		<tr>
			<td>Push-to-Connect Tube Fitting for Air and Water&#160;Straight Adapter, 1/8&#34; Tube OD x 1/8 NPT Male</td>
			<td>McMAster-Carr</td>
			<td>7880T113</td>
			<td></td>
		</tr>
		<tr>
			<td>Push-to-Connect Tube Fitting with Universal Thread&#160;for Air and Water, Adapter, 1/8&#34; Tube OD x 1/8 Pipe</td>
			<td>McMAster-Carr</td>
			<td>51235K101</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain Tygon S3 E-3603 Flexible Tubing 500 ft.</td>
			<td>Fisher Scientific</td>
			<td>14-171-268</td>
			<td></td>
		</tr>
		<tr>
			<td>Superblock T20</td>
			<td>Fisher Scientific</td>
			<td>PI37536</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Scissors - Sharp-Blunt</td>
			<td>F.S.T.</td>
			<td>14001-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Forceps Delicate 1x2 Teeth Curved</td>
			<td>Katena</td>
			<td>K5-4110</td>
			<td></td>
		</tr>
		<tr>
			<td>Translaminar Autonomous System (TAS)</td>
			<td>University of North Texas Health Science Center</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>USA Size 030 O-ring Buna-N, B1000, 70 Durometer, Black, Buna-N 
			(NBR, Nitrile, Buna)</td>
			<td>Marco Rubber &#38; Plastics</td>
			<td>B1000-030</td>
			<td></td>
		</tr>
	</tbody>
</table>

translaminar autonomous system model for the modulation of intraocular and intracranial pressure in human donor posterior segments

There is a current unmet need for a new preclinical human model that can target disease etiology ex vivo using intracranial pressure (ICP) and intraocular pressure (IOP) which&#160;can identify various pathogenic paradigms related to the glaucoma pathogenesis. Ex vivo human anterior segment perfusion organ culture models have previously been successfully utilized and applied as effective technologies for the discovery of glaucoma pathogenesis and testing of therapeutics. Preclinical drug screening and research performed on ex vivo human organ systems can be more translatable to clinical research. This article describes in detail the generation and operation of a novel ex vivo human translaminar pressure model called the translaminar autonomous system (TAS). The TAS model can independently regulate ICP and IOP using human donor posterior segments. The model allows for studying pathogenesis in a preclinical manner. It can reduce the use of living animals in ophthalmic research. In contrast to in vitro experimental models, optic nerve head (ONH) tissue structure, complexity, and integrity can also be maintained within the ex vivo TAS model.

Design and creation of the translaminar autonomous system 
Translaminar pressure differential is a potential key mechanism in the pathogenesis of various diseases, including glaucoma. Uses for the model described include, but are not limited to, the study of glaucoma (elevated IOP, perhaps decreased ICP), traumatic brain injury (elevated ICP), and long-term exposure to microgravity-associated visual impairment (elevated ICP, elevated IOP). To help discover molecular pathogenesis targeting translamin...

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Translaminar Autonomous System Model for the Modulation of Intraocular and Intracranial Pressure in Human Donor Posterior Segments

We describe and detail the use of the translaminar autonomous system. This system utilizes the human posterior segment to independently regulate the pressure inside the segment (intraocular) and surrounding the optic nerve (intracranial) to generate a translaminar pressure gradient that mimics features of glaucomatous optic neuropathy.

There is a current unmet need for a new preclinical human model that can target disease etiology ex vivo using intracranial pressure (ICP) and intraocular ...

translaminar-autonomous-system-model-for-modulation-intraocular

Research

JoVE Journal

Neuroscience

2.9K Views.  University of North Texas Health Science Center. We describe and detail the use of the translaminar autonomous system. This system utilizes the human posterior segment to independently regulate the pressure inside the segment (intraocular) and surrounding the optic nerve (intracranial) to generate a translaminar pressure gradient that mimics features of glaucomatous optic neuropathy.

Video: Translaminar Autonomous System Model for the Modulation of Intraocular and Intracranial Pressure in Human Donor Posterior Segments

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Enrichment of Bruch's Membrane from Human Donor Eyes

Age-related macular degeneration (AMD) is a leading cause of visual impairment in the developed world. The disease manifests itself by the destruction of the center&#160;of the retina, called the macula, resulting in the loss of central vision. Early AMD is characterised by the presence of small, yellowish lesions called soft drusen that can progress onto late AMD such as geographic atrophy (dry AMD) or neovascularisation (wet AMD). Although the clinical changes are well described, and the understanding of genetic influences on conferring AMD risk are getting ever more detailed, one area lacking major progress is an understanding of the biochemical consequences of genetic risk. This is partly due to difficulties in understanding the biochemistry of Bruch&#8217;s membrane, a very thin extracellular matrix that acts as a biological filter of material from the blood supply and a scaffold on which the retinal pigment epithelial (RPE) cell monolayer resides. Drusen form within Bruch&#8217;s membrane and their presence disrupts nutrient flow to the RPE cells. Only by investigating the protein composition of Bruch&#8217;s membrane, and indeed how other proteins interact with it, can researchers hope to unravel the biochemical mechanisms underpinning drusen formation, development of AMD and subsequent vision loss. This paper details methodologies for enriching either whole Bruch&#8217;s membrane, or just from the macula region, so that it can be used for downstream biochemical analysis, and provide examples of how this is already changing the understanding of Bruch&#8217;s membrane biochemistry.

Enrichment of Bruch&#039;s Membrane from Human Donor Eyes

Identifying proteins specifically associated with Bruch&#8217;s membrane in human eyes is an important step in understanding the biochemical mechanisms behind eye diseases such as age-related macular degeneration. This protocol describes how to enrich this sheet of extracellular matrix for down-stream biochemical analysis.

Age-related macular degeneration (AMD) is a leading cause of visual impairment in the developed world. The disease manifests itself by the destruction of ...

Autonomous and Rechargeable Microneurostimulator Endoscopically Implantable into the Submucosa

Gastric dysmotility can be a sign of common diseases such as longstanding diabetes mellitus. It is known that the application of high-frequency low-energetic stimulation can help to effectively moderate and alleviate the symptoms of gastric dysmotility. The goal of the research was the development of a miniature, endoscopically implantable device to a submucosal pocket. The implantable device is a fully customized electronic package which was specifically designed for the purpose of experiments in the submucosa. The device is equipped with a lithium-ion battery which can be recharged wirelessly by receiving an incident magnetic field from the charging/transmitting coil. The uplink communication is achieved in a MedRadio band at 432 MHz. The device was endoscopically inserted into the submucosal pocket of a live domestic pig used as an in vivo model, specifically in the stomach antrum. The experiment confirmed that the designed device can be implanted into the submucosa and is capable of bidirectional communication. The device can perform bipolar stimulation of muscle tissue.

The application of high-frequency low-energetic stimulation can alleviate the symptoms of gastric dysmotility. In this research, a miniature, endoscopically implantable and wirelessly rechargeable device which is implanted into a submucosal pocket is presented. Successful both-way communication and stimulation control were achieved during an experiment on live pig.

Gastric dysmotility can be a sign of common diseases such as longstanding diabetes mellitus. It is known that the application of high-frequency ...

Analyzing Neural Activity and Connectivity Using Intracranial EEG Data with SPM Software

Measuring neural activity and connectivity associated with cognitive functions at high spatial and temporal resolutions is an important goal in cognitive neuroscience. Intracranial electroencephalography (EEG) can directly record electrical neural activity and has the unique potential to accomplish this goal. Traditionally, averaging analysis has been applied to analyze intracranial EEG data; however, several new techniques are available for depicting neural activity and intra- and inter-regional connectivity. Here, we introduce two analytical protocols we recently applied to analyze intracranial EEG data using the Statistical Parametric Mapping (SPM) software: time-frequency SPM analysis for neural activity and dynamic causal modeling of induced responses for intra- and inter-regional connectivity. We report our analysis of intracranial EEG data during the observation of faces as representative results. The results revealed that the inferior occipital gyrus (IOG) showed gamma-band activity at very early stages (110 ms) in response to faces, and both the IOG and amygdala showed rapid intra- and inter-regional connectivity using various types of oscillations. These analytical protocols have the potential to identify the neural mechanisms underlying cognitive functions with high spatial and temporal profiles.

We present two analytical protocols that can be used to analyze intracranial electroencephalography data using the Statistical Parametric Mapping (SPM) software: time-frequency statistical parametric mapping analysis for neural activity, and dynamic causal modeling of induced responses for intra- and inter-regional connectivity.

Measuring neural activity and connectivity associated with cognitive functions at high spatial and temporal resolutions is an important goal in cognitive ...

How to Obtain Reliable Visual Event-related Potentials in Newborns

The present study discusses the characteristics of visual event-related potentials (VEPs) and outlines methodological steps for obtaining reliable measurements in newborns. Obtaining high-quality, reliable VEPs is crucial for the early detection of abnormal development of the central nervous system in at-risk newborns, and for implementing successful early interventions. Recommendations are based on a previous study which showed that when post-conceptional age, polysomnography-identified sleep stages, and light-emitting diodes (LEDs) googles as the luminous source are controlled, no more than 4 repetitions of VEP averages are required to obtain replicable recordings, variability decreases, and reliable VEPs can be obtained. By controlling for these sources of variability and using statistical analyses, we were able to clearly and reliably identify the amplitude and latency of three main components (NII, PII and NIII) present in 100% of newborns (n = 20) during active sleep. Recording VEPs during awake states, quiet sleep and transitional sleep is not recommended because VEP morphology may differ significantly from one average to the next, leading to the risk of misleading clinical prognoses. Moreover, it is easier to obtain VEPs during active sleep because this state can be clearly and reliably identified at this stage of development, sleep cycles are short enough to allow measurements to be taken in a reasonable time, and the method does not require new o expensive equipment.

Several important points for obtaining high-quality reliable visual evoked potentials (VEPs) in newborns while minimizing variability and the risk of misleading prognoses are presented.

The present study discusses the characteristics of visual event-related potentials (VEPs) and outlines methodological steps for obtaining reliable ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Simultaneous Application of Transcranial Direct Current Stimulation during Virtual Reality Exposure

Transcranial direct current stimulation (tDCS) is a form of non-invasive brain stimulation that changes the likelihood of neuronal firing through modulation of neural resting membranes. Compared to other techniques, tDCS is relatively safe, cost-effective, and can be administered while individuals are engaged in controlled, specific cognitive processes. This latter point is important as tDCS may predominantly affect intrinsically active neural regions. In an effort to test tDCS as a potential treatment for psychiatric illness, the protocol described here outlines a novel procedure that allows the simultaneous application of tDCS during exposure to trauma-related cues using virtual reality (tDCS+VR) for veterans with posttraumatic stress disorder (NCT03372460). In this double-blind protocol, participants are assigned to either receive 2 mA tDCS, or sham stimulation, for 25 minutes while passively watching three 8-minute standardized virtual reality drives through Iraq or Afghanistan, with virtual reality events increasing in intensity during each drive. Participants undergo six sessions of tDCS+VR over the course of 2-3 weeks, and psychophysiology (skin conductance reactivity) is measured throughout each session. This allows testing for within and between session changes in hyperarousal to virtual reality events and adjunctive effects of tDCS. Stimulation is delivered through a built-in rechargeable battery-driven tDCS device using a 1 (anode) x 1 (cathode) unilateral electrode set-up. Each electrode is placed in a 3 x 3 cm (current density 2.22 A/m2) reusable sponge pocket saturated with 0.9% normal saline. Sponges with electrodes are attached to the participant&#8217;s skull using a rubber headband with the electrodes placed such that they target regions within the ventromedial prefrontal cortex. The virtual reality headset is placed over the tDCS montage in such a way as to avoid electrode interference.

This manuscript outlines a novel protocol to allow the simultaneous application of transcranial direct current stimulation during exposure to warzone trauma-related cues using virtual reality for veterans with posttraumatic stress disorder.

Transcranial direct current stimulation (tDCS) is a form of non-invasive brain stimulation that changes the likelihood of neuronal firing through ...

Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates reafferent cues that animals must disentangle from relevant sensory cues of the surrounding environment. For example, when a fish swims, flow generated from body undulations is detected by the mechanoreceptive neuromasts, comprising hair cells, that compose the lateral line system. The hair cells then transmit fluid motion information from the sensor to the brain via the sensory afferent neurons. Concurrently, corollary discharge of the motor command is relayed to hair cells to prevent sensory overload. Accounting for the inhibitory effect of predictive motor signals during locomotion is, therefore, critical when evaluating the sensitivity of the lateral line system. We have developed an in vivo electrophysiological approach to simultaneously monitor posterior lateral line afferent neuron and ventral motor root activity in zebrafish larvae (4-7 days post fertilization) that can last for several hours. Extracellular recordings of afferent neurons are achieved using the loose patch clamp technique, which can detect activity from single or multiple neurons. Ventral root recordings are performed through the skin with glass electrodes to detect motor neuron activity. Our experimental protocol provides the potential to monitor endogenous or evoked changes in sensory input across motor behaviors in an intact, behaving vertebrate.

We describe a protocol to monitor changes in the afferent neuron activity during motor commands in a model vertebrate hair cell system.

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates ...

Caenorhabditis elegans as a Model System for Discovering Bioactive Compounds Against Polyglutamine-Mediated Neurotoxicity

Age-related misfolding and aggregation of pathogenic proteins are responsible for several neurodegenerative diseases. For example, Huntington&#39;s disease (HD) is principally driven by a CAG nucleotide repeat that encodes an expanded glutamine tract in huntingtin protein. Thus, the inhibition of polyglutamine (polyQ) aggregation and, in particular, aggregation-associated neurotoxicity is a useful strategy for the prevention of HD and other polyQ-associated conditions. This paper introduces generalized experimental protocols to assess the neuroprotective capacity of test compounds against HD using established polyQ transgenic Caenorhabditis elegans models. The AM141 strain is chosen for the polyQ aggregation assay as an age-associated phenotype of discrete fluorescent aggregates can be easily observed in its body wall at the adult stage due to muscle-specific expression of polyQ::YFP fusion proteins. In contrast, the HA759 model with strong expression of polyQ-expanded tracts in ASH neurons is used to examine neuronal death and chemoavoidance behavior. To comprehensively evaluate the neuroprotective capacity of target compounds, the above test results are ultimately presented as a radar chart with profiling of multiple phenotypes in a manner of direct comparison and direct viewing.

Caenorhabditis elegans as a Model System for Discovering Bioactive Compounds Against Polyglutamine-Mediated Neurotoxicity

Here, we present a protocol to assess the neuroprotective activities of test compounds in Caenorhabditis elegans, including polyglutamine aggregation, neuronal death, and chemoavoidance behavior, as well as an exemplary integration of multiple phenotypes.

Age-related misfolding and aggregation of pathogenic proteins are responsible for several neurodegenerative diseases. For example, Huntington&#39;s ...

Intracranial Pressure Monitoring In Nontraumatic Intraventricular Hemorrhage Rodent Model

Survivors of intraventricular hemorrhage are often left with significant long-term memory impairment; thus, research utilizing intraventricular hemorrhage animal models is essential. In this study, we sought out ways to measure intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during nontraumatic intraventricular hemorrhage in rats. The experimental design included three Sprague Dawley groups: sham, standard 200 &#181;l intraventricular hemorrhage, and vehicle control groups. By introducing an intraparenchymal fiberoptic pressure sensor, precise intracranial pressure measurements were obtained in all groups. Cerebral perfusion pressures were calculated with the knowledge of intracranial pressure and mean arterial pressure values. As expected, the intraventricular hemorrhage and vehicle control groups both experienced a rise in the intracranial pressure and subsequent decline in cerebral perfusion pressure during intraventricular injection of autologous blood and artificial cerebrospinal fluid, respectively. The addition of an intraparenchymal fiberoptic pressure sensor is beneficial in monitoring precise intracranial pressure changes.

Monitoring intracranial pressure in rodent&#160;models of nontraumatic intraventricular hemorrhage is not common in the current literature. Herein, we demonstrate a technique for measuring intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during intraventricular hemorrhage in a rat&#160;animal model.

Survivors of intraventricular hemorrhage are often left with significant long-term memory impairment; thus, research utilizing intraventricular hemorrhage ...