Name: corneal donor tissue preparation for endothelial keratoplasty
Uploaded: 2012-06-12T12:00:00
Duration: 8 min 37 s
Description: Watch this Scientific Journal Video about Corneal Donor Tissue Preparation for Endothelial Keratoplasty at JoVE.com

Supported in part by NIH/NEI EY017885 (RMS).

1. Setup: Aseptic Technique11
<ol>
 <li>Turn on Laminar Flow Hood (LFH).</li>
 <li>Turn on the Nitrogen gas control unit. Ensure that hose is connected between console unit and Nitrogen tank. Turn gas on. Gas setting should be set to 50-60 PSI on regulator tank. Turbine pressure on unit should read 3.3 to 3.4 Bars.</li>
 <li>Press Test button. Atmospheric pressure will read in the 700s and pumps 1 and 2 should be below 200. The unit will beep when testing is complete. Don mask with eye protection, as applicable, and cap. Wash hands and don non-sterile gloves. Place sterile kits inside LFH and open wraps to establish sterile field.</li>
 <li>Place one sterile medicine cup and one sterile plastic basin with sterile gauze next to the established sterile field. Pour sterile isopropyl alcohol into medicine cup. Wipe pachymetry probe with alcohol swab/pad and place probe, tip down, into medicine cup. The probe tip should soak in isopropyl alcohol for 10 minutes. After pachymetry probe has soaked for 10 minutes, rinse the probe tip with sterile water from the prefilled syringe to remove alcohol and place probe in the plastic basin.</li>
 <li>Using aseptic technique, open and drop sterile items (blade, sterile gloves, and corneal viewing chambers) onto sterile field unless already part of the pre-made pack.</li>
 <li>Obtain corneal tissue in preservation media and place inside LFH (Figure 2).</li>
 <li>Open vial(s) and dispose of the lid(s) into an appropriate biohazard receptacle. Discard non-sterile gloves.</li>
 <li>Don sterile gloves. Place pre-packed plastic basin to the side of the field where pachymeter is located. Connect infusion line to stopcock and place spike end of infusion line outside of hood securing the line in such a way as to prevent the line outside of the hood from reentering the hood and touching the sterile field.</li>
 <li>Perform corneal tissue transfer. Gently pour the tissue and medium, in a single motion, from the existing vial to a sterile viewing chamber. Be sure to keep the endothelial side up. Place the cap on the chamber. Keep chambers containing tissue and media on sterile field. Remove vial(s) from LFH. Remove and discard sterile gloves.</li>
 <li>Spike balanced salt solution (BSS) bottle with infusion line. Hang bottle of sterile BSS approximately four feet from working area surface.</li>
 <li>Open two pairs of sterile gloves and sterile gown with towel.</li>
 <li>Perform six minute surgical scrub and dry hands.</li>
 <li>Put on sterile gown, then both pairs of sterile gloves.</li>
 <li>Open instrument boxes and place the contents onto the sterile field.</li>
 <li>Dispense 10-20 cc of BSS solution into metal basin or medicine cup. Fill sterile pipette with BSS solution. Connect stopcock and infusion line to infusion port of the anterior chamber and flush with BSS so that a small amount is visible on top of the piston.</li>
 <li>Position artificial chamber in LFH on top of gauze. Lower piston by turning graft tightening ring counterclockwise.</li>
</ol>
2. Procedure
<ol>
 <li>Using forceps, gently grasp cornea by scleral rim. Open stopcock to flush system with BSS while carefully centering cornea on top of the piston with the endothelial side down. If bubbles are visible beneath the cornea, continue infusing with BSS and slightly moving the cornea to remove them. Discontinue infusion once air bubbles are removed. Ensure that the cornea is centered on the piston (Figure 3a).</li>
 <li>Position anterior chamber cover on base locking ring so that the tabs of the cover fit into the open spaces on the base. Lock cover into base by turning cover clockwise 15&deg;. Raise piston up and tighten completely (Figure 3b).</li>
 <li>With a sterile gloved finger, touch top of cornea to ensure that there is sufficient pressure to perform procedure.</li>
 <li>Check pachymetry by placing probe on surface of cornea to obtain thickness reading (Figure 4). The standard deviation must be less than 10. If not, retake measurement. With the sterile gloved hand that was not used to perform the pachymetry, pass the turbine hose to the hand that performed pachymetry and screw the turbine hose into the control unit. Have the assisting technician record pachymetry reading.</li>
 <li>Remove contaminated gloves.</li>
 <li>Using sterile marking pen, draw a 4mm line from limbus toward center of the cornea (or do not mark if surgeon prefers no marking). Use pipette to place five drops of BSS on top of the cornea.</li>
 <li>Unless specific instructions are provided by the surgeon, the 300 &mu;m cutting head will be used if the pre-cut pachymetry measurement is less than 600 &mu;m. If over 600 &mu;m, the 350 &mu;m cutting head will be used.</li>
 <li>With thread facing toward you and number facing up, load blade into right side of microkeratome head, pointed end first, avoiding contact with the blade.</li>
 <li>Holding head at each side, screw head onto turbine. Tighten the artificial chamber with fingers.</li>
 <li>Screw turbine onto turbine hose.</li>
 <li>Test the blade oscillation by pressing the vacuum pedal once, submerging the cutting head in the sterile water and pulse the turbine for 5 seconds.</li>
 <li>Retighten piston.</li>
 <li>Place microkeratome onto the base with mounting post and position at five o'clock. Press and hold turbine pedal to oscillate blade (Figure 5a).</li>
 <li>Using index finger and thumb, hold microkeratome midway between the base and the top of the turbine. Turn wrist to make a smooth, even cut through and across the cornea. Release turbine pedal and lift microkeratome straight off unit. Press vacuum pedal to turn off vacuum (Figure 5b).</li>
 <li>Don a sterile glove and recheck pachymetry reading after corneal cap is removed by placing probe on corneal surface to obtain second thickness reading being mindful that the tissue is very thin and easily damaged. Remove contaminated glove. Have assisting technician record second pachymetry reading.</li>
 <li>Using sterile marking pen, place a mark if requested by surgeon, on the mid-peripheral edge of the residual stromal bed in the artificial chamber. Place a few drops of sterile BSS onto residual stromal bed.</li>
 <li>Remove the anterior cornea cap from the microkeratome (Figure 6) and replace it onto the cornea before removing cornea from artificial anterior chamber. Cap should be centered on cornea using peripheral marking to align the cap correctly if applicable. Use swab spear to gently reposition the cap onto the cornea. Use fresh spear to absorb any BSS underneath anterior cap.</li>
 <li>Open stopcock, turn chamber upside-down and lower piston slow enough to prevent cornea from deforming. Unlock base and allow pressure from the BSS flow to push the artificial chamber cup from the base. With forceps, gently pull at the corneal rim at 9, 12 and 3 o'clock positions to loosen it from the cup. Once loosened, use forceps at the 12 o'clock position to remove cornea. Place the cornea in the viewing chamber, endothelial side up (Figure 7). Place the lid on the viewing chamber. Remove sterile glove.</li>
 <li>Tighten viewing chamber lid.</li>
 <li>Perform post-cut slit lamp evaluation and specular microscopy of cornea and record findings.</li>
 <li>Place chamber(s) in appropriate section of refrigerator.</li>
 <li>Package for transport to surgeon (Figure 8).</li>
</ol>
3. Representative Results
Proper mechanical microkeratome dissection of the donor cornea results in smooth, uniform lamellar corneal donor tissue. This is shown by an optical coherence tomography (OCT) image of the cornea in cross section (Figure 9). The final posterior corneal tissue should be of adequate thickness as measured by pachymetry. Over the past 6 months at the Michigan Eye Bank, the mean pre-processing corneal thickness was 558 microns, and post-processing thickness of the posterior corneal tissue was 158 microns. Eighty-seven percent of microkeratome-dissected donor tissue measured between 100-200 microns, 10.9% measured &gt;200 microns, and 2.1% measured &lt;100 microns.
The posterior corneal tissue must retain high endothelial cell density (ECD) to provide improved endothelial function to the transplant recipient. Specular microscopy is used to measure ECD. In the past 6 months, all of the corneal tissue deemed eligible for endothelial keratoplasty from the Michigan Eye Bank had an ECD of &gt;2200 cells/mm2. Ninety-nine percent of the precut DSAEK tissue had a post-processing ECD &gt;2400 cells/mm2, and 19% had an ECD &gt; 3000 cells/mm2. The mean change in pre- to post-processing ECD was negligible at 1.2% (p=0.0003, paired two-tailed t-test). Complications that can occur during the procedure include loss of pressure, asymmetric dissection, and significant tissue indentation (resulting in corneal endothelial cell loss). Improper dissection can also result in tissue perforation.
<img alt="Figure 1" src="/files/ftp_upload/3847/3847fig1.jpg" /> 
Figure 1. Overall Scheme of Corneal Tissue Preparation for Endothelial Keratoplasty Figures 1a to 1e are cross-section representations of the donor cornea preparation. Figures 1f to 1i are cross-section representations of the transplant recipient's cornea during DSAEK surgery.
<ol class="lalpha">
 <li>The donor cornea is mounted onto an artificial anterior chamber and dissected with the microkeratome head.</li>
 <li>After tissue dissection, the two lamellae are separate but the anterior lamella (cap) is positioned back onto the remained corneal bed for transportation to the surgeon.</li>
 <li>In the operating room, the surgeon uses a trephine to cut the central corneal tissue vertically through both anterior and posterior lamella.</li>
 <li>After trephine cutting, the peripheral donor cornea is removed from the field.</li>
 <li>The posterior lamella is gently removed from the anterior lamella with a spatula. The donor posterior lamella (DSAEK tissue) replaces the diseased portion of the recipient cornea (1i).</li>
 <li>The patient has a diseased posterior cornea to be removed.</li>
 <li>The diseased posterior cornea is scored with an intraocular instrument (reverse Sinsky hook) in a large diameter circle.</li>
 <li>After scoring the diseased cornea, the posterior corneal membrane is removed.</li>
 <li>The donor posterior lamella (DSAEK tissue) is placed in the recipient's eye.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/3847/3847fig2.jpg" /> 
Figure 2. Corneal Tissue in Media. Corneal tissue is in preservation media after harvesting from donor.
<img alt="Figure 3" src="/files/ftp_upload/3847/3847fig3.jpg" /> 
Figure 3. Corneal Tissue on Artificial Anterior Chamber.
<ol class="lalpha">
 <li>Corneal tissue is mounted onto the artificial anterior chamber prior base locking ring or tissue dissection.</li>
 <li>Corneal tissue is mounted onto artificial anterior chamber with base locking ring sealed over the tissue.</li>
</ol>
<img alt="Figure 4" src="/files/ftp_upload/3847/3847fig4.jpg" /> 
Figure 4. Corneal Tissue Thickness Check. The corneal thickness is checked using ultrasound pachymetry probe on surface of cornea.
<img alt="Figure 5" src="/files/ftp_upload/3847/3847fig5.jpg" /> 
Figure 5. Corneal Tissue Dissection.
<ol class="lalpha">
 <li>Microkeratome blade is placed on the base with mounting post and position at five o'clock.</li>
 <li>Microkeratome is passed through cornea to make the lamellar dissection.</li>
</ol>
<img alt="Figure 6" src="/files/ftp_upload/3847/3847fig6.jpg" /> 
Figure 6. Anterior Cap of Cornea after Dissection. The anterior cornea cap is free after microkeratome dissection is complete.
<img alt="Figure 7" src="/files/ftp_upload/3847/3847fig7.jpg" /> 
Figure 7. Dissected Cornea in Viewing Chamber. After dissection is completed, the cornea is returned to the viewing chamber for transportation.
<img alt="Figure 8" src="/files/ftp_upload/3847/3847fig8.jpg" /> 
Figure 8. Cornea in Packaging for Transportation. The cornea in the viewing chamber is then securely packaged for transportation to the surgeon.
<img alt="Figure 9" src="/files/ftp_upload/3847/3847fig9.jpg" /> 
Figure 9. OCT Appearance of Cornea. The OCT of the final dissected cornea (in cross-section) shows that the cornea is split into two halves. The interface is more intensely reflective than the surrounding tissue as indicated by the arrow.

<ol>
	<li>Melles, G. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+surgical+technique+for+posterior+lamellar+keratoplasty.">A surgical technique for posterior lamellar keratoplasty.</a> Cornea. 17, 618-626 (1998).</li><li>Melles, G. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preliminary+clinical+results+of+posterior+lamellar+keratoplasty+through+a+sclerocorneal+pocket+incision.">Preliminary clinical results of posterior lamellar keratoplasty through a sclerocorneal pocket incision.</a> Ophthal. 107, 1850-1856 (2000).</li><li>Terry, M. A., Ousley, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Replacing+the+endothelium+without+corneal+surface+incisions+or+sutures:+the+first+United+States+clinical+series+using+the+deep+lamellar+endothelial+keratoplasty+procedure.">Replacing the endothelium without corneal surface incisions or sutures: the first United States clinical series using the deep lamellar endothelial keratoplasty procedure.</a> Ophthal. 110, 755-764 (2003).</li><li>Price, F. W., Price, M. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Descemet's+stripping+with+endothelial+keratoplasty+in+50+eyes:+a+refractive+neutral+corneal+transplant.">Descemet's stripping with endothelial keratoplasty in 50 eyes: a refractive neutral corneal transplant.</a> J. Refract Surg. 21, 339-345 (2005).</li><li>Price, M. O., Price, F. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Descemet's+stripping+with+endothelial+keratoplasty:+comparative+outcomes+with+microkeratome-dissected+and+manually+dissected+donor+tissue.">Descemet's stripping with endothelial keratoplasty: comparative outcomes with microkeratome-dissected and manually dissected donor tissue.</a> Ophthal. 113, 1936-1942 (2006).</li><li>Bahar, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retrospective+contralateral+study+comparing+descemet+stripping+automated+endothelial+keratoplasty+with+penetrating+keratoplasty.">Retrospective contralateral study comparing descemet stripping automated endothelial keratoplasty with penetrating keratoplasty.</a> Cornea. 28, 485-488 (2009).</li><li>Rose, L., Briceno, C. A., Stark, W. J., Gloria, D. G., Jun, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+Eye+Bank-Prepared+Posterior+Lamellar+Corneal+Tissue+for+Endothelial+Keratoplasty.">Assessment of Eye Bank-Prepared Posterior Lamellar Corneal Tissue for Endothelial Keratoplasty.</a> Ophthal. 115, 279-286 (2008).</li><li>Price, M. O., Baig, K. M., Brubaker, J. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=prospective+comparison+of+precut+versus+surgeon-dissected+grafts+for+Descemet+stripping+automated+endothelial+keratoplasty.">prospective comparison of precut versus surgeon-dissected grafts for Descemet stripping automated endothelial keratoplasty.</a> Am. J. Ophthalmol. 146, 36-41 (2008).</li><li>Chen, E. S., Terry, M. A., Shamie, N., Hoar, K. L., Friend, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precut+tissue+in+Descemet's+stripping+automated+endothelial+keratoplasty+donor+characteristics+and+early+postoperative+complications.">Precut tissue in Descemet's stripping automated endothelial keratoplasty donor characteristics and early postoperative complications.</a> Ophthalmology. 115, 497-502 (2008).</li><li>Terry, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endothelial+Keratoplasty:+A+Comparision+of+Complication+Rates+and+Endothelial+Survival+between+Precut+Tissue+and+Surgeon-cut+Tissue+by+a+Single+DSAEK+Surgeon.">Endothelial Keratoplasty: A Comparision of Complication Rates and Endothelial Survival between Precut Tissue and Surgeon-cut Tissue by a Single DSAEK Surgeon.</a> Trans. Am. Ophthalmol. Soc. 107, 184-193 (2009).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Reference document E1.710 Corneal Tissue Preparation for Lamellar Procedures. , 1-5 (2010).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Eye Banking Statistical Report. , 1-5 (2009).</li><li>Glasser, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+complications+during+endothelial+keratoplasty.">Tissue complications during endothelial keratoplasty.</a> Cornea. 29, 1428-1429 (2010).</li><li>Kelliher, C., Engler, C., Speck, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comprehensive+analysis+of+eye+bank-prepared+posterior+lamellar+corneal+tissue+for+use+in+endothelial+keratoplasty.">A comprehensive analysis of eye bank-prepared posterior lamellar corneal tissue for use in endothelial keratoplasty.</a> Cornea. 28, 966-970 (2009).</li><li>Price, F. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precut+tissue+for+Descemet+stripping+automated+endothelial+keratoplasty.">Precut tissue for Descemet stripping automated endothelial keratoplasty.</a> Cornea. 27, 630-631 (2008).</li></ol>

Drs. Woodward and Shtein have nothing to disclose. Mr. Mavin and Mr. Titus are employees of Midwest Eye Banks.

The inner-most layer of the cornea, the endothelium, maintains the optical clarity of the cornea by maintaining corneal dehydration. Numerous disease states specifically affect the health and viability of the corneal endothelium including dystrophies, infections, inflammatory processes, and degenerations. Until recently, corneal transplantation to replace diseased corneal endothelium required full thickness transplantation of the cornea. In the past decade, targeted replacement of the posterior cornea by partial thickness corneal transplantation, such as DSAEK surgery, has been able to achieve the same goal with a shorter recovery time. Regardless of technique, transplanted tissues must retain a healthy corneal endothelium to maintain optical clarity in the transplanted cornea; as a result, all tissue processing techniques have additional steps to protect the corneal endothelium.
Automated microkeratome dissection is currently the standard for corneal tissue preparation for endothelial keratoplasty. Use of the microkeratome was shown to decrease donor tissue perforation and speed visual recovery as compared to manual dissection 5. When DSAEK was initially introduced, some surgeons purchased microkeratome equipment and performed donor lamellar dissection in the operating room at the start of surgery. If tissue preparation failed, the surgery would have to be cancelled causing inconvenience and expense to the patient and the surgeon. In 2006 eye banks began performing microkeratome dissection "off site" under controlled conditions in the eye bank laboratory a day prior to surgery 12. If tissue preparation fails in this setting, there is time to perform another dissection with fresh tissue.
Microkeratome dissection of donor posterior lamellar corneal tissue requires gentle and precise technique to minimize complications. The cornea must be placed gently on the artificial anterior chamber (AAC) to avoid any damage to the delicate endothelial cells. Once the cornea is mounted and aligned on the AAC, the locking ring must be positioned and adjusted for a tight seal to maintain high pressure within the chamber and decrease the chances of tissue collapse during the microkeratome dissection. To ensure uniform dissection, the microkeratome head should be rotated in a constant, smooth manner. After dissection, the tissue must again be handled carefully to minimize endothelial cell damage. Further, it is critical to confirm that the lamellar cut is centered and complete in order to ensure that the posterior lamella will be suitable for use in surgery 13.
Corneal donor perforation is a dreaded complication of this procedure; it becomes less frequent with experience. If repeated perforations occur during microkeratome passes, it is important to check multiple steps in the procedure. The cornea needs to be properly mounted and aligned on the AAC. A tight seal and high pressure should be obtained prior to microkeratome pass and maintained throughout the procedure. On occasion, residual conjunctiva exists on the sclera rim and needs to be dissected off prior to mounting. The microkeratome blade should be sharp and without notches. The correct microkeratome head thickness should be selected based on the estimated corneal thickness. The microkeratome head must be oscillating properly prior to the pass. With further problems with this technique, consult the microkeratome provider to verify that all equipment is working properly.
Even when performed perfectly, there are some limitations to this technique. Microkeratome heads are available to cut corneal tissue at various thicknesses; however, the relationship of microkeratome head to final tissue thickness is imprecise with the same head providing a wide range of final tissue thicknesses 7. In addition, a significant learning curve exists and complications in preparation are more frequent for new users. Performing the lamellar dissection in the eye bank setting allows trained technicians who perform this procedure many times per day to optimize their technique, thereby minimizing complications due to inexperience.
Modifications of the technique for microkeratome-assisted corneal dissection for endothelial keratoplasty exist. Some prefer to use viscoelastic material or corneal storage media rather than balanced salt solution beneath the corneal button on the artificial anterior chamber with the intent of providing increased protection to the corneal endothelium. There are also variations in the method for determining the location in which the microkeratome incision should be initiated. Different eye banks and different corneal surgeons may have particular preferences as to how the cornea is marked for identification of the anterior corneal cap or posterior lamellar orientation. It should be noted that manufacturers make similar equipment to that used in this protocol. The particular manufacturer's instructions should be followed with whichever equipment is used.
Dependable eye bank preparation of dissected corneal tissue for DSAEK surgery has been beneficial for the widespread acceptance of this relatively new surgical technique. Once surgeons became confident in eye banks performing the "first step" in corneal transplantation - namely tissue preparation, many practical limitations to DSAEK were alleviated. Eye banks have been able to reliably keep up with the demand for processed tissue with safe, effective, and timely supply of donor tissue. High quality DSAEK tissue can now be ordered in advance and operating room time and tissue preparation complications have decreased 9, 14-15.
This protocol describes the preparation of DSAEK grafts from the perspective of eye bank processing of corneal donor tissue; however, this technique is directly applicable to surgeons interested in preparing their own lamellar corneal donor tissue as well as those interested in performing laboratory-based or clinical studies on lamellar corneal transplantation.
Future directions for corneal lamellar preparation include use of the femtosecond laser for tissue processing, creation of "ultra-thin" DSAEK corneal tissue, or preparation of corneal endothelial scrolls for Descemet's membrane endothelial keratoplasty (DMEK) surgery. Endothelial keratoplasty is a surgical field in evolution and the symbiotic relationship between the eye bank and corneal surgeon will continue to change as surgical techniques are optimized.

<table border="1">
<tbody>
 <tr>
 <td>Name of the Equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Microkeratome</td>
 <td>Moria</td>
 <td>1021174</td>
 </tr>
 <tr>
 <td>Artificial Anterior Chamber base</td>
 <td>Moria</td>
 <td>19161-562</td>
 </tr>
 <tr>
 <td>Artificial Anterior Chamber cap</td>
 <td>Moria</td>
 <td>19172-220</td>
 </tr>
 <tr>
 <td>Microkeratome Head 300</td>
 <td>Moria</td>
 <td>O304</td>
 </tr>
 <tr>
 <td>Microkeratome Head 350</td>
 <td>Moria</td>
 <td>N791</td>
 </tr>
 <tr>
 <td>Console</td>
 <td>Moria</td>
 <td>19360-3482</td>
 </tr>
 </tbody>
</table>

corneal donor tissue preparation for endothelial keratoplasty

Over the past ten years, corneal transplantation surgical techniques have undergone revolutionary changes1,2. Since its inception, traditional full thickness corneal transplantation has been the treatment to restore sight in those limited by corneal disease. Some disadvantages to this approach include a high degree of post-operative astigmatism, lack of predictable refractive outcome, and disturbance to the ocular surface. The development of Descemet's stripping endothelial keratoplasty (DSEK), transplanting only the posterior corneal stroma, Descemet's membrane, and endothelium, has dramatically changed treatment of corneal endothelial disease. DSEK is performed through a smaller incision; this technique avoids 'open sky' surgery with its risk of hemorrhage or expulsion, decreases the incidence of postoperative wound dehiscence, reduces unpredictable refractive outcomes, and may decrease the rate of transplant rejection3-6.
Initially, cornea donor posterior lamellar dissection for DSEK was performed manually1 resulting in variable graft thickness and damage to the delicate corneal endothelial tissue during tissue processing. Automated lamellar dissection (Descemet's stripping automated endothelial keratoplasty, DSAEK) was developed to address these issues. Automated dissection utilizes the same technology as LASIK corneal flap creation with a mechanical microkeratome blade that helps to create uniform and thin tissue grafts for DSAEK surgery with minimal corneal endothelial cell loss in tissue processing.
Eye banks have been providing full thickness corneas for surgical transplantation for many years. In 2006, eye banks began to develop methodologies for supplying precut corneal tissue for endothelial keratoplasty. With the input of corneal surgeons, eye banks have developed thorough protocols to safely and effectively prepare posterior lamellar tissue for DSAEK surgery. This can be performed preoperatively at the eye bank. Research shows no significant difference in terms of the quality of the tissue7 or patient outcomes8,9 using eye bank precut tissue versus surgeon-prepared tissue for DSAEK surgery. For most corneal surgeons, the availability of precut DSAEK corneal tissue saves time and money10, and reduces the stress of performing the donor corneal dissection in the operating room. In part because of the ability of the eye banks to provide high quality posterior lamellar corneal in a timely manner, DSAEK has become the standard of care for surgical management of corneal endothelial disease.
The procedure that we are describing is the preparation of the posterior lamellar cornea at the eye bank for transplantation in DSAEK surgery (Figure 1).

Watch this Scientific Journal Video about Corneal Donor Tissue Preparation for Endothelial Keratoplasty at JoVE.com

Corneal Donor Tissue Preparation for Endothelial Keratoplasty

Endothelial corneal transplantation is a surgical technique for treatment of posterior corneal diseases. Mechanical microkeratome dissection to prepare tissue results in thinner, more symmetric grafts with less endothelial cell loss and improved outcomes. Dissections can be performed at the eye bank prior to corneal transplantation surgery.

Over the past ten years, corneal transplantation surgical techniques have undergone revolutionary changes1,2. Since its inception, traditional full ...

corneal-donor-tissue-preparation-for-endothelial-keratoplasty

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In 2007, Rane presented the first single port nephrectomy for a small non-functioning kidney at the World Congress of Endourology.  Since that time, the ...

Corneal Donor Tissue Preparation for Descemet's Membrane Endothelial Keratoplasty

Descemet&#8217;s Membrane Endothelial Keratoplasty (DMEK) is a form of corneal transplantation in which only a single cell layer, the corneal endothelium, along with its basement membrane (Descemet&#39;s membrane) is introduced onto the recipient&#39;s posterior stroma3. Unlike Descemet&#8217;s Stripping Automated Endothelial Keratoplasty (DSAEK), where additional donor stroma is introduced, no unnatural stroma-to-stroma interface is created. As a result, the natural anatomy of the cornea is preserved as much as possible allowing for improved recovery time and visual acuity4. Endothelial Keratoplasty (EK) is the procedure of choice for treatment of endothelial dysfunction. The advantages of EK include rapid recovery of vision, preservation of ocular integrity and minimal refractive change due to use of a small, peripheral incision1. DSAEK utilizes donor tissue prepared with partial thickness stroma and endothelium. The rapid success and utilization of this procedure can be attributed to availability of eye-bank prepared precut tissue. The benefits of eye-bank preparation of donor tissue include elimination of need for specialized equipment in the operating room and availability of back up donor tissue in case of tissue perforation during preparation. In addition, high volume preparation of donor tissue by eye-bank technicians may provide improved quality of donor tissue. DSAEK may have limited best corrected visual acuity due to creation of a stromal interface between the donor and recipient cornea. Elimination of this interface with transplantation of only donor Descemet&#39;s membrane and endothelium in DMEK may improve visual outcomes and reduce complications after EK5. Similar to DSAEK, long term success and acceptance of DMEK is dependent on ease of availability of precut, eye-bank prepared donor tissue. Here we present a stepwise approach to donor tissue preparation which may reduce some barriers eye-banks face in providing DMEK grafts.

Corneal Donor Tissue Preparation for Descemet&#039;s Membrane Endothelial Keratoplasty

Endothelial keratoplasty is a surgical technique continuously evolving as advancements result in transplantation of thinner grafts with each succession1. Here we present a technique for controlled manual tissue dissection to allow safe and repeatable separation of endothelium and Descemet&#39;s membranes from donor corneoscleral buttons for transplantation2.

Descemet&#8217;s Membrane Endothelial Keratoplasty (DMEK) is a form of corneal transplantation in which only a single cell layer, the corneal endothelium, ...

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene and cell therapies are highly promising alternative approaches for CVD treatment. However, the broad clinical application of gene therapy is greatly limited by the lack of suitable gene delivery systems. The development of appropriate gene delivery vectors can provide a solution to current challenges in cell therapy. In particular, existing drawbacks, such as limited efficiency and low cell retention in the injured organ, could be overcome by appropriate cell engineering (i.e., genetic) prior to transplantation. The presented protocol describes the efficient and safe transient modification of endothelial cells using a polyethyleneimine superparamagnetic magnetic nanoparticle (PEI/MNP)-based delivery vector. Also, the algorithm and methods for cell characterization are defined. The successful intracellular delivery of microRNA (miR) into human umbilical vein endothelial cells (HUVECs) has been achieved without affecting cell viability, functionality, or intercellular communication. Moreover, this approach was proven to cause a strong functional effect in introduced exogenous miR. Importantly, the application of this MNP-based vector ensures cell magnetization, with accompanying possibilities of magnetic targeting and non-invasive MRI tracing. This may provide a basis for magnetically guided, genetically engineered cell therapeutics that can be monitored non-invasively with MRI.

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

This manuscript describes the efficient, non-viral delivery of miR to endothelial cells by a PEI/MNP vector and their magnetization. Thus, in addition to genetic modification, this approach allows for magnetic cell guidance and MRI detectability. The technique can be used to improve the characteristics of therapeutic cell products.

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene ...

A Porcine Corneal Endothelial Organ Culture Model Using Split Corneal Buttons

Experimental research on corneal endothelial cells is associated with several difficulties. Human donor corneas are scarce and rarely available for experimental investigations as they are normally needed for transplantation. Endothelial cell cultures often do not translate well to in vivo situations. Due to the biostructural characteristics of non-human corneas, stromal swelling during cultivation induces substantial corneal endothelial cell loss, which makes it difficult to perform cultivation for an extended period of time. Deswelling agents such as dextran are used to counteract this response. However, they also cause significant endothelial cell loss. Therefore, an ex vivo organ culture model not requiring deswelling agents was established. Pig eyes from a local slaughterhouse were used to prepare split corneal buttons. After partial corneal trephination, the outer layers of the cornea (epithelium, bowman layer, parts of the stroma) were removed. This significantly reduces corneal endothelial cell loss induced by massive stromal swelling and Descemet&#39;s membrane folding throughout longer cultivation periods and improves general preservation of the endothelial cell layer. Subsequent complete corneal trephination was followed by the removal of the split corneal button from the remaining eye bulb and cultivation. Endothelial cell density was assessed at follow-up times of up to 15 days after preparation (i.e., days 1, 8, 15) using light microscopy. The preparation technique used allows a better preservation of the endothelial cell layer enabled by less stromal tissue swelling, which results in slow and linear decline rates in split corneal buttons comparable to human donor corneas. As this standardized organo-typically cultivated research model for the first time allows a stable cultivation for at least two weeks, it is a valuable alternative to human donor corneas for future investigations of various external factors with regards to their effects on the corneal endothelium.

Here, a step-by-step protocol for the preparation and cultivation of porcine split corneal buttons is presented. As this organo-typically cultivated organ culture model shows cell death rates within 15 days, comparable to human donor corneas, it represents the first model allowing long-term cultivation of non-human corneas without adding toxic dextran.

Experimental research on corneal endothelial cells is associated with several difficulties. Human donor corneas are scarce and rarely available for ...

Development of a Noninvasive, Laser-Assisted Experimental Model of Corneal Endothelial Cell Loss

Nd:YAG lasers have been used to perform noninvasive intraocular surgery, such as capsulotomy for several decades now. The incisive effect relies on the optical breakdown at the laser focus. Acoustic shock waves and cavitation bubbles are generated, causing tissue rupture. Bubble sizes and pressure amplitudes vary with pulse energy and position of the focal point. In this study, enucleated porcine eyes were positioned in front of a commercially available Nd:YAG laser. Variable pulse energies as well as different positions of the focal spots posterior to the cornea were tested. Resulting lesions were evaluated by two-photon microscopy and histology to determine the best parameters for an exclusive detachment of corneal endothelial cells (CEC) with minimum collateral damage. The advantages of this method are the precise ablation of CEC, reduced collateral damage, and above all, the non-contact treatment.

Here, we present a protocol to detach corneal endothelial cells (CEC) from Descemet&#8217;s membrane (DM) using a neodymium:YAG (Nd:YAG) laser as an ex vivo disease model for bullous keratopathy (BK).

Nd:YAG lasers have been used to perform noninvasive intraocular surgery, such as capsulotomy for several decades now. The incisive effect relies on the ...

An Epithelial Abrasion Model for Studying Corneal Wound Healing

The cornea is critical for vision, accounting for about two-thirds of the refractive power of the eye. Crucial to the role of the cornea in vision is its transparency. However, due to its external position, the cornea is highly susceptible to a wide variety of injuries that can lead to the loss of corneal transparency and eventual blindness. Efficient corneal wound healing in response to these injuries is pivotal for maintaining corneal homeostasis and preservation of corneal transparency and refractive capabilities. In events of compromised corneal wound healing, the cornea becomes vulnerable to infections, ulcerations, and scarring. Given the fundamental importance of corneal wound healing to the preservation of corneal transparency and vision, a better understanding of the normal corneal wound healing process is a prerequisite to understanding impaired corneal wound healing associated with infection and disease. Toward this goal, murine models of corneal wounding have proven useful in furthering our understanding of the corneal wound healing mechanisms operating under normal physiological conditions. Here, a protocol for creating a central corneal epithelial abrasion in mouse using a trephine and a blunt golf club spud is described. In this model, a 2 mm diameter circular trephine, centered over the cornea, is used to demarcate the wound area. The golf club spud is used with care to debride the epithelium and create a circular wound without damaging the corneal epithelial basement membrane. The resulting inflammatory response proceeds as a well-characterized cascade of cellular and molecular events that are critical for efficient wound healing. This simple corneal wound healing model is highly reproducible and well-published and is now being used to evaluate compromised corneal wound healing in the context of disease.

Here, a protocol for creating a central corneal epithelial abrasion wound in the mouse using a trephine and a blunt golf club spud is described. This corneal wound healing model is highly reproducible and is now being used to evaluate compromised corneal wound healing in the context of diseases.

The cornea is critical for vision, accounting for about two-thirds of the refractive power of the eye. Crucial to the role of the cornea in vision is its ...

A Porcine Heterotopic Heart Transplantation Protocol for Delivery of Therapeutics to a Cardiac Allograft

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts and complications such as primary graft dysfunction and graft rejection. The recent clinical use of an ex vivo perfusion device in cardiac transplantation introduces a unique opportunity for treating cardiac allografts with therapeutic interventions to improve function and avoid deleterious recipient responses. Establishing a translational, large-animal model for therapeutic delivery to the entire allograft is essential for testing novel therapeutic approaches in cardiac transplantation. The porcine, heterotopic heart transplantation model in the intraabdominal position serves as an excellent model for assessing the effects of novel interventions and the immunopathology of graft rejection. This model additionally offers long-term survival for the pig, given that the graft is not required to maintain the recipient's circulation. The aim of this protocol is to provide a reproducible and robust approach for achieving ex vivo delivery of a therapeutic to the entire cardiac allograft prior to transplantation and provide technical details to perform a survival heterotopic transplant of the ex vivo perfused heart.

We present a protocol for utilizing a normothermic ex vivo sanguinous perfusion system for the delivery of therapeutics to an entire cardiac allograft in a porcine heterotopic heart transplant model.

Cardiac transplantation is the gold standard treatment for end-stage heart failure. However, it remains limited by the number of available donor hearts ...

Preparation of Human Myocardial Tissue for Long-Term Cultivation

Cardiomyocyte cultivation has seen a vast number of developments, ranging from two-dimensional (2D) cell cultivation to iPSC derived organoids. In 2019, an ex vivo way to cultivate myocardial slices obtained from human heart samples was demonstrated, while approaching in vivo condition of myocardial contraction. These samples originate mostly from heart transplantations or left-ventricular assist device placements. Using a vibratome and a specially developed cultivation system, 300 &#181;m thick slices are placed between a fixed and a spring wire, allowing for stable and reproducible cultivation for several weeks. During cultivation, the slices are continuously stimulated according to individual settings. Contractions can be displayed and recorded in real-time, and pharmacological agents can be readily applied. User-defined stimulation protocols can be scheduled and performed to assess vital contraction parameters like post-pause-potentiation, stimulation threshold, force-frequency relation, and refractory period. Furthermore, the system enables a variable pre- and afterload setting for a more physiological cultivation.
Here, we present a step-by-step guide on how to generate a successful long-term cultivation of human left ventricular myocardial slices, using a commercial biomimetic cultivation solution.

We present a protocol for ex vivo cultivation of human ventricular myocardial tissue. It allows for detailed analysis of contraction force and kinetics, as well as the application of pre- and afterload to mimic the in vivo physiological environment more closely.

Cardiomyocyte cultivation has seen a vast number of developments, ranging from two-dimensional (2D) cell cultivation to iPSC derived organoids. In 2019, ...

In Vivo Heart Transplantation to Study Endothelial-to-Mesenchymal Transition

A Neonatal Heterotopic Rat Heart Transplantation Model for the Study of Endothelial-to-Mesenchymal Transition

Endocardial fibroelastosis (EFE), defined by subendocardial tissue accumulation, has major impacts on the development of the left ventricle (LV) and precludes patients with congenital critical aortic stenosis and hypoplastic left heart syndrome (HLHS) from curative anatomical biventricular surgical repair. Surgical resection is currently the only available therapeutic option, but EFE often recurs, sometimes with an even more infiltrative growth pattern into the adjacent myocardium.
To better understand the underlying mechanisms of EFE and to explore therapeutic strategies, an animal model suitable for preclinical testing was developed. The animal model takes into consideration that EFE is a disease of the immature heart and is associated with flow disturbances, as supported by clinical observations. Thus, the heterotopic heart transplantation of neonatal rat donor hearts is the basis for this model.
A neonatal rat heart is transplanted into an adolescent rat&#39;s abdomen and connected to the recipient&#39;s infrarenal aorta and inferior vena cava. While perfusion of the coronary arteries preserves the viability of the donor heart, flow stagnation within the LV induces EFE growth in the very immature heart. The underlying mechanism of EFE formation is the transition of endocardial endothelial cells to mesenchymal cells (EndMT), which is a well-described mechanism of early embryonic development of the valves and septa but also the leading cause of fibrosis in heart failure. EFE formation can be macroscopically observed within days after transplantation. Transabdominal echocardiography is used to monitor the graft viability, contractility, and the patency of the anastomoses. Following euthanasia, the EFE tissue is harvested, and it shows the same histopathological characteristics as human EFE tissue from HLHS patients.
This in vivo model allows for studying the mechanisms of EFE development in the heart and testing treatment options to prevent this pathological tissue formation and provides the opportunity for a more generalized examination of EndMT-induced fibrosis.

Author Spotlight: A Neonatal Heterotopic Rat Heart Transplantation Model for the Study of Endothelial-to-Mesenchymal Transition  

This work presents an animal model of endothelial-to-mesenchymal transition-induced fibrosis, as seen in congenital cardiac defects such as critical aortic stenosis or hypoplastic left heart syndrome, which allows for detailed histological tissue evaluation, the identification of regulatory signaling pathways, and the testing of treatment options.

Endocardial fibroelastosis (EFE), defined by subendocardial tissue accumulation, has major impacts on the development of the left ventricle (LV) and ...

Oral Submucous Fibrosis Tissue Sample Preparation for Hematoxylin-Eosin Staining

To begin, using a microtome, cut the formalin fixed paraffin embedded oral submucous fibrosis tissues into continuous sections of three in 10 micrometers thickness, then unfold the sections in water. Place three micron sections on the adhesion microscope slide or slide A, and 10 microns on pen membrane slide or slide B.Place both slides on a hot plate at 60 degrees Celsius. After two hours, place slides A and B in xylene for 10 minutes, then hydrate the slides in graded ethanol concentrations for one minute in each concentration.Next, place the slides in ultrapure distilled water for 30 seconds. Soak slides in Harris hematoxylin dye solution for 90 seconds. Again wash the slides thrice in ultrapure distilled water for 30 seconds each.After that, submerge the slides in div-hematoxylin solution for two seconds. Return slides into ultrapure distilled water for 30 seconds, repeating three times. Next, immerse slides A and B in 1%re-blue solution for two seconds.After washing the slides with distilled water, place them in an eosin solution for two seconds. Again immerse slides in ultrapure distilled water for 30 seconds before dehydrating and increasing ethanol concentrations, then immerse slide A in xylene for three minutes. Finally, apply a drop of the mounting medium to slide A, and cover it with a glass cover.