This work was supported by the National Eye Institute (EY015279 and EY028597) and Delaware INBRE (P20 GM103446).

The following protocol was approved by the University of Delaware Institutional Animal Care and Use Committee under protocol #1039-2021-1. In compliance with the Association for Research in Vision and Ophthalmology (ARVO) for the Use of Animals in Ophthalmic and Vision Research24, all ocular survival surgeries can only be performed in one eye. See Table of Materials for details about reagents and instruments used in this protocol. Alternative approaches for extracting the lens fibers&#160;and suturing&#160;are shown in Figure 2&#160; and&#160;Figure 3.
1. Animals
<ol>
	<li>Use any wild-type or genetically modified laboratory mouse strain that has eyes with intact lenses for this procedure.
	<ol>
		<li>Use C57BL/6 mice as a starting point, as their responses are highly reproducible at both the morphological and molecular levels25,26. 
		NOTE: C57BL/6 or any mouse strain can be used. In the video protocol, the FVB mouse is used for better visual demonstration.</li>
		<li>Perform studies of the response of mutants on other mouse strains with strain-matched controls. 
		NOTE: With practice, this procedure may be performed on mice whose lenses have cataracts as long as the lens capsule is intact and the lens has not undergone significant liquefaction. See the study of mice with lens conditional deletion of the transcription factor Sip127.</li>
	</ol>
	</li>
	<li>Perform this surgery on mice of either sex as young as 8 weeks to as old as 24 months25,26.
	<ol>
		<li>Perform this surgery in even younger mice (after eyelid opening) with practice, although it will be more difficult as these eyes will be significantly smaller.</li>
		<li>Take particular care for surgeries on elderly mice due to their lower tolerance for anesthesia25,26. 
		&#8203;NOTE: Sex differences in surgery responses are more prevalent in elderly mice26.</li>
	</ol>
	</li>
</ol>
2. Preparation of solutions and surgical tools
<ol>
	<li>Obtain anesthetizing and analgesic agents, dilation drops, sterile disposable scalpel, #5 Dumont forceps, 2-110 suture forceps, needles (26 G 1/2 and 27 G 45&#176; bent dispensing tip 1&#34;), syringes, and balanced saline solution (BSS).</li>
	<li>Sterilize surgical instruments in an autoclave.</li>
</ol>
3. Prior to anesthesia
<ol>
	<li>Prior to anesthesia, give the mouse one drop each of 1% atropine and 2.5% phenylephrine to dilate the pupil (see Figure 1A) of the eye undergoing surgery. Apply ophthalmic ointment to the non-operated eye to keep the surface from drying. 
	NOTE: The ARVO statement on the use of animals in eye research24 only allows a single eye to undergo survival surgery, leaving the other eye untouched so that the animal retains vision. Thus, survival surgeries are always unilateral; however, the contralateral, unoperated eye can be operated on just prior to sacrifice to create animal-matched &#39;time zero&#39; controls.</li>
	<li>Make sure the eye drops do not run down the mouse&#39;s face and enter the nose, as this can obstruct its breathing.</li>
	<li>Prepare a syringe with sterile BSS using a sterile 26 G 1/2 needle. Fit the syringe with a sterile 27 G 45&#176; bent blunt dispensing tip of 1 inch (referred to as a dispensing tip below).</li>
</ol>
4. Anesthesia
<ol>
	<li>Following eye drops, anesthetize the mouse with an intraperitoneal injection of xylazine/ketamine/acepromazine (35 mg/kg, 80 mg/kg, and 0.5 mg/kg) solution (Figure 1B). 
	NOTE: Inhaled anesthesia might not be possible unless a specialized nose cone can be utilized to allow for access to the eye needed for surgery.</li>
</ol>
5. Making the incision
<ol>
	<li>Once the mouse is completely under anesthesia, as measured by the lack of toe pinch reflex, and its pupil has dilated completely (2 mm) (Figure 4A), the mouse is ready for surgery.
	<ol>
		<li>Move the anesthetized mouse to the stage of a compound dissecting light microscope. Use a heater pad to ensure the mouse has adequate thermal regulation and use sterile drapes.</li>
		<li>View the eye under 10x-20x magnification using a dissecting microscope (see Table of Materials for specifics).</li>
	</ol>
	</li>
	<li>Make a 1-1.5 mm incision in the center of the cornea using a sterile surgical scalpel. Ensure the incision extends through the cornea and incises the anterior lens capsule (Figure 1C and Figure 4B).</li>
</ol>
6. Separating the lens fibers from the capsular bag
<ol>
	<li>Grasp the syringe with a dispensing tip containing sterile BSS and gently expel 1 drop of BSS above the cornea to wet the surface for further manipulation.</li>
	<li>Separate the lens fiber mass from the lens capsule.
	<ol>
		<li>Insert the dispensing tip through the central corneal incision into the lens capsule incision, then gently move the needle in a circular motion while slowly releasing BSS to separate apical-apical connections between lens epithelial and fiber cells (hydro dissection), ultimately expelling the lens fiber mass (Figure 1D).</li>
		<li>In most cases, the fiber mass will eventually adhere to the dispensing tip during this procedure (Figure 2A). If this happens, gently remove the dispensing tip from the inside eye; and clean with a sterile towel to remove this tissue. Repeat until all lens fiber cells are removed.</li>
		<li>Alternatively, use micro forceps (number 5 Dumont) to extract the lens nucleus (Figure 2B). If using this method, be careful that the original corneal incision exceeds the diameter of the nucleus to ensure easy removal.</li>
	</ol>
	</li>
	<li>Rinse the lens capsule with sterile BSS to remove any remaining fibers.</li>
	<li>Confirm that the lens capsule is left inside the eye (Figure 1E and Figure 4C) by viewing its translucent/glassy appearance through the microscope.</li>
</ol>
7. Inflating the anterior chamber and suturing the corneal incision
<ol>
	<li>Inflate the anterior chamber to its normal depth by injecting BSS through the incision into the anterior chamber.
	<ol>
		<li>Add inhibitors or activators of signaling pathways of interest to the BSS to test their roles in the lens injury response12,15. 
		NOTE: Animals can also be treated systemically with pathway inhibitors, which may be needed depending on the pharmacokinetics of drug turnover as the aqueous humor balance is re-established after surgery15.</li>
	</ol>
	</li>
	<li>Suture the corneal incision with square knots using 10-0 nylon thread&#160;with a tapered, curved 5.3 mm 1/2c needle (Figure 1F and Figure 4D-F).
	<ol>
		<li>Place either one square knot stitch in the central part of the incision or two square knot stitches at both the upper and lower halves of the original incision (see Figure 3). Number of stitches depends on the length of the corneal incision.</li>
		<li>Do not puncture both corneal flaps at one time. Instead, hold one corneal flap using forceps in the non-dominant hand and push the needle/nylon through. Once one side of the cornea has been punctured, push the needle/nylon through the next corneal flap. Close the corneal flaps using square knot stitches.</li>
	</ol>
	</li>
</ol>
8. Post-surgical care
NOTE: Topical antibiotic ointment is applied to the eye, and analgesic agents are administered. Mice do not&#160;usually show signs of distress after initial surgery. If needed, slow-release version of buprenorphine could be used.
<ol>
	<li>Immediately following this surgery, do the following sub-steps.
	<ol>
		<li>Apply one drop of 0.5% erythromycin ophthalmic ointment directly above the suture. Make sure the entire incision is covered in ointment.</li>
		<li>Administer an intraperitoneal injection of buprenorphine (0.1 mg/kg).</li>
	</ol>
	</li>
	<li>Allow the mouse to recover from anesthesia in a cage placed on a heating pad to optimize thermal support.&#160; 
	NOTE: Monitor the animal for signs of trauma, distress, or discomfort. Some signs include infection/scratching at the surgical site, displaced suture, loss of fur, etc.</li>
</ol>
9. Monitoring animal recovery
<ol>
	<li>Monitor the animal&#39;s recovery after surgery until it awakes from anesthesia.</li>
	<li>Check mice every day for the first 3 days following surgery. 
	NOTE: It is usually not needed to remove stitches because mice are either sacrificed prior to full healing (2 weeks PCS) or stitches spontaneously fall out.</li>
	<li>Administer buprenorphine orally, as needed for pain management after 4-8 h post-surgery, at a dosage of 0.5 mg/kg (in Jello) or a dosage of 0.1 mg/kg for those not eating. Alternatively, administer buprenorphine extended-release just prior to surgery. 
	NOTE: When possible, preemptive analgesics should be utilized, yet this may require adjustments to the analgesic protocol that require review and approval from the institutional animal care and use committee.</li>
</ol>
10. Euthanasia
<ol>
	<li>Euthanize the mouse at a desired time post-surgery
	<ol>
		<li>Removed the operated eye using the unoperated eye as control tissue; or re-anesthetize the animal, perform lens fiber cell removal surgery on the previously unoperated contralateral eye, then immediately sacrifice the animal (without allowing it to awake from anesthesia) to provide a time zero operated control.</li>
		<li>Ensure quick dissection and tissue harvest. 
		NOTE: LECs undergo transcriptomic changes within minutes of surgery so rapid tissue harvest and fixation/freezing are necessary when studying early LEC responses to surgery.</li>
	</ol>
	</li>
</ol>
11. Performing further analysis on harvested lens tissues
<ol>
	<li>Use the obtained tissue for immunostaining, RNA sequencing, cell culture, or&#160;western analysis12,15,23,25,26,28. Flow cytometry is theoretically possible as well12, although the small number of LECs in the mouse lens (~40,000)29 and the presence of capsule associated cell debris following surgery make this challenging.</li>
	<li>Immunostaining
	<ol>
		<li>Euthanize the mouse in a chamber via CO2 inhalation (4 L/min) from a gas cylinder. Increase the flow rate to 8-10 L/min once the mouse is unconscious and continue this exposure until complete cessation of breathing plus 1 min. 
		NOTE: The mouse can be euthanized using an appropriate method per AVMA Guidelines for the Euthanasia of Animals or per Institutional Animal Care and Use Committee approved guidelines or policies.</li>
		<li>Ensure proper sacrifice by employing a secondary method of euthanasia, e.g., cervical dislocation. 
		NOTE: The Duncan lab has investigated time points ranging from minutes to months PCS12,15,23.</li>
		<li>After the mouse has been humanely sacrificed, isolate the operated eye for analysis.
		<ol>
			<li>Hold micro forceps in the non-dominant hand to grasp the operated eye while holding micro scissors in the dominant hand.</li>
			<li>Gently clip the optic nerve and muscles anchoring the eye. 
			NOTE: Be particularly careful handling eyes that were recently operated on versus those that have had longer to heal PCS. Recently operated eyes are more structurally fragile.</li>
		</ol>
		</li>
		<li>Embed the dissected eye in OCT media for later cryo-sectioning, immunostaining, and semi-quantitative confocal microscopy28,30.</li>
	</ol>
	</li>
	<li>Isolation of the capsular bag with associated cells for RNA sequencing, cell culture, or western blot analysis
	<ol>
		<li>Euthanize the mouse in a chamber via CO2 inhalation (4 L/min) from a gas cylinder. Increase the flow rate to 8-10 L/min once the mouse is unconscious and continue this exposure until complete cessation of breathing plus 1 min.</li>
		<li>Ensure proper sacrifice by employing a secondary method of euthanasia, e.g., cervical dislocation.</li>
		<li>If an early timepoint PCS (within 1-2 days) is required, either remove stitches prior to tissue extraction or make another incision in the cornea through which to extract the lens capsule. 
		NOTE: At later time points PCS, the stitches will likely have fallen out, so a new incision will be needed.</li>
	</ol>
	</li>
	<li>Locate the translucent lens capsule.
	<ol>
		<li>Gently insert the micro forceps through the corneal opening, grasp the capsule, and remove it from the anterior chamber.</li>
		<li>Place the capsule in a 1.5 mL microcentrifuge tube or culture dish for further analysis.</li>
	</ol>
	</li>
</ol>

Replicating Cataract Surgical Techniques in a Mouse Model Through LEC Extraction

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The Duncan lab has received financial support from Pliantx to test anti-PCO therapies using this surgery model.

This surgical technique requires advanced mouse handling and micro-surgical skills that take practice to develop. The most difficult to master is the placement of fine sutures in the cornea to close the wound. Suppose the experimenter is a total novice to suturing. In that case, it is recommended to first practice using string and cloth, then move to practice using large-diameter sutures on small fruits to master the needed hand movements to make surgical square knots. Then, the experimenter must practice making incision...

The lens is a highly organized, transparent tissue that refracts light to produce a clearly focused image onto the retina1,2,3. This specialized organ is surrounded by an uninterrupted basement membrane (the capsule), which isolates the lens from other parts of the eye. The inner anterior surface of the capsule anchors a monolayer of lens epithelial cells (LECs), which then differentiate at the lens equator into lens fiber cells, which comprise the vast majority of the lens3. A cataract occurs when the lens loses its transparency due to factors like aging, genetic mutation, UV radiation, oxidative stress, and ocular trauma4. Cataract is a leading cause of blindness worldwide, particularly in countries with poor medical care5. However, this condition is now readily treated by surgical removal of opaque lens cells by phacoemulsification in which the central lens capsule and attached lens epithelium are removed, followed by the use of a vibrating probe to break the cellular mass of the lens into smaller fragments that can be suctioned out; leaving behind a capsular bag with some attached equatorial LECs1. Vision is then most commonly restored by post-surgical implantation of an artificial intraocular lens (IOL).
While cataract surgery (CS) is a highly effective and minimally invasive treatment for cataracts, a patient&#39;s post-surgical recovery can be acutely hampered by the development of ocular inflammation5,6,8. This inflammation can cause post-surgical pain, retinal edema leading to retinal detachment, as well as exacerbation of other inflammatory and fibrotic conditions like uveitis and glaucoma4,7,8,9. Ocular inflammation post-CS (PCS) is generally treated with anti-inflammatory eye drops which are plagued by poor patient compliance or dropless cataract surgery that can lead to an increased prevalence of floaters8,10,11. In the long term, the outcomes of CS can be compromised by the development of posterior capsular opacification (PCO)12. PCO occurs when residual LECs left behind after surgery undergo a wound healing response, proliferate, and migrate onto the posterior lens capsule at&#160;months to years PCS contributing to secondary visual obstruction13,14.
Understanding the molecular mechanisms by which CS results in such acute and chronic responses represents a major challenge in the field, as much of the literature investigating PCO relies on the induction of LEC conversion to myofibroblasts in culture via treatment with active transforming growth factor beta (TGF&#946;)12,15. Human capsular bag models created from cadaver lenses cultured in vitro better reflect the biology of the lens PCS as LECs are cultured on their native basement membrane but are difficult to manipulate mechanistically, do not recapitulate the intraocular environment, and are inherently variable due to lens to lens (or donor to donor) variability16,17. Rabbit1,18 and non-human primate19,20&#160;in vivo models of cataract surgery overcome some of these issues but still are not easy to manipulate for mechanistic studies. Notably, a prior study of mammalian lens regeneration found significant lens regeneration within four weeks after lens fiber cells were removed from mice, leaving the lens epithelial cells and capsule behind, while no lens regrowth occurred when the entire lens was removed21,22. We subsequently streamlined this procedure and optimized it for the study of the acute response of LECs to lens fiber cell removal and their subsequent conversion to a mixed population of cells with myofibroblast and fiber cell properties.
Using this mouse model of cataract surgery, we have shown that lens epithelial cells drastically remodel their transcriptome by 6 hours PCS to produce numerous proinflammatory cytokines23, while they begin expressing fibrotic markers as early as 24 h PCS12,23, prior to the onset of canonical TGF&#946; signaling. Mechanistic experiments in knockout mice using this model revealed that cellular fibronectin expression by LECs is required for a sustained fibrotic response PCS, likely due to both its role in the assembly of the fibrotic ECM and cell signaling12. Other studies showed that &#945;V&#946;8-integrin is required for the transition of LECs to myofibroblasts due to its function in TGF&#946; activation, and the potential of this approach to identify anti-PCO therapeutic leads was confirmed as a &#945;V&#946;8-integrin antagonist also blocked LEC EMT15.
Here, we provide a detailed protocol describing how to remove the lens fiber cells from living mice (Figure 1and&#160;Figure 4) while leaving behind the lens capsule and LECs to model the LEC response to cataract surgery.&#160;

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modeling cataract surgery in mice

Cataract surgery (CS) is an effective treatment for cataracts, a major cause of visual disability worldwide. However, CS leads to ocular inflammation, and in the long term, it can result in posterior capsular opacification (PCO) and/or lens dislocation driven by the post-surgical overgrowth of lens epithelial cells (LECs) and their conversion to myofibroblasts and/or aberrant fiber cells. However, the molecular mechanisms by which CS results in inflammation and PCO are still obscure because most in vitro models do not recapitulate the wound healing response of LECs seen in vivo, while traditional animal models of cataract surgery, such as rabbits, do not allow the genetic manipulation of gene expression to test mechanisms. Recently, our laboratory and others have successfully used genetically modified mice to study the molecular mechanisms that drive the induction of proinflammatory signaling and LEC epithelial to mesenchymal transition, leading to new insight into PCO pathogenesis. Here, we report the established protocol for modeling cataract surgery in mice, which allows for robust transcriptional profiling of the response of LECs to lens fiber cell removal via RNAseq, the evaluation of protein expression by semi-quantitative immunofluorescence, and the use of modern mouse genetics tools to test the function of genes that are hypothesized to participate in the pathogenesis of acute sequelae like inflammation as well as the later conversion of LECs to myofibroblasts and/or aberrant lens fiber cells.

Based on the results from this surgical method, the Duncan Lab has used this mouse model of cataract surgery to construct an injury response time course of lens epithelial cells post-surgery (Figure 5). By 6 h after surgery, 5% of the lens epithelial transcriptome is differentially expressed (Figure 6A), including the upregulation of numerous immediate early response genes and proinflammatory cytokines23 (Figure 6B</s...

Watch this Scientific Journal Video about Unraveling Molecular Mechanisms in PCO and Fibrosis Following Cataract Surgery at JoVE.com

Author Spotlight: Unraveling the Molecular Mechanisms in PCO and Fibrosis Following Cataract Surgery

This method models cataract surgery in vivo by removing lens fiber cells from adult mice and leaving behind the capsular bag with attached lens epithelial cells (LECs). The injury response is then assessed at various times post-surgery using molecular and morphological criteria.

Modeling Cataract Surgery in Mice

Cataract surgery (CS) is an effective treatment for cataracts, a major cause of visual disability worldwide. However, CS leads to ocular inflammation, and ...

Research

JoVE Journal

Biology

837 Views.  University of Delaware. This method models cataract surgery in vivo by removing lens fiber cells from adult mice and leaving behind the capsular bag with attached lens epithelial cells (LECs). The injury response is then assessed at various times post-surgery using molecular and morphological criteria. 

Video: Author Spotlight: Unraveling the Molecular Mechanisms in PCO and Fibrosis Following Cataract Surgery

Heterotopic Heart Transplantation in Mice

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable for studying rejection and immune response now that newer transgenic and gene knockout mice are available, and a large number of immunologic reagents have been developed. The heart transplant model is less stringent than the skin transplant models, although technically more challenging. We have developed a modified technique and have completed over 1000 successful cases of heterotopic heart transplantation in mice. When making anastomosis of the ascending aorta and abdominal aorta, two stay sutures are placed at the proximal and distal apexes of recipient abdominal aorta with the donor s ascending aorta, then using 11-0 suture for anastomosis on both side of aorta with continuing sutures. The stay sutures make the anastomosis easier and 11-0 is an ideal suture size to avoid bleeding and thrombosis.
When making anastomosis of pulmonary artery and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s pulmonary artery. The left wall of the inferior vena cava and donor s pulmonary artery is closed with continuing sutures in the inside of the inferior vena cava after, one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s pulmonary artery are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis. In this article, we provide details of the technique to supplement the video.

The mouse heterotopic heart transplantation model has been proven by many investigators to be an important method for studying mechanisms of rejection and immune response. However, the techniques involved are still challenging. By modifying standard techniques we have had success with more than 1000 transplants.

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable ...

Small Bowel Transplantation In Mice

Since 1990, the development of tacrolimus-based immunosuppression and improved surgical techniques, the increased array of potent immunosuppressive medications, infection prophylaxis, and suitable patient selection helped improve actuarial graft and patient survival rates for all types of intestine transplantation. Patients with irreversible intestinal failure and complications of parenteral nutrition should now be routinely considered for small intestine transplantation. However, Survival rates for small intestinal transplantation have been slow to improve compares increasingly favorably with renal, liver, heart and lung. The small bowel transplantation is still unsatisfactory compared with other organs. Further progress may depend on better understanding of immunology and physiology of the graft and can be greatly facilitated by animal models. A wider use of mouse small bowel transplantation model is needed in the study of immunology and physiology of the transplantation gut as well as efficient methods in diagnosing early rejection. However, this model is limited to use because the techniques involved is an extremely technically challenging. We have developed a modified technique. When making anastomosis of portal vein and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s portal vein. The left wall of the inferior vena cava and donor s portal vein is closed with continuing sutures in the inside of the inferior vena cava after, after one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s portal vein are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis.  In this article, we provide details of the technique to supplement the video.

The mouse small bowel transplantation model has been recognized as an important tool to study mechanismes of immune rejection and screen new immunosuppressive drugs. However, this model is limited to use because the techniques involved is an extremely technically challenge. Now we introduce the modified technique.

Since 1990, the development of tacrolimus-based immunosuppression and improved surgical techniques, the increased array of potent immunosuppressive ...

Survivable Stereotaxic Surgery in Rodents 

The ability to measure extracellular basal levels of neurotransmitters in the brain of awake animals allows for the determination of effects of different systemic challenges (pharmacological or physiological) to the CNS. For example, one can directly measure how the animal's midbrain dopamine projections respond to dopamine-releasing drugs like d-amphetamine or natural stimuli like food. In this video, we show you how to implant guide cannulas targeting specific sites in the rat brain, how to insert and implant a microdialysis probe and how to use high performance liquid chromatography coupled with electrochemical detection (HPLC-EC) to measure extracellular levels of oxidizable neurotransmitters and metabolites. Local precise introduction of drugs through the microdialysis probe allows for refined work on site specificity in a compound s mechanism of action. This technique has excellent anatomical and chemical resolution but only modest time resolution as microdialysis samples are usually processed every 20-30 minutes to ensure detectable neurotransmitter levels. Complementary ex vivo tools (i.e., slice and cell culture electrophysiology) can assist with monitoring real-time neurotransmission.

Survivable Stereotaxic Surgery in Rodents

The monitoring of extracellular neurotransmitter levels in distinct brain regions of freely moving animals offers insights on the link between neurotransmitter release and behavior. In vivo microdialysis coupled with electrochemical detection provides excellent anatomical and chemical resolution; and information on how basal neurotransmission is altered by pharmacological or physiological manipulations.

The ability to measure extracellular basal levels of neurotransmitters in the brain of awake animals allows for the determination of effects of different ...

Computer-assisted Large-scale Visualization and Quantification of Pancreatic Islet Mass, Size Distribution and Architecture

The pancreatic islet is a unique micro-organ composed of several hormone secreting endocrine cells such as beta-cells (insulin), alpha-cells (glucagon), and delta-cells (somatostatin) that are embedded in the exocrine tissues and comprise 1-2% of the entire pancreas. There is a close correlation between body and pancreas weight. Total beta-cell mass also increases proportionately to compensate for the demand for insulin in the body. What escapes this proportionate expansion is the size distribution of islets. Large animals such as humans share similar islet size distributions with mice, suggesting that this micro-organ has a certain size limit to be functional. The inability of large animal pancreata to generate proportionately larger islets is compensated for by an increase in the number of islets and by an increase in the proportion of larger islets in their overall islet size distribution. Furthermore, islets exhibit a striking plasticity in cellular composition and architecture among different species and also within the same species under various pathophysiological conditions. In the present study, we describe novel approaches for the analysis of biological image data in order to facilitate the automation of analytic processes, which allow for the analysis of large and heterogeneous data collections in the study of such dynamic biological processes and complex structures. Such studies have been hampered due to technical difficulties of unbiased sampling and generating large-scale data sets to precisely capture the complexity of biological processes of islet biology. Here we show methods to collect unbiased "representative" data within the limited availability of samples (or to minimize the sample collection) and the standard experimental settings, and to precisely analyze the complex three-dimensional structure of the islet. Computer-assisted automation allows for the collection and analysis of large-scale data sets and also assures unbiased interpretation of the data. Furthermore, the precise quantification of islet size distribution and spatial coordinates (i.e. X, Y, Z-positions) not only leads to an accurate visualization of pancreatic islet structure and composition, but also allows us to identify patterns during development and adaptation to altering conditions through mathematical modeling. The methods developed in this study are applicable to studies of many other systems and organisms as well.

Novel computer-assisted methods of large-scale procurement and analysis of immunohistochemically stained pancreatic specimens are described: (1) Virtual Slice capture of the entire section; (2) Mass analysis of large-scale data; (3) Reconstruction of 2D Virtual Slices; (4) 3D islet mapping; and (5) Mathematical analysis.

The pancreatic islet is a unique micro-organ composed of several hormone secreting endocrine cells such as beta-cells (insulin), alpha-cells (glucagon), ...

Bioengineering Human Microvascular Networks in Immunodeficient Mice

The future of tissue engineering and cell-based therapies for tissue regeneration will likely rely on our ability to generate functional vascular networks in vivo. In this regard, the search for experimental models to build blood vessel networks in vivo is of utmost importance 1. The feasibility of bioengineering microvascular networks in vivo was first shown using human tissue-derived mature endothelial cells (ECs) 2-4; however, such autologous endothelial cells present problems for wide clinical use, because they are difficult to obtain in sufficient quantities and require harvesting from existing vasculature. These limitations have instigated the search for other sources of ECs. The identification of endothelial colony-forming cells (ECFCs) in blood presented an opportunity to non-invasively obtain ECs 5-7. We and other authors have shown that adult and cord blood-derived ECFCs have the capacity to form functional vascular networks in vivo 7-11. Importantly, these studies have also shown that to obtain stable and durable vascular networks, ECFCs require co-implantation with perivascular cells. The assay we describe here illustrates this concept: we show how human cord blood-derived ECFCs can be combined with bone marrow-derived mesenchymal stem cells (MSCs) as a single cell suspension in a collagen/fibronectin/fibrinogen gel to form a functional human vascular network within 7 days after implantation into an immunodeficient mouse. The presence of human ECFC-lined lumens containing host erythrocytes can be seen throughout the implants indicating not only the formation (de novo) of a vascular network, but also the development of functional anastomoses with the host circulatory system. This murine model of bioengineered human vascular network is ideally suited for studies on the cellular and molecular mechanisms of human vascular network formation and for the development of strategies to vascularize engineered tissues.

Here, we describe a methodology to deliver human cord blood-derived endothelial colony-forming cells (ECFCs) and bone marrow-derived mesenchymal stem cells (MSCs), embedded in a collagen/fibronectin gel, subcutaneously into immunodeficient mice. This cell/gel combination generates a human vascular network that connects with the mouse vasculature.

The future of tissue engineering and cell-based therapies for tissue regeneration will likely rely on our ability to generate functional vascular networks ...

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present methods for the development of assays to query potentially clinically significant nonsynonymous changes using in vivo complementation in zebrafish. Zebrafish (Danio rerio) are a useful animal system due to their experimental tractability; embryos are transparent to enable facile viewing, undergo rapid development ex vivo, and can be genetically manipulated.1 These aspects have allowed for significant advances in the analysis of embryogenesis, molecular processes, and morphogenetic signaling. Taken together, the advantages of this vertebrate model make zebrafish highly amenable to modeling the developmental defects in pediatric disease, and in some cases, adult-onset disorders. Because the zebrafish genome is highly conserved with that of humans (~70% orthologous), it is possible to recapitulate human disease states in zebrafish. This is accomplished either through the injection of mutant human mRNA to induce dominant negative or gain of function alleles, or utilization of morpholino (MO) antisense oligonucleotides to suppress genes to mimic loss of function variants. Through complementation of MO-induced phenotypes with capped human mRNA, our approach enables the interpretation of the deleterious effect of mutations on human protein sequence based on the ability of mutant mRNA to rescue a measurable, physiologically relevant phenotype. Modeling of the human disease alleles occurs through microinjection of zebrafish embryos with MO and/or human mRNA at the 1-4 cell stage, and phenotyping up to seven days post fertilization (dpf). This general strategy can be extended to a wide range of disease phenotypes, as demonstrated in the following protocol. We present our established models for morphogenetic signaling, craniofacial, cardiac, vascular integrity, renal function, and skeletal muscle disorder phenotypes, as well as others.

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present a systematic approach for developing physiologically relevant, sensitive and specific in vivo assays for interpreting variation in human pathology. Transient genetic manipulation via microinjection of WT and mutant human mRNA and morpholino (MO) antisense oligonucleotides harness the tractability of the developing zebrafish embryo to rapidly assay pathogenic mutations, especially, but not exclusively, in the context of human developmental disorders.

Here,  we present methods for the development of assays to query potentially clinically  significant nonsynonymous changes using in  vivo complementation ...

Modeling Age-Associated Neurodegenerative Diseases in Caenorhabditis elegans

Battling human neurodegenerative pathologies and managing their pervasive socioeconomic impact is becoming a global priority. Notwithstanding their detrimental effects on the human life quality and the healthcare system, the majority of human neurodegenerative disorders still remain incurable and non-preventable. Therefore, the development of novel therapeutic interventions against such maladies is becoming a pressing urgency. Age-associated deterioration of neuronal circuits and function is evolutionarily conserved in organisms as diverse as the lowly worm Caenorhabditis elegans and humans, signifying similarities in the underlying cellular and molecular mechanisms. C. elegans is a highly malleable genetic model, which offers a well-characterized nervous system, body transparency and a diverse repertoire of genetic and imaging techniques to assess neuronal activity and quality control during ageing. Here, we introduce and describe methodologies utilizing some versatile nematode models, including hyperactivated ion channel-induced necrosis (e.g., deg-3(d) and mec-4(d)) and protein aggregate (e.g., &#945;-syunclein and poly-glutamate)-induced neurotoxicity, to monitor and dissect the cellular and molecular underpinnings of age-related neuronal breakdown. A combination of these animal neurodegeneration models, together with genetic and pharmacological screens for cell death modulators will lead to an unprecedented understanding of age-related breakdown of neuronal function and will provide critical insights with broad relevance to human health and quality of life.

Modeling Age-Associated Neurodegenerative Diseases in Caenorhabditis elegans

Here, we introduce and describe widely accessible methodologies utilizing some versatile nematode models, including hyperactivated ion channel-induced necrosis and protein aggregate-induced neurotoxicity, to monitor and dissect the cellular and molecular underpinnings of age-associated neurodegenerative diseases.

Battling human neurodegenerative pathologies and managing their pervasive socioeconomic impact is becoming a global priority. Notwithstanding their ...

A Modified Surgical Model of Hind Limb Ischemia in ApoE-/- Mice using a Miniature Incision

The purpose of this study is to introduce and evaluate a modified surgical approach to induce acute ischemia in mice that can be implemented in most animal laboratories.&#160;Contrary to the conventional approach for double ligation of the femoral artery (DLFA), a smaller incision on the right inguinal region was made to expose the proximal femoral artery (FA) to perform DLFA. Then, using a 7-0 suture, the incision was dragged to the knee region to expose the distal FA. Magnetic resonance imaging (MRI) on bilateral hind limbs was used to detect FA occlusion after the surgery. At 0, 1, 3, 5, and 7 days after the surgery, functional recovery of the hind limbs was visually assessed and graded using the Tarlov scale. Histologic evaluation was performed after euthanizing the animals 7 days after DLFA. The procedures were successfully performed on the right leg in ten ApoE-/- mice, and no mice died during subsequent observation. The incision sizes in all 10 mice were less than 5 mm (4.2 &#177; 0.63 mm). MRI results showed that FA blood flow in the ischemic side was clearly blocked. The Tarlov scale results demonstrated that hind limb function significantly decreased after the procedure and slowly recovered over the following 7 days. Histologic evaluation showed a significant inflammatory response on the ischemic side and reduced microvascular density in the ischemic hind limb. In conclusion, this study introduces a modified technique using a miniature incision to perform hind limb ischemia (HLI) using DLFA.

A Modified Surgical Model of Hind Limb Ischemia in ApoE-/- Mice using a Miniature Incision

This article demonstrates an efficient surgical approach to establish acute ischemia in mice with a small incision. This approach can be applied by most research groups without any laboratory upgrades.

The purpose of this study is to introduce and evaluate a modified surgical approach to induce acute ischemia in mice that can be implemented in most ...

Isolation of Human Primary Valve Cells for In vitro Disease Modeling

Calcific aortic valve disease (CAVD) is present in nearly a third of the elderly population. Thickening, stiffening, and calcification of the aortic valve causes aortic stenosis and contributes to heart failure and stroke. Disease pathogenesis is multifactorial, and stresses such as inflammation, extracellular matrix remodeling, turbulent flow, and mechanical stress and strain contribute to the osteogenic differentiation of valve endothelial and valve interstitial cells. However, the precise initiating factors that drive the osteogenic transition of a healthy cell into a calcifying cell are not fully defined. Further, the only current therapy for CAVD-induced aortic stenosis is aortic valve replacement, whereby the native valve is removed (surgical aortic valve replacement, SAVR) or a fully collapsible replacement valve is inserted via a catheter (transcatheter aortic valve replacement, TAVR). These surgical procedures come at a high cost and with serious risks; thus, identifying novel therapeutic targets for drug discovery is imperative. To that end, the present study develops a workflow where surgically removed tissues from patients and donor cadaver tissues are used to create patient-specific primary lines of valvular cells for in vitro disease modeling. This protocol introduces the utilization of a cold storage solution, commonly utilized in organ transplant, to reduce the damage caused by the often-lengthy procurement time between tissue excision and laboratory processing with the benefit of greatly stabilizing cells of the excised tissue. The results of the present study demonstrate that isolated valve cells retain their proliferative capacity and endothelial and interstitial phenotypes in culture upwards of several days after valve removal from the donor. Using these materials allows for the collection of control and CAVD cells, from which both control and disease cell lines are established.

This protocol describes the collection of human aortic valves extracted during surgical aortic valve replacement procedures or from cadaveric tissue, and the subsequent isolation, expansion, and characterization of patient specific primary valve endothelial and interstitial cells. Included are important details regarding the processes needed to ensure cell viability and phenotype specificity.

Calcific aortic valve disease (CAVD) is present in nearly a third of the elderly population. Thickening, stiffening, and calcification of the aortic valve ...

Establishing a Diaphyseal Femur Fracture Model in Mice

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age and overall health status of the patient, failures can occur, leading to delayed union or nonunion. Due to the increasing number of fractures resulting from high-energy trauma and aging, the development of innovative therapeutic strategies to improve bone repair based on the combination of skeletal/mesenchymal stem/stromal cells and biomimetic biomaterials is urgently needed. To this end, the use of reliable animal models is fundamental to better understanding the key cellular and molecular mechanisms that determine the healing outcomes. Of all the models, the mouse is the preferred research model because it offers a wide variety of transgenic strains and reagents for experimental analysis. However, the establishment of fractures in mice may be technically challenging due to their small size. Therefore, this article aims to demonstrate the procedures for the surgical establishment of a diaphyseal femur fracture in mice, which is stabilized with an intramedullary wire and resembles the most common bone repair process, through cartilaginous callus formation.

This protocol describes a surgical procedure for the establishment of a diaphyseal fracture in the femur of mice, which is stabilized with an intramedullary wire, for fracture healing studies.

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age ...

Modeling Cataract Surgery in Mice

Summary

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Chapters in this video

Heterotopic Heart Transplantation in Mice

Small Bowel Transplantation In Mice

Survivable Stereotaxic Surgery in Rodents

Computer-assisted Large-scale Visualization and Quantification of Pancreatic Islet Mass, Size Distribution and Architecture

Bioengineering Human Microvascular Networks in Immunodeficient Mice

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Modeling Age-Associated Neurodegenerative Diseases in Caenorhabditis elegans

A Modified Surgical Model of Hind Limb Ischemia in ApoE^-/- Mice using a Miniature Incision

Isolation of Human Primary Valve Cells for In vitro Disease Modeling

Establishing a Diaphyseal Femur Fracture Model in Mice