This work was supported by funds from Midwestern University College of Health Sciences Research Facilitation Grant awarded to EEH and Midwestern University Office of Research and Sponsored Programs Intramural Grant awarded to KJL.

This protocol conforms to the AVMA Animal Welfare Principles for the care and use of laboratory animals and has been approved by the Institutional Animal Care and Use Committee (IACUC) at Midwestern University.
1. Preparation of Anesthesia, Equipment, &#38; Media
<ol>
	<li>Dechlorinate approximately 1.5 L of tap water using either a commercial dechlorinating agent (available at pet stores) or by letting water stand for 24 hr. Obtain three 1 L beakers and fill each with 500 ml dechlorinated water. Label one for holding fish before any procedure and one for recovery. To the third beaker, add 100 &#181;g/ml of tricaine methanesulfonate; this becomes the beaker in which fish will be anesthetized.</li>
	<li>Fill a shallow tray with ice, warm the culture medium (RPMI 1640, 10% FBS (fetal bovine serum), 50 &#181;g/ml gentamycin, 100 &#181;g/ml kanamycin, 25 mM HEPES) to RT, and gather clean jeweler&#8217;s tweezers. Clean tweezers by immersing the tips in 70% ethanol and pass through a Bunsen burner, allowing the ethanol to burn off.</li>
	<li>Decide if the procedure will be performed without magnification, with a commercial, table-top lighted magnifying glass with 2 - 10X magnification, or dissecting microscope (entirely dependent on personal preference).</li>
</ol>
2. Establish Explant Cultures
<ol>
	<li>Catch the desired number of fish and place in a holding beaker. Transfer a single fish to the anesthesia beaker; it is best to anesthetize fish individually.</li>
	<li>Monitor the effects of anesthesia by observing the swimming and gill movement of the fish. As anesthesia progresses, gill movement slows and the fish will swim erratically and frequently upside down; consider fish anesthetized as soon as gill movement ceases. Do not leave the fish for longer periods in the anesthesia bath as this may result in the death of the fish.</li>
	<li>Remove fish from the anesthesia beaker and transfer to ice, placing the fish horizontally with the tail facing the hand holding the tweezers. If experienced with the technique and if multiple fish are to be used to make explants, consider placing the next fish in the anesthesia beaker at this time.</li>
	<li>To remove scales, position tweezers parallel to the fish with tip at edge of scale. Depress flat side of the tweezers slightly into the side of the fish to cause the scale to elevate from the body of the fish. Grasp scale with tweezers, pluck, and place (without inversion) in tissue culture dish. Repeat the procedure, removing a maximum of 12 scales from a large adult (6 from each side), avoiding the gill region, the lateral line, and fins.</li>
	<li>Place the fish into the recovery beaker and monitor to ensure it recovers from the effects of anesthesia. At this time, fish can be transferred back to an individual tank. Allow the fish to recover for at least 21 days to allow for complete scale regeneration.</li>
	<li>Depending on the experimental design, place up to 20 scales per plastic 35 mm tissue culture-treated dish or 4 - 6 scales per glass bottom dish. Allow scales to adhere to the dish before gently adding media so as not to detach scales from the dish. 
	NOTE: With experience, multiple fish can be processed before media is added to the first dish.</li>
	<li>Add 1.2 ml of complete media (see 1.2 above) into each 35 mm dish.</li>
	<li>Preferably incubate explant cultures at 28 &#176;C in 5% CO2 for the desired period of time with or without additional treatment. Alternatively, incubate cultures on a heated microscope stage or within a live cell culture chamber for video microscopy.</li>
</ol>
3. Brightfield and Video Microscopy
<ol>
	<li>Position dishes containing cell sheets onto the microscope stage and initially focus using the 4X objective lens. 
	NOTE: Higher magnifications may be used, depending on the experiment. Marking the location of each cell sheet and/or orientation of the dish on the stage will facilitate taking images over time with the sheets in approximately the same orientation.</li>
	<li>Remove dishes from the incubator and photograph at various time points, according to each experimental protocol and place back into the incubator if only still images are needed over the experimental time frame.</li>
	<li>While performing video microscopy, pay attention to the following:
	<ol>
		<li>Position the scale/emerging cell sheet within the field of view such that the migrating cell sheet remains in the field of view for the duration of the video. 
		NOTE: This may take some experience in observing how cells migrate from under the scale. For short videos, this is less of a concern than for longer videos.</li>
		<li>For shorter time frames, incubate at RT. For longer time frames (18+ hr), use a heated stage or an incubation chamber to maintain temperature, pH balance, and to minimize evaporation of the culture medium.</li>
	</ol>
	</li>
</ol>
4. Fixation for Immunofluorescence Microscopy
<ol>
	<li>Prepare and pre-warm all solutions to 28 &#176;C. In order to view keratocytes under fluorescence, grow explant cultures as described above using glass-bottom 35 mm tissue culture dishes. Alternatively, use glass chamber slides.</li>
	<li>Add 0.8 ml of 4% paraformaldehyde in 1.1x PBS to each dish. Do not remove the culture media first. Incubate at RT for 5 min then aspirate to remove the solution.</li>
	<li>Add 0.8 ml of 4% paraformaldehyde in 1.1x PBS to each dish. Incubate at RT for 5 min then aspirate to remove the solution.</li>
	<li>Unless using external ligands or external epitopes, permeabilize cells by adding 0.5 ml of 0.2% Triton X-100 in 1X PBS. Incubate at RT for 5 min then aspirate to remove the solution.</li>
	<li>Add 0.5 ml of 1% BSA in 1x PBS and incubate for 1 hr (at RT) or O/N (at 4 &#176;C). 
	NOTE: Depending on experimental conditions, phalloidin can be added at this step.</li>
	<li>After incubation, aspirate to remove the solution and proceed with antibody staining. Alternatively, store the culture dishes by adding at least 1 - 2 ml of 1% BSA in 1x PBS for 3 - 4 weeks at 4 &#176;C.</li>
</ol>
5. Immunofluorescence Protocol
<ol>
	<li>Prepare primary and secondary antibody solutions according to the manufacturer&#8217;s recommendations. 
	NOTE: If manufacturer recommendations are not available, a good starting dilution of primary antibodies (in 1% BSA in 1x PBS) is 1:500 for polyclonal and 1:1,000 for monoclonal antibodies; a good starting dilution for secondary antibodies is 1:1,000. Adjust these dilutions of the original stock obtained from the manufacturer higher if the signal is weak, lower if the signal is strong and background is a problem.</li>
	<li>Aspirate the 1% BSA in 1x PBS solution. Add 0.5 ml of primary antibody solution and incubate for 1 hr (at RT) or O/N (at 4 &#176;C).</li>
	<li>Remove the primary antibody solution, place into a clean and labeled 1.5 ml microcentrifuge tube, and freeze at -20 &#176;C. 
	NOTE: This allows for the reuse of the primary antibody, if needed.</li>
	<li>Wash the dish 3 - 5 times with 1x PBS, removing and discarding the PBS after each wash.</li>
	<li>Add 0.5 ml of secondary antibody solution. If desired, add fluorescently-labeled phalloidin (following any manufacturer&#8217;s recommended concentration) during this step. Incubate for 1 hr (at RT) or O/N (at 4 &#176;C). 
	NOTE: Use of fluorescently-labeled phalloidin allows for visualization of actin filaments; we have found that 488-labeled phalloidin works best in our zebrafish keratocytes, but other fluorophores can be used.</li>
	<li>Remove and discard the secondary antibody solution.</li>
	<li>Wash the dish 3 - 5 times with 1x PBS, removing and discarding the PBS after each wash. 
	NOTE: The dish can be stored, covered from light, at 4 &#176;C if unable to view immediately; make sure 1x PBS is added into the dish before storing to ensure the cells do not dry out but warm to RT (20 - 30 min), then remove the PBS just prior to proceeding to the next step.</li>
	<li>When ready to observe, add 2 - 3 drops of a mounting medium, with or without DAPI, depending on the experimental needs, into the dish and let sit for 10 min at RT. 
	NOTE: We use a commercially-available, glycerol-based mounting medium with DAPI, but other mounting media can be used. Allowing 10 min for the mounting medium to sit in the dish facilitates easy removal of the glass coverslip from the bottom of the dish.</li>
	<li>After incubation, gently squeeze on the sides of the dish to loosen and remove the coverslip. Add an additional drop of mounting media onto a glass slide and place the coverslip, cells down, onto the slide. 
	NOTE: Consider sealing the coverslip to the slide using clear fingernail polish if the slide needs to be kept for longer than 1 - 2 days.</li>
	<li>Observe the slide under a fluorescence microscope. 
	NOTE: Most of our images were taken using a 40X or 63X objective lens under oil immersion, but the specific objective lens used will depend on the experimental protocol.</li>
</ol>

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The authors have nothing to disclose.

The most critical step for success of keratocyte explant culture is allowing the scale to adhere to the culture dish for approximately 2 - 3 min before addition of the culture medium. Approximately 75% of scales will adhere and grow sheets (the number of cultures established for each experiment will need to be adjusted accordingly). Keratocyte sheets do not form around every scale removed from the fish and placed in culture for several reasons. First, DAPI staining of explants reveals that some scales have few cells atta...

Studies of cell motility have traditionally focused on individually migrating cells. Keratocytes cultured from fish and amphibian explants have proven to be a powerful model system to study the mechanisms of cell motility using both experimental and mathematical modeling approaches1-6. Experimental work on individually migrating cells is aided by the rapid, smooth gliding motion of the cells, while maintaining a uniform shape, speed, and direction. However, in vivo, cells frequently move collectively while maintaining cell-cell junctions, particularly during embryogenesis, wound healing, and metastasis. Thus, collective cell migration is a growing and increasingly prominent area of research within the field of cell migration. The collective migration of these cells has recently been described7 and it has many unique features which make it a powerful system to study collective cell migration in a wound healing model.
When scales are plucked from an anesthetized adult zebrafish, some epithelial tissue remains attached to the underside of the scale. When cell culture medium is added, these epithelial cells, or keratocytes, migrate rapidly as a collective unit from the scale. The explant culture can be viewed as an epithelial wound healing model, in which the cells migrate away from the explant as they would across the provisional matrix of a wound bed to reestablish an intact epithelium. Recent data suggests that this ex vivo culture mimics many features of in vivo wound healing in adult zebrafish. Wound closure rates8 translate to a migration speed similar to that observed in the ex vivo system7. In addition, in an in vivo wound healing model, treatment with warfarin suggests that hemostasis has no effect on rate of healing, while treatment with hydrocortisone to decrease immune response does not delay reepithelialization8. As the zebrafish does not bleed when the scale is removed from the animal, hemostasis is not a major player. Although we have found immune cells in explants, we have not studied the effects they might have on collective cell migration.
The protocol presented here describes how to establish zebrafish keratocyte explant cultures that ensure consistency and reproducibility for use in many biological assays. These explant cultures are a particularly compelling in vitro model for collective cell migration for several reasons. First, zebrafish keratocytes are primary cells used within hours of establishing explant cultures. Therefore, these cells have not undergone the morphological and gene expression changes associated with passage of primary cells9-13. Second, this system represents a model for the study of reepithelialization in response to wounding14 and, as part of the response to wounding, an epithelial to mesenchymal transition (EMT) process is initiated as evidenced by changes in gene expression and cytoskeletal rearrangements14,15. Thus, the background changes in gene expression and motility which occur in untreated cells have been characterized and provide a context to interpret changes with treatment. Furthermore, the fish keratocyte and the human keratinocyte are functionally equivalent; both are the primary epithelial cells of their respective species and play a key role in epithelial wound healing. Third, adult zebrafish are gaining recognition as a model system for a variety of human diseases16-18 including melanoma and other cancers19-21, cutaneous wound healing8,14 and tissue regeneration22-24.
Technical advantages of this model system are equally compelling. A growing number of mutant and transgenic lines are available from non-profit, centralized facilities. Specifically, the Zebrafish Mutation Project (ZMP) at the Wellcome Trust Sanger Institute aims to create a knockout allele in every protein coding gene in the zebrafish genome. Currently, they have mutated 11,892 genes, approximately 45% of the genome, with 24,088 alleles characterized. In addition, ZMP accepts proposals and will generate knock-outs free of charge. Recent methods in gene knockdown in larval and adult zebrafish25-28 provide flexibility to study the function of developmentally important genes for which transgenic approaches are problematic or impractical.
Due to their extremely rapid rates of motility of ~145 &#181;m/hr shortly after establishment of cultures29, assays can be completed rapidly (usually in 24 hr or less). As experiments can be completed rapidly and cultures grow well at RT, several of the technical hurdles associated with mammalian cell migration assays involving video microscopy are avoided. In addition, in the age of tightening research budgets, zebrafish are easily and inexpensively maintained.
The structure and behavior of the explant is complex. As the fish do not bleed when the scale is removed, it is unlikely that much more than the superficial epidermal layers are removed with the scale. Keratocytes appear to be the predominant cell type in the explant when scales are initially removed from the fish as the vast majority of cells appear to stain with an anti-E-cadherin antibody14. Furthermore, there is no evidence of fibroblasts in the initial culture as judged by the absence of vimentin staining by immunofluorescence14. However, with repeated scale removal, there appear to be numerous neutrophils in the explant (see Video 1). We have identified these cells as neutrophils based on morphological observations of their size, rapidly motility, and their migration out of the cell sheet when exposed to LPS (data not shown). Additionally, these cells stain brightly with an antibody to neutrophil cytosolic factor as well as with a variety of secondary IgG antibodies, which indicates that they have abundant FcRs on their surface (data not shown). Multiple cell layers are present within the explant. Confocal microscopy reveals the presence of a single layer near the leading edge to more than two layers of keratocytes near the scale with neutrophils visible above, below, and between the cell keratocyte sheets.
At early time points, cells on the edge of the multi-layer explant polarize, initiating collective migration of keratocytes from the explant at initial rates of ~145 &#181;m/hr. The area covered by the explant rapidly increases, leading to cell spreading, tension within the sheet, and a decreased rate of advance of the leading edge. Rapid interconversions between leader and follower cells are observed at the leading edge during the formation and closure of spontaneously formed holes within the sheet7. As the EMT process initiated with the explant continues, the sheet fragments and the keratocytes lose their specialized, rapid motility. Gene expression and morphological changes consistent with EMT, wound healing, and inflammatory responses occur within 7 days of culture with explants being considered viable for approximately 10 days14.

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			<td height="21" style="height:21px;width:267px;">Name of Material/ Equipment</td>
			<td style="width:71px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:320px;">Comments/Description</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:267px;">35 mm Glass Bottom Culture Dishes (Poly-D Lysine Coated)&#160;</td>
			<td style="width:71px;">MatTek</td>
			<td style="width:119px;">P35GC-1.5-14-C</td>
			<td style="width:320px;">1.5 mm&#160; thickness is best for coverslip removal.&#160; 14 mm diameter provides more culture area.&#160;&#160;</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Swiss Jewelers Style Forceps, Integra, Miltex 17-303</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">21909-460</td>
			<td style="width:320px;">We find that narrow, fine forceps work the best for scale removal.&#160;&#160;</td>
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zebrafish keratocyte explants to study collective cell migration and reepithelialization in cutaneous wound healing

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell migration. However, when explants are established, these cells also move collectively, maintaining many of the features which make individual keratocytes an attractive model to study migration: rapid rates of motility, extensive actin-rich lamellae with a perpendicular actin cable, and relatively constant speed and direction of migration. In early explants, the rapid interconversion of cells migrating individually with those migrating collectively allows the study of the role of cell-cell adhesions in determining the mode of migration, and emphasizes the molecular links between the two modes of migration. Cells in later explants lose their ability to migrate rapidly and collectively as an epithelial to mesenchymal transition occurs and genes associated with wound healing and inflammation are differentially expressed. Thus, keratocyte explants can serve as an in vitro model for the reepithelialization that occurs during cutaneous wound healing and can represent a unique system to study mechanisms of collective cell migration in the context of a defined program of gene expression changes. A variety of mutant and transgenic zebrafish lines are available, which allows explants to be established from fish with different genetic backgrounds. This allows the role of different proteins within these processes to be uniquely addressed. The protocols outlined here describe an easy and effective method for establishing these explant cultures for use in a variety of assays related to collective cell migration.

As illustrated in Figure 1, sheets of keratocytes can be observed migrating out from beneath the scale within hours of establishing the explant culture. Occasionally, an individually migrating keratocyte which has broken away from the collectively migrating sheet can be observed. In addition, as the explant was established from a fish which had been previously plucked, there are abundant neutrophils migrating through the sheet. At longer culture periods, the keratocyte sheet fragments as cells undergo EM...

Watch this Scientific Journal Video about Zebrafish Keratocyte Explants to Study Collective Cell Migration and Reepithelialization in Cutaneous Wound Healing at JoVE.com

Zebrafish Keratocyte Explants to Study Collective Cell Migration and Reepithelialization in Cutaneous Wound Healing

Zebrafish keratocytes migrate in cell sheets from explants and provide an in vitro model for the study of the mechanisms of collective cell migration in the context of epithelial wound healing. These protocols detail an effective way to establish primary explant cultures for use in collective cell migration assays.

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell ...

zebrafish-keratocyte-explants-to-study-collective-cell-migration

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JoVE Journal

Developmental Biology

9.9K 次观看.  Midwestern University. 斑马鱼角膜基质细胞的表外植迁移，并提供了一个体外模型集体细胞迁移的上皮伤口愈合的情况下机制的研究。这些协议的细节，以建立初级外植体培养在集体细胞迁移测定法使用的有效方式。 

视频: Zebrafish Keratocyte Explants to Study Collective Cell Migration and Reepithelialization in Cutaneous Wound Healing

Optimization of the Wound Scratch Assay to Detect Changes in Murine Mesenchymal Stromal Cell Migration After Damage by Soluble Cigarette Smoke Extract

Cell migration is vital to many physiological and pathological processes including tissue development, repair, and regeneration, cancer metastasis, and inflammatory responses. Given the current interest in the role of mesenchymal stromal cells in mediating tissue repair, we are interested in quantifying the migratory capacity of these cells, and understanding how migratory capacity may be altered after damage. Optimization of a rigorously quantitative migration assay that is both easy to customize and cost-effective to perform is key to answering questions concerning alterations in cell migration in response to various stimuli. Current methods for quantifying cell migration, including scratch assays, trans-well migration assays (Boyden chambers), micropillar arrays, and cell exclusion zone assays, possess a range of limitations in reproducibility, customizability, quantification, and cost-effectiveness. Despite its prominent use, the scratch assay is confounded by issues with reproducibility related to damage of the cell microenvironment, impediments to cell migration, influence of neighboring senescent cells, and cell proliferation, as well as lack of rigorous quantification. The optimized scratch assay described here demonstrates robust outcomes, quantifiable and image-based analysis capabilities, cost-effectiveness, and adaptability to other applications.

The capacity to migrate is a key function of many different cell types, including mesenchymal stromal cells (MSCs). However, quantifying alterations in migratory capacity after damage is challenging. This protocol describes an easily adaptable migration assay that uses rigorous statistics to quantify changes in MSC migratory capacity after damage.

Cell migration is vital to many physiological and pathological processes including tissue development, repair, and regeneration, cancer metastasis, and ...

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, ...

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Development of an Ethanol-induced Fibrotic Liver Model in Zebrafish to Study Progenitor Cell-mediated Hepatocyte Regeneration

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading to cirrhosis with hepatocellular dysfunction. Among multiple hepatic insults, alcohol abuse can lead to significant health problems including liver failure and hepatocellular carcinoma. Nonetheless, the identity of endogenous cellular sources that regenerate hepatocytes in response to alcohol has not been properly investigated. Moreover, few studies have effectively modeled hepatocyte regeneration upon alcohol-induced injury. We recently reported on establishing an ethanol (EtOH)-induced fibrotic liver model in zebrafish in which hepatic progenitor cells (HPCs) gave rise to hepatocytes upon near-complete hepatocyte loss in the presence of fibrogenic stimulus. Furthermore, through chemical screens using this model, we identified multiple small molecules that enhance hepatocyte regeneration. Here we describe in detail the procedures to develop an EtOH-induced fibrotic liver model and to perform chemical screens using this model in zebrafish. This protocol will be a critical tool to delineate the molecular and cellular mechanisms of how hepatocyte regenerates in the fibrotic liver. Furthermore, these methods will facilitate potential discovery of novel therapeutic strategies for chronic liver disease in vivo.

Sustained fibrosis with deposition of excessive extracellular matrix proteins leads to cirrhosis. Alcohol abuse is one of the main causes of severe liver disease. We established an ethanol-induced zebrafish fibrotic liver model to study the mechanisms and strategies of promoting hepatocyte regeneration upon alcohol-induced injury.

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading ...

Adenoviral Gene Therapy for Diabetic Keratopathy: Effects on Wound Healing and Stem Cell Marker Expression in Human Organ-cultured Corneas and Limbal Epithelial Cells

The goal of this protocol is to describe molecular alterations in human diabetic corneas and demonstrate how they can be alleviated by adenoviral gene therapy in organ-cultured corneas. The diabetic corneal disease is a complication of diabetes with frequent abnormalities of corneal nerves and epithelial wound healing. We have also documented significantly altered expression of several putative epithelial stem cell markers in human diabetic corneas. To alleviate these changes, adenoviral gene therapy was successfully implemented using the upregulation of c-met proto-oncogene expression and/or the downregulation of proteinases matrix metalloproteinase-10 (MMP-10) and cathepsin F. This therapy accelerated wound healing in diabetic corneas even when only the limbal stem cell compartment was transduced. The best results were obtained with combined treatment. For possible patient transplantation of normalized stem cells, an example is also presented of the optimization of gene transduction in stem cell-enriched cultures using polycationic enhancers. This approach may be useful not only for the selected genes but also for the other mediators of corneal epithelial wound healing and stem cell function.

An example of adenoviral gene therapy in the human diabetic organ-cultured corneas is presented towards the normalization of delayed wound healing and markedly reduced epithelial stem cell marker expression in these corneas. It also describes the optimization of this process in stem cell-enriched limbal epithelial cultures.

The goal of this protocol is to describe molecular alterations in human diabetic corneas and demonstrate how they can be alleviated by adenoviral gene ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

A Device for Performing Cell Migration/Wound Healing in a 96-Well Plate

The cell migration/wounding assay is a commonly used method to study cell migration and other biological processes, such as angiogenesis and tumor metastasis. In this assay, cells are grown to form a confluent monolayer and a mechanical wound is created by scratching with a device. Then the migration rate of the cells towards the denuded area can be monitored by imaging. Our 8-channel mechanical wounder is designed to tackle most of the problems associated with the cell migration assay. Firstly, our wounder can be easily sterilized by autoclaving or with common disinfectants. Secondly, the individual adjustable pins allow even contact with the cell culture plate so that sharp and reproducible wounds can be created. Thirdly, the guiding bars on both sides of the wounder ensure consistent wounding position in each well. The use of disposable plastic pipette tips for wounding can further provide better handling of the wounder as well as to minimize cross-contamination. In conclusion, our cell wounder can provide researchers with a user friendly and reproducible device for performing the cell migration assay using the standard 96-well culture plate.

Here, we present a protocol to perform a semi-high throughput cell migration assay on a 96-well cell culture plate. This protocol is a fast, simple and economical method to create consistent scratch wounds on a cell monolayer.

The cell migration/wounding assay is a commonly used method to study cell migration and other biological processes, such as angiogenesis and tumor ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...

A Drosophila Model to Study Wound-induced Polyploidization

Polyploidy is a frequent phenomenon whose impact on organismal health and disease is still poorly understood. A cell is defined as polyploid if it contains more than the diploid copy of its chromosomes, which is a result of endoreplication or cell fusion. In tissue repair, wound-induced polyploidization (WIP) has been found to be a conserved healing strategy from fruit flies to vertebrates. WIP has several advantages over cell proliferation, including resistance to oncogenic growth and genotoxic stress. The challenge has been to identify why polyploid cells arise and how these unique cells function. Provided is a detailed protocol to study WIP in the adult fruit fly epithelium where polyploid cells are generated within 2 days after a puncture wound. Taking advantage of D. melanogaster&#39;s extensive genetic tool kit, the genes required to initiate and regulate WIP, including Myc, have begun to be identified. Continued studies using this method can reveal how other genetic and physiological variables including sex, diet, and age regulate and influence WIP&#39;s function.

A Drosophila Model to Study Wound-induced Polyploidization

Wound-induced polyploidization is a conserved tissue repair strategy where cells grow in size instead of dividing to compensate for cell loss. Here is a detailed protocol on how to use the fruit fly as a model to measure ploidy and its genetic regulation in epithelial wound repair.

Polyploidy is a frequent phenomenon whose impact on organismal health and disease is still poorly understood. A cell is defined as polyploid if it ...

Murine Excisional Wound Healing Model and Histological Morphometric Wound Analysis

The murine excisional wound model has been used extensively to study each of the sequentially overlapping phases of wound healing: inflammation, proliferation and remodeling. Murine wounds have a histologically well-defined and easily recognizable wound bed over which these different phases of the healing process are measurable. Within the field, it is common to use an arbitrarily defined &#8220;middle&#8221; of the wound for histological analyses. However, wounds are a three-dimensional entity and often not histologically symmetrical, supporting the need for a well-defined and robust method of quantification to detect morphometric defects with a small effect size. In this protocol, we describe the procedure for creating bilateral, full-thickness excisional wounds in mice as well as a detailed instruction on how to measure morphometric parameters using an image processing program on select serial sections. The two-dimension measurements of wound length, epidermal length, epidermal area, and wound area are used in combination with the known distance between sections to extrapolate the three-dimension epidermal area covering the wound, overall wound area, epidermal volume and wound volume. Although this detailed histological analysis is more time and resource consuming than conventional analyses, its rigor increases the likelihood of detecting novel phenotypes in an inherently complex wound healing process.

This protocol describes how to generate bilateral, full-thickness excisional wounds in mice and how to subsequently monitor, harvest, and prepare the wounds for morphometric analysis. Included is an in-depth description of how to use serial histological sections to define, precisely quantify and detect morphometric defects.