Authors
Contact Us

Sign In

A subscription to JoVE is required to view this content. Sign in or start your free trial.

In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

These protocols were developed to analyze cortical lens morphology, structural integrity of the zebrafish lens sutures in fixed and live lenses and to measure the position of the zebrafish lens nucleus along the anterior-posterior axis.

Abstract

The zebrafish is uniquely suited to genetic manipulation and in vivo imaging, making it an increasingly popular model for reverse genetic studies and for generation of transgenics for in vivo imaging. These unique capabilities make the zebrafish an ideal platform to study ocular lens development and physiology. Our recent findings that an Aquaporin-0, Aqp0a, is required for stability of the anterior lens suture, as well as for the shift of the lens nucleus to the lens center with age led us to develop tools especially suited to analyzing the properties of zebrafish lenses. Here we outline detailed methods for lens dissection that can be applied to both larval and adult lenses, to prepare them for histological analysis, immunohistochemistry and imaging. We focus on analysis of lens suture integrity and cortical cell morphology and compare data generated from dissected lenses with data obtained from in vivo imaging of lens morphology made possible by a novel transgenic zebrafish line with a genetically encoded fluorescent marker. Analysis of dissected lenses perpendicular to their optical axis allows quantification of the relative position of the lens nucleus along the anterior-posterior axis. Movement of the lens nucleus from an initial anterior position to the center is required for normal lens optics in adult zebrafish. Thus, a quantitative measure of lens nuclear position directly correlates with its optical properties.

Introduction

The zebrafish is an excellent model for studying lens development and physiology due to the anatomical similarities to mammalian lenses, relative ease of genetic and pharmacological manipulation, speed of embryonic eye development, small size and transparency at early stages allowing for in vivo imaging. The methods described here were developed to analyze zebrafish lens morphology at embryonic and adult stages with a focus on sutural integrity, cortical membrane morphology in vitro and in vivo, and location of lens nuclear position along the anterior-posterior axis ex vivo. These methods serve as a starting point for functional studies of lens development and physiology, as well as reverse genetic screens for lens phenotypes in zebrafish.

Imaging zebrafish lens morphology

Aquaporin 0 (AQP0) is the most abundant lens membrane protein and is critical for both, lens development and clarity, due to multiple essential functions in mammals. Zebrafish have two Aqp0s (Aqp0a and Aqp0b) and we have developed methods to analyze loss of their functions in both embryonic and adult lenses. Our studies reveal that aqp0a-/- mutants develop anterior polar opacity due to instability of the anterior suture, and aqp0a/b double mutants develop nuclear opacity1. AQP0 has been shown to play roles in water transport2, adhesion3,4, cytoskeletal anchoring5 and generation of the refractive index gradient6, but these studies have largely been performed in vitro. The zebrafish provides a unique opportunity to study how loss of function, or perturbed function of Aqp0a or Aqp0b would affect morphology and function in a living lens. To assess lens cell morphology and sutural integrity during development, we modified existing in vitro immunohistochemical methods7 for use in embryonic and adult lenses, and generated transgenics to monitor this process in vivo.

Immunohistochemical analysis of plasma membrane structure and sutural integrity was performed in whole fixed embryos and adult lenses. Zebrafish lenses are extremely small (lens diameter is ~100 µm in embryos and up to 1 mm in adults) compared with their mammalian counterparts and have point sutures8, which are infrequently captured in cryosections. Thus, whole lenses are essential for analyzing sutural integrity. For in vivo analysis of anterior suture formation, and imaging of precise lens cell architecture, we generated transgenics expressing mApple specifically labeling lens membranes.

Advantages of live imaging of lens membrane transgenics include: 1) avoiding fixation artifacts, 2) studying dynamic morphological changes in time-lapse movies, and 3) enabling longitudinal studies in which earlier events can be correlated with later phenotypes. Pigmentation of the iris normally prevents clear imaging of the lens periphery. Addition of 1-phenyl 2-thiourea (PTU) before the primordia-5 (prim-5) stage9 prevents melanogenesis and eye pigmentation up to around 4 days postfertilization (dpf). However, after 4 dpf, the lens periphery is obscured in vivo, particularly at older stages. Furthermore, the density of the lens itself obscures imaging of its posterior pole. Therefore, to study morphology of the lens periphery, or the posterior suture, after 4 dpf, lenses need to be excised and fixed.

Transgenic zebrafish lines have been used to analyze embryonic lens membrane structure in vivo10. The Q01 transgenic expresses a cyan fluorescent protein fused to a membrane targeting sequence, Gap43, driven by the EF1α promoter and a hexamer of the DF4 pax6 enhancer element ubiquitously in lens fiber cells11. Q01 does have extra-lenticular expression, including amacrine cells in the retina, which adds background signal if the primary interest is the lens. We developed a novel transgenic line that expresses a membrane-tethered mApple specifically in the lens, with the aim of avoiding any extra-lenticular signal.

Lens nucleus localization

We discovered that the lens nucleus moves from an initial anterior location in larval zebrafish to a central location in the anterior-posterior axis in adult lenses. This shift in the position of the high refractive index lens nucleus is thought to be a requirement for normal development of zebrafish lens optics1. Our methods allow quantification of lens nuclear centralization in relation to the lens radius. Using this method, we showed that Aqp0a is required for lens nuclear centralization1, and this simple method can be applied to other studies of the development and physiology of the lens and its optical properties in the zebrafish model.

Protocol

The animal protocols used in this study adhere to the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and have been approved by the Institutional Animal Care and Use Committee (IACUC) of University of California, Irvine.

1. Zebrafish Husbandry and Euthanasia

  1. Raise and maintain zebrafish (AB strain) under standard laboratory conditions12. Raise embryos in embryo medium (EM)12. Add 0.003% PTU to EM from 20-24 h postfertilization (before the prim-5 stage) to prevent pigment formation in embryos12 used for imaging at embryonic stages9. Raise larvae from 6-30 days postfertilization (dpf) on a diet of live rotifers until 14 dpf, and live artemia after 14 dpf12.
    CAUTION: PTU is very toxic
  2. Anesthetize fish using tricaine until non-responsive to touch.
    1. Prepare tricaine stock by combining 400 mg of 3-amino benzoic acidethylester, with 2.1 mL of 1 M Tris (pH 9), in 100 mL of ddH2O. Adjust to pH 7.0 and store at -20 °C.
      CAUTION: Tricaine is toxic.
    2. Dilute 4.2 mL of tricaine stock in 100 mL of tank water to anesthetize larvae/adults or in EM to anesthetize embryos (final concentration of tricaine at 0.0165% w/v).

2. Fixation of Embryos and Larvae

NOTE: Immunohistochemical protocols were adapted from previously published materials7.

  1. Fix dechorionated embryos or larvae up to 2 weeks postfertilization in 4% (v/v) paraformaldehyde (PFA) in phosphate buffered saline (PBS) overnight at 4 °C on a rocker.
    CAUTION: PFA is combustible and is carcinogenic.
  2. Wash embryos three times for 10 min in PBS, and permeabilize in PBS with 10% Triton and 1% DMSO (PBS-T) overnight at 4 °C.
    CAUTION: DMSO is toxic, is harmful by ingestion or skin absorption, carries hazardous materials through skin, and is combustible. Triton is toxic, corrosive and is hazardous to aquatic environments.

3. Dissection of Larval and Adult Zebrafish Lenses

  1. Anesthetize fish from 6 dpf larvae to adulthood with tricaine until non-responsive to touch but still showing a strong heartbeat. Measure fish standard length as per Schilling (2002) for staging13.
  2. Immediately excise eyes using micro-dissection scissors and place into a dissection dish in PBS (Figure 2 shown for adult eyes).
    1. Make a custom 35 mm dish filled with silicone for lens dissections. Once silicone has set, excise a divot around 2-3 mm in diameter and 0.5 mm deep for immobilizing adult eyes/lenses during dissection.
  3. Dissection of adult zebrafish lenses
    1. Place an adult eye into the dish divot posterior side up filled with PBS. Immobilize the eye by inserting forceps at <45° angle through the optic disc. Be careful not to nick or compress the lens. Make two or three radial incisions through retina and sclera from the optic disc to the ciliary zone with dissection scissors.
    2. Peel back the retina and sclera like flower petals and invert the eye, cornea side up. Immobilize the lens indirectly via manipulation of the sclera and cornea with the flat side of the scissors, while pulling away the retina and attached tissues with forceps. Carefully trim excess tissue from the lens.
      NOTE: It is vital to dissect carefully to obtain consistently healthy lenses.
  4. Dissection of larval zebrafish lenses
    1. Place a larval eye posterior side up onto the flat part of the silicone dish filled with PBS and use a sharpened tungsten needle to make radial cuts through the retina and sclera while immobilizing the eye with another tungsten needle or forceps.
      NOTE: Be careful not to damage the lens.
      1. Sharpen the tip of a 2 cm length of 0.1 mm tungsten wire electrolytically by suspending the wire tip into 10% (w/v) NaOH and applying a low voltage alternating-current14. Secure the needle into a Pasteur pipette by melting the glass end using a Bunsen burner.
        NOTE: Alternative fine dissection tools can also be used.
        CAUTION: NaOH is corrosive.
    2. Gently scoop out the lens from the dissociated eye with a blunt side of the needle, and carefully pull away attached tissue.

4. Fixation of Dissected Lenses

  1. Immediately fix dissected lenses in 1.5% (v/v) PFA in PBS for 24 h at room temperature (RT). Wash lenses three times in PBS for 10 min each.
  2. Permeabilize lenses for whole mount analysis or cryoprotect for cryosectioning (see section 8).
    1. Permeabilize lenses in PBS-T overnight at 4 °C.

5. Lens Immunohistochemistry

  1. Label membranes of fixed embryo/larval or adult lenses by incubation in Phalloidin- Alexa Fluor 546 (1:200) and cell nuclei with DAPI (1:1,000 of 5 mg/mL stock) in PBS-T overnight at 4 °C.
    CAUTION: DAPI is a skin and eye irritant.
  2. Wash three times with PBS-T for 10 min each. Clear tissue by incubation in 30%, 50% and 70% glycerol in PBS-T (v/v) for at least 1 h at RT each, or overnight at 4 °C.
  3. Mount fish in 70% glycerol in PBS-T (v/v) onto glass bottom 35 mm microwell dishes with eyes as flat and straight against the cover slip as possible. For younger fish, a slight anterior-lateral tilt may help to orient the lens suture to be parallel to the coverslip.

6. Analysis of Zebrafish Anterior Lens Sutures Using a Transgenic Line In Vivo

  1. Generate Tg(βB1cry:mAppleCAAX) lines using the Tol2 kit15. The Tol2 transposable element system15 enables stable integration of the construct where mApple is driven by 300 bp of the human βB1-crystallin promoter16 tethered to the membrane by the CAAX sequence resulting in expression specifically in lens fiber cell membranes.
    NOTE: This construct (ID:122451) is available for purchase.
    1. On the morning of injection, prepare the injection mixture and keep on ice. To reach a final volume of 10 µL of containing RNase- and DNase-free H2O, add: 1.5 µL of huβB1cry:mAppleCAAX DNA construct at 50 ng/µL - [0.375-0.75 pg/final injection], 1.5 µL of Tol2 transposase mRNA at 300 ng/µL (Tol2 kit)15 [15-30 pg/final injection], and 1 µL of phenol red indicator at 1% w/v [1 x10-5% (w/v)/final injection].
    2. Inject 50-100 pL of injection mixture into 1-cell stage embryos12.
  2. Measure transgenesis efficiency in F0 injected embryos by measuring frequency of embryos with mApple positive lenses at 3 dpf.
    NOTE: Expect to have >80% transgenesis efficiency using this method. F0 mosaics allow detailed analysis of membrane morphologies of individual cells. Varying levels of mosaicism allows one to image the morphologies of single or small groups of cells, while more global lens expression allows analysis of multicellular structures like lens sutures in vivo.
  3. Outcross F0 mosaic fish to generate stable lines, which label all fiber cell membranes. F0 founders with the transgene integrated into the germline will generate offspring with stable integrations that can be maintained as transgenic lines.

7. Analysis of Zebrafish Anterior Lens Sutures Using a Transgenic Line In Vivo

  1. Anesthetize 3 dpf or adult mosaic or stable transgenic fish and mount in 1% low melt agarose (LMA) with tricaine with the eye flat against the cover slip of glass bottom microwell dishes.
    1. Dissolve 1 g of LMA in 100 mL of EM (without methylene blue) to make 1% LMA. Store long term as 15 mL stocks at 4 °C. Microwave to dissolve stock and store at 42 °C until each tube is used.
  2. Cover fish up to 6 dpf with EM with tricaine, or with tank water with tricaine for adults when LMA is set.
    1. Mix 5 mL of EM without methylene blue or tank water with 250 µL of tricaine stock and 7 µL of PTU if imaging PTU treated fish.

8. Lens Cryosectioning and Immunohistochemistry

  1. Cryoprotect fixed embryos or adult lenses by placing into 10% sucrose in PBS (w/v), 20% sucrose for 1 h at RT each (or overnight at 4 °C) and overnight in 30% sucrose at 4 °C.
  2. Embed tissue in OCT in base molds, and then freeze onto chucks with OCT. Cryosection at 12-14 µm and collect onto a Superfrost/Plus microscope slides. Warm sections onto slide for 10 min on a slide warmer prewarmed to ~35 °C.
  3. Wash sections three times with PBS for 10 min each. Label sections with Phalloidin-Alexa Flour 546 (1:200) with DAPI (1:1,000) or WGA-Alexa Flour 594 (1:200), followed by three PBS washes. Mount with anti-fade mounting medium, and seal coverslip with nail polish.

9. Imaging

  1. Acquire images with a confocal microscope. Acquire z-stacks or optical slices using a 60x N/A 1.2 water-immersion objective, or similar, at specific regions from the anterior to posterior lens pole. Use the following acquisition settings: 1.2 airy disc using the 561 nm laser, Texas red filter for phalloidin-Alexa Flour 546 or mApple, or the 405 laser with the UV filter for DAPI.
  2. Correct z-intensity laser power to counteract signal loss with depth into the lens. This allows for a strong signal at the posterior pole of fixed and live 3 dpf embryos, and strong signal in equatorial optical slices in adult fish lenses.
  3. Compile and view images using image processing software.

10. Measurement of Lens Nucleus Localization in the Anterior-posterior Axis

  1. Orient freshly excised lenses axially in PBS in a 35 mm dish with a coverglass bottom, with poles and sutures oriented parallel to the plane of focus. Look for a difference in the refractive index, which usually occurs at the interface of the lens cortex and lens nucleus to identify the lens nucleus.
    NOTE: Healthy wild type adult lenses are extremely transparent making it difficult to see the lens sutures and nucleus, or to determine lens orientation. In this case, a slight nick with forceps to the lens capsule causes minor damage, which makes the sutures and lens nucleus apparent in about 10 min helping to orient the lens for this measurement. However, the lenses become unusable for downstream applications as they are now damaged.
  2. Take images of lenses with the lens nucleus in focus under bright field illumination with or without DIC optics using a dissection microscope with a camera attached. Image a micrometer under the same magnification for calibration.
    1. Click on the live view button, adjust the exposure settings to visualize the lens nucleus and lens periphery, and take an image by clicking the snap shot button. Save as a tif file.
    2. Use image processing software to calibrate the acquired lens images. Select the straight-line tool and draw a line of known length on the micrometer image, click analyze | set scale. Enter known distance, units and select 'global' calibration, and click ok.
  3. Measure the distance of the center of the lens nucleus to the anterior pole (a - r).
    1. Use the straight-line tool in image processing software to draw a line across the imaged center of the lens nucleus in an axial orientation. Take the center of this line as the center of the lens nucleus.
    2. Draw another line from this point to the anterior pole of the lens and select 'measure' in the 'analyze' menu to measure the distance (a - r), where a is the radius of the lens and r is the distance from the center of the lens to the center of the nucleus.
    3. Draw another line from the anterior to the posterior poles and measure this distance as the lens diameter (2a). Copy these lengths from the 'results' window and transfer to a spreadsheet or statistics program.
      NOTE: It is easier to precisely measure from the center of the nucleus to the anterior pole, than measuring from the center of the lens nucleus to the center of the lens. This measurement is converted by the formula in step 10.3.5 to report the distance of the center of the lens nucleus to the center of the lens. Always use the adult nucleus for measurements, even if the embryonic nucleus is evident.
    4. Divide the diameter by 2, to calculate the lens radius, a.
    5. Calculate r/a, as figure-protocol-13570, which is the normalized localization of the lens nucleus with respect to the lens radius.
      NOTE: For a nucleus placed closer to the anterior pole, r/a will be >0.0, while a centrally localized nucleus will have an r/a of 0.0.
  4. Graph the log of the normalized axial nucleus measurement as a function of the standard length to determine changes in lens nucleus localization during zebrafish development.

Results

Adult zebrafish eye anatomy closely resembles that of mammals (Figure 1A). Despite some differences between zebrafish and mammalian eyes, such as having a ciliary zone instead of a ciliary body17, differences in optical properties18, and differences in morphogenesis during embryonic development19, the zebrafish eye is an excellent model for studying eye development and understanding ophth...

Discussion

Analysis of zebrafish lens morphology is the initial step in understanding phenotypes in mutants, or the effects of pharmacological interventions aimed at studying biology of the ocular lens. We outline methods to analyze lens sutures, cortical fiber cell morphology and aspects of the lens nucleus. These approaches are a combination of in vitro and in vivo (compared in Table 1). The in vitro methods allow for greater detail of the outer cortical cell morphology, as well as acce...

Disclosures

We have no disclosures.

Acknowledgements

We would like to acknowledge our funding source: NIH R01 EY05661 to J.E.H, Ines Gehring for assisting with generating the aqp0a/b double mutants and zebrafish husbandry, Dr Daniel Dranow for discussions leading to generation of the transgenics, Dr Bruce Blumberg and Dr Ken Cho’s labs for use of their dissecting microscopes, and Dr Megan Smith for help with statistical analysis.

Materials

NameCompanyCatalog NumberComments
1-phenyl-2-thiourea (PTU)SigmaP7629CAUTION – very toxic
4% Paraformaldehyde aqueous solutionElectron Microscopy SciencesRT 157-4CAUTION – health hazard, combustible
Confocal microscopeNikonEclipse Ti-E
CryostatLeicaCM3050SObjective and chamber temperature set to -21˚C
DAPIInvitrogenD1306CAUTION – irritating to eyes and skin
Dimethyl Sulfoxide (DMSO)Fisher ScientificD128CAUTION – combustible, penetrates skin 
Disposable base moldVWR Scientific15154-631
Disposable Pasteur glass pipetsFisherbrand13-678-20A
Dumont # 5 forcepsDumont & FilsKeep forceps sharpened
Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine)Sigma-AldrichA5040CAUTION - toxic
Glass bottom microwell dish (35mm petri dish, 14mm microwell, #1.5 coverglass)MatTek CorporationP35G-1.5-14-C
GlycerolSigmaG2025
huβB1cry:mAppleCAAX DNA constructAddgeneID:122451
ImageJWayne Rasband, NIHv1.51n
Low Melt Agarose (LMA)Apex902-36-6
NIS-ElementsNikonV 4.5
NIS-Elements AR softwareNikon
Olympus  with a model 2.1.1.183 controllerOlympus CorpDP70
Olympus microscope Olympus CorpSZX12
Phalloidin-Alexa Flour 546Thermo FisherA22283
Phenol Red indicator (1% w/v)Ricca Chemical Company5725-16
Phosphate buffered saline (PBS)Fisher ScientificBP399
PhotoshopAdobeCS5 v12.0
Photoshop softwareAdobeCS5 v12.0
Plan Apo 60x/1.2 WD objectiveNikon
Power sourceWild HeerbruggMTr 22Or equivalent power source 
Slide warmer model No. 26020FSFisher Scientific12-594
Sodium Hydroxide beadsFisher ScientificS612-3CAUTION - corrosive/irritating to eyes and skin, target organ - respiratory system, corrosive to metals
Superfrost/Plus microscope slideFisher Scientific12-550-15
Sylgard 184 silicone Dow CorningWorld Prevision InstrumentsSYLG184
Tissue-Tek O.C.T. CompoundSakura Finetek4583
Triton X-100 BioXtraSigmaT9284CAUTION – Toxic, hazardous to aquatic environment, corrosive
Vannas micro-dissection scissorsTed Pella Inc1346Sharp/sharp straight tips
Vectashield antifade mouting mediumVector laboratoriesH-1000
Wheat Germ Agglutinin (WGA)-Alexa Flour-594Life TechnologiesW11262

References

  1. Vorontsova, I., Gehring, I., Hall, J. E., Schilling, T. F. Aqp0a Regulates Suture Stability in the Zebrafish Lens. Investigative Ophthalmology & Visual Science. 59 (7), 2869-2879 (2018).
  2. Hall, J. E., Mathias, R. T. The Aquaporin Zero Puzzle. Biophysical Journal. 107 (1), 10-15 (2014).
  3. Nakazawa, Y., Oka, M., Funakoshi-Tago, M., Tamura, H., Takehana, M. The Extracellular C-loop Domain Plays an Important Role in the Cell Adhesion Function of Aquaporin 0. Current Eye Research. 42 (4), 617-624 (2017).
  4. Kumari, S. S., Varadaraj, K. Intact AQP0 performs cell-to-cell adhesion. Biochemical and Biophysical Research Communications. 390 (3), 1034-1039 (2009).
  5. Lindsey Rose, K. M., et al. The C Terminus of Lens Aquaporin 0 Interacts with the Cytoskeletal Proteins Filensin and CP49. Investigative Ophthalmology & Visual Science. 47 (4), 1562-1570 (2006).
  6. Kumari, S. S., Varadaraj, K. Aquaporin 0 plays a pivotal role in refractive index gradient development in mammalian eye lens to prevent spherical aberration. Biochemical and Biophysical Research Communications. 452 (4), 986-991 (2014).
  7. Uribe, R. A., Gross, J. M. Immunohistochemistry on Cryosections from Embryonic and Adult Zebrafish Eyes. Cold Spring Harbor Protocols. 2007 (7), (2007).
  8. Dahm, R., Schonthaler, H. B., Soehn, A. S., van Marle, J., Vrensen, G. F. J. M. Development and adult morphology of the eye lens in the zebrafish. Experimental Eye Research. 85 (1), 74-89 (2007).
  9. Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. Stages of embryonic development of the zebrafish. Developmental Dynamics. 203 (3), 253-310 (1995).
  10. Greiling, T. M. S., Clark, J. I. Early lens development in the zebrafish: A three-dimensional time-lapse analysis. Developmental Dynamics. 238 (9), 2254-2265 (2009).
  11. Godinho, L., et al. Targeting of amacrine cell neurites to appropriate synaptic laminae in the developing zebrafish retina. Development. 132 (22), 5069-5079 (2005).
  12. Westerfield, M. . The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Danio rerio). , (2002).
  13. Schilling, T. F., Nusslein-Volhard, C., Dahm, R. . Zebrafish. 261, 59-94 (2002).
  14. Brady, J. A simple technique for making very fine, durable dissecting needles by sharpening tungsten wire electrolytically. Bulletin of the World Health Organization. 32 (1), 143-144 (1965).
  15. Kwan, K. M., et al. The Tol2kit: A multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Developmental Dynamics. 236 (11), 3088-3099 (2007).
  16. Hou, H. H., Kuo, M. Y. P., Luo, Y. W., Chang, B. E. Recapitulation of human βB1-crystallin promoter activity in transgenic zebrafish. Developmental Dynamics. 235 (2), 435-443 (2006).
  17. Soules, K., Link, B. Morphogenesis of the anterior segment in the zebrafish eye. BMC Developmental Biology. 5 (1), 12 (2005).
  18. Greiling, T. M. S., Clark, J. I. The transparent lens and cornea in the mouse and zebra fish eye. Seminars in Cell & Developmental Biology. 19 (2), 94-99 (2008).
  19. Greiling, T. M. S., Aose, M., Clark, J. I. Cell Fate and Differentiation of the Developing Ocular Lens. Investigative Ophthalmology & Visual Science. 51 (3), 1540-1546 (2010).
  20. Richardson, R., Tracey-White, D., Webster, A., Moosajee, M. The zebrafish eye-a paradigm for investigating human ocular genetics. Eye. 31, 68 (2016).
  21. Gestri, G., Link, B. A., Neuhauss, S. C. F. The Visual System of Zebrafish and its Use to Model Human Ocular Diseases. Developmental Neurobiology. 72 (3), 302-327 (2012).
  22. Chhetri, J., Jacobson, G., Gueven, N. Zebrafish on the move towards ophthalmological research. Eye. 28 (4), 367-380 (2014).
  23. Lim, J. C., Walker, K. L., Sherwin, T., Schey, K. L., Donaldson, P. J. Confocal Microscopy Reveals Zones of Membrane Remodeling in the Outer Cortex of the Human Lens. Investigative Ophthalmology & Visual Science. 50 (9), 4304-4310 (2009).
  24. Lim, J. C., Vaghefi, E., Li, B., Nye-Wood, M. G., Donaldson, P. J. Characterization of the Effects of Hyperbaric Oxygen on the Biochemical and Optical Properties of the Bovine LensEffects of HBO on the Bovine Lens. Investigative Ophthalmology & Visual Science. 57 (4), 1961-1973 (2016).
  25. Gold, M. G., et al. AKAP2 anchors PKA with aquaporin-0 to support ocular lens transparency. EMBO Molecular Medicine. 4 (1), 15-26 (2012).
  26. Petrova, R. S., Schey, K. L., Donaldson, P. J., Grey, A. C. Spatial distributions of AQP5 and AQP0 in embryonic and postnatal mouse lens development. Experimental Eye Research. 132, 124-135 (2015).
  27. Chee, K. N., Vorontsova, I., Lim, J. C., Kistler, J., Donaldson, P. J. Expression of the sodium potassium chloride cotransporter (NKCC1) and sodium chloride cotransporter (NCC) and their effects on rat lens transparency. Molecular Vision. 16, 800-812 (2010).
  28. Hayes, J. M., et al. Integrin α5/fibronectin1 and focal adhesion kinase are required for lens fiber morphogenesis in zebrafish. Molecular Biology of the Cell. 23 (24), 4725-4738 (2012).
  29. Imai, F., Yoshizawa, A., Fujimori-Tonou, N., Kawakami, K., Masai, I. The ubiquitin proteasome system is required for cell proliferation of the lens epithelium and for differentiation of lens fiber cells in zebrafish. Development. 137 (19), 3257-3268 (2010).
  30. Biswas, S. K., Lee, J. E., Brako, L., Jiang, J. X., Lo, W. K. Gap junctions are selectively associated with interlocking ball-and-sockets but not protrusions in the lens. Molecular Vision. 16, 2328-2341 (2010).

Reprints and Permissions

Request permission to reuse the text or figures of this JoVE article

Request Permission

Explore More Articles

ZebrafishLens DevelopmentLens MorphologyLens OpticsSutural IntegrityMicrodissectionCataract PreventionGenetic ScreeningDissection TechniquesOptical DiscSagittal OrientationRefractive IndexLarval ZebrafishPBS SolutionTissue Harvesting

This article has been published

Video Coming Soon

JoVE Logo

Privacy

Terms of Use

Policies

Contact Us

Recommend to library

JoVE NEWSLETTERS

Research

Education

Authors

Librarians

ABOUT JoVE

Copyright © 2025 MyJoVE Corporation. All rights reserved