The Zebrafish: Cellular and Developmental Biology, Part B, Volume 101, Third Edition (Methods in Cell Biology)

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Author: H. William Detrich III | Monte Westerfield | Leonard Zon

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Methods in Cell Biology VOLUME 101 The Zebrafish: Cellular and Developmental Biology, Part B Third Edition

Series Editors Leslie Wilson Department of Molecular, Cellular and Developmental Biology University of California Santa Barbara, California

Paul Matsudaira Department of Biological Sciences National University of Singapore Singapore

Methods in Cell Biology VOLUME 101 The Zebrafish: Cellular and Developmental Biology, Part B Third Edition

Edited by

H. William Detrich III Department of Biology, Northeastern University, Boston, MA, USA

Monte Westerfield Institute of Neuroscience, University of Oregon, Eugene, OR, USA

Leonard I. Zon Division of Hematology/Oncology, Children’s Hospital of Boson, Department of Pediatrics and Howard Hughes Medical Institute, Harvard Medical School, Boston, MA, USA

AMSTERDAM BOSTON HEIDELBERG LONDON NEW YORK OXFORD PARIS SAN DIEGO SAN FRANCISCO SINGAPORE SYDNEY TOKYO Academic Press is an imprint of Elsevier

Academic Press is an imprint of Elsevier 225 Wyman Street, Waltham, MA 02451, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 32, Jamestown Road, London NW1 7BY, UK Linacre House, Jordan Hill, Oxford OX2 8DP, UK First edition 2011 Copyright # 2011 Elsevier Inc. All rights reserved No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made ISBN: 978-0-12-387036-0 ISSN: 0091-679X For information on all Academic Press publications visit our website at elsevierdirect.com

Printed and bound in USA 11 12 13

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DEDICATION

We dedicate the Third Edition of Methods in Cell Biology: The Zebrafish to Wolfgang Driever, Mark C. Fishman, Charles Kimmel, and Christiane N€ ussleinVolhard. Through their foresighted embrace of the zebrafish as a model vertebrate and their pursuit of genetic screens to illuminate vertebrate development, they fostered the emergence of the vibrant zebrafish research community.

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CONTRIBUTORS

Numbers in parentheses indicate the pages on which the author’s contributions begin. R. Craig Albertson (225), Department of Biology, Syracuse University, Syracuse NY, Tufts Univeristy, Boston, Massachusetts James F. Amatruda (19), Department of Pediatrics; Molecular Biology; Internal Medicine, UT Southwestern Medical Center, Dallas, Texas, USA Jennifer L. Anderson (111), Carnegie Institution for Science, Department of Embryology, Baltimore, Maryland, USA Viktoria Andreeva (225), Department of Oral and Maxillofacial Pathology, Tufts Univeristy, Boston, Massachusetts Andrei Avanesov (39), Division of Craniofacial and Molecular Genetics, Tufts University, Massachusetts, USA Maira Carrillo (197), Department of Biological Sciences, University of North Texas, Denton, Texas, USA Juliana D. Carten (111), Carnegie Institution for Science, Department of Embryology, Baltimore, Maryland, USA Joanne Chan (181), Vascular Biology Program and Department of Surgery, Children’s Hospital Boston, Harvard Medical School, Boston, Massachusetts, USA W. James Cooper (225), Department of Biology, Syracuse University, Syracuse NY, Tufts Univeristy, Boston, Massachusetts H. William Detrich III (1), Department of Biology, Northeastern University, Boston, Massachusetts 02115, USA Judith Eisen (143), Institute of Neuroscience, 1254 University of Oregon, Eugene, Oregon Steven A. Farber (111), Carnegie Institution for Science, Department of Embryology, Baltimore, Maryland, USA Robert Gerlai (249), Department of Psychology, University of Toronto, Mississauga, Ontario, Canada Wolfram Goessling (205), Genetics Division, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts, USA; Harvard Stem Cell Institute, Cambridge, Massachusetts, USA Sean Hasso (181), Vascular Biology Program and Department of Surgery, Children’s Hospital Boston, Harvard Medical School, Boston, Massachusetts, USA Pudur Jagadeeswaran (197), Department of Biological Sciences, University of North Texas, Denton, Texas, USA Seongcheol Kim (197), Department of Biological Sciences, University of North Texas, Denton, Texas, USA

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Contributors

Jade Li (39), Departments of Genetics, Yale University School of Medicine, New Haven, Connecticut, USA Jarema Malicki (39), Division of Craniofacial and Molecular Genetics, Tufts University, Massachusetts, USA Sean G. Megason (1), Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115, USA Timothy J. Mitchison (1), Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115, USA Grant I. Miura (161), Division of Biological Sciences, University of California, San Diego, La Jolla, California Trista E. North (205), Department of Pathology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, Massachusetts, USA; Harvard Stem Cell Institute, Cambridge, Massachusetts, USA Nikolaus D. Obholzer (1), Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115, USA Kevin J. Parsons (225), Department of Biology, Syracuse University, Syracuse NY, Tufts Univeristy, Boston, Massachusetts Uvaraj P. Radhakrishnan (197), Department of Biological Sciences, University of North Texas, Denton, Texas, USA Surendra K. Rajpurohit (197), Department of Biological Sciences, University of North Texas, Denton, Texas, USA Iain Shepherd (143), Department of Biology, Emory University Rollins Research Building, Atlanta Georgia David L. Stachura (75), Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego; Department of Cellular and Molecular Medicine, University of California San Diego School of Medicine, La Jolfla, California, USA Zhaoxia Sun (39), Departments of Genetics, Yale University School of Medicine, New Haven, Connecticut, USA David Traver (75), Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego; Department of Cellular and Molecular Medicine, University of California San Diego School of Medicine, La Jolfla, California, USA Daniel Verduzco (19), Department of Pediatrics; Molecular Biology, UT Southwestern Medical Center, Dallas, Texas, USA Martin W€ uhr (1), Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115, USA Pamela C. Yelick (225), Department of Oral and Maxillofacial Pathology, Tufts Univeristy, Boston, Massachusetts Deborah Yelon (161), Division of Biological Sciences, University of California, San Diego, La Jolla, California Shiaulou Yuan (39), Departments of Genetics, Yale University School of Medicine, New Haven, Connecticut, USA

PREFACE

Building on the foundation of our first (1999) and second (2004) editions of Methods in Cell Biology: The Zebrafish, Monte, Len, and I are pleased to continue this Third Edition with Methods in Cell Biology Volume 101, Cellular and Developmental Biology, Part B. In this volume (and its previously released companion, Volume 100, Part A), our contributors present the latest technical advances in the Cell, Developmental and Neural Biology of the zebrafish that have appeared since the second edition. One theme that clearly emerges from these chapters is that the zebrafish is the preeminent vertebrate model for mechanistic cellular studies of developmental processes in vivo. Subsequently, Genetics, Genomics, and Informatics will cover new technologies in Forward and Reverse Genetics, Transgenesis, The Zebrafish Genome and Mapping Technologies, Informatics and Comparative Genomics, and Infrastructure. The Third Edition will also introduce Disease Models and Chemical Screens, two rapidly emerging and compelling applications of the zebrafish. We trust that the Third Edition will prove valuable to both seasoned zebrafish investigators and those who are newly adopting the zebrafish model as part of their research armamentarium. We thank the Series Editors, Leslie Wilson and Paul Matsudaira, and the staff of Elsevier/Academic Press, especially Zoe Kruze, for their enthusiastic support of our Third Edition of Methods in Cell Biology: The Zebrafish. Their help, patience, and encouragement are profoundly appreciated. H. William Detrich, III Monte Westerfield Leonard I. Zon

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CHAPTER 1

Live Imaging of the Cytoskeleton in Early Cleavage-Stage Zebrafish Embryos M. W€ uhr,* N.D. Obholzer,* S.G. Megason,* H.W. Detrich IIIy and T.J. Mitchison* *

Department of Systems Biology, Harvard Medical School, Boston, Massachusetts 02115, USA

y

Department of Biology, Northeastern University, Boston, Massachusetts 02115, USA

Abstract I. Introduction II. Maintaining the Breeding Competence of Zebrafish throughout the Day III. Mounting Zebrafish Embryos for Live Imaging A. Rationale B. Methods IV. Live Imaging of Microtubules in Cleaving Zebrafish Embryos A. Rationale B. Methods C. Results V. Live Imaging of Microfilaments in Cleaving Zebrafish Embryos A. Rationale B. Method C. Results VI. Comparison of Microscopic Techniques for Imaging the Cytoskeleton of Cleaving Zebrafish Embryos VII. Discussion and Future Directions Acknowledgments Appendix A Supplementary Movies References

Abstract The large and transparent cells of cleavage-stage zebrafish embryos provide unique opportunities to study cell division and cytoskeletal dynamics in very large animal cells. Here, we summarize recent progress, from our laboratories and others, METHODS IN CELL BIOLOGY, VOL 101 Copyright 2011. Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-387036-0.00001-3

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on live imaging of the microtubule and actin cytoskeletons during zebrafish embryonic cleavage. First, we present simple protocols for extending the breeding competence of zebrafish mating ensembles throughout the day, which ensures a steady supply of embryos in early cleavage, and for mounting these embryos for imaging. Second, we describe a transgenic zebrafish line [Tg(bactin2:HsENSCONSIN17282-3xEGFP)hm1] that expresses the green fluorescent protein (GFP)-labeled microtubule-binding part of ensconsin (EMTB-3GFP). We demonstrate that the microtubule-based structures of the early cell cycles can be imaged live, with single microtubule resolution and with high contrast, in this line. Microtubules are much more easily visualized using this tagged binding protein rather than directly labeled tubulin (injected Alexa-647-labeled tubulin), presumably due to lower background from probe molecules not attached to microtubules. Third, we illustrate live imaging of the actin cytoskeleton by injection of the actin-binding fragment of utrophin fused to GFP. Fourth, we compare epifluorescence-, spinning-disc-, laser-scanning-, and two-photon-microscopic modalities for live imaging of the microtubule cytoskeleton in early embryos of our EMTB-3GFP-expressing transgenic line. Finally, we discuss future applications and extensions of our methods.

I. Introduction The zebrafish embryo has long been recognized as an excellent model system for molecular–genetic analysis of vertebrate embryonic development (Detrich et al., 1999), one whose advantages complement, and perhaps exceed, those of the mouse (Orkin and Zon, 1997). Forward genetic screens using large-scale zygotic (Driever et al., 1996; Haffter et al., 1996), maternal (Pelegri and Mullins, 2004), and numerous targeted strategies have generated thousands of mutations in the zebrafish that affect all levels of development. Systematic identification and cloning of the mutated genes, whether by candidate (Skromne and Prince, 2008), positional (Bahary et al., 2004), or insertional (Amsterdam and Hopkins, 2004) approaches, has greatly enhanced our understanding of the signaling pathways that regulate expression of the vertebrate body plan. Modern deep sequencing methods will make gene identification even faster. The advantages of the zebrafish for mechanistic studies of developmental processes in vivo at the cellular level have been less well appreciated although the tide is clearly turning (Beis and Stainier, 2006). The remarkable optical clarity of the large blastomeres of the pre-pigmentation embryo facilitates the microscopic examination of cellular processes that underlie morphogenesis. The reduced pigmentation mutant lines nacre (Lister et al., 1999) and casper (White et al., 2008) extend tissue and organ transparency to juvenile and adult animals. As researchers apply transgenic approaches to tag proteins of interest with a fluorescent protein (FP), we foresee a major shift of cellular research to the context of the living fish. Zebrafish excel over amphibian models for live imaging of early development because their meroblastic cleavage separates the transparent blastodisc from the opaque yolk, whereas the

1. Live Imaging of the Cytoskeleton in Early Cleavage-Stage Zebrafish Embryos

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holoblastic cleavage of amphibian embryos renders cells nontransparent at early stages due to distributed yolk particles. The high fecundity of the zebrafish and its low maintenance costs are also major advantages, particularly in comparison to the mouse. Characterization of the cytoskeleton of zebrafish eggs and embryos and its role in morphogenesis of the zygote began in the early 1990s. These studies, which had been stimulated by the pioneering work of J. P. Trinkaus on epiboly and gastrulation in embryos of Fundulus heteroclitus (Betchaku and Trinkaus, 1978; Trinkaus, 1949, 1951), focused initially on microtubules and microfilaments. Using ultraviolet irradiation and antimitotic drugs, Str€ ahle and Jesuthasan (1993) and Solnica-Krezel and Driever (1994) demonstrated that microtubules participate either directly or indirectly in epibolic cell movements, and Jesuthasan and Str€ ahle (1997) concluded that specification of the zebrafish dorsoventral axis required the microtubule-dependent transport of dorsal determinants from the vegetal pole to marginal blastomeres. In recent years, numerous studies have shown that maternal products of the zebrafish oocyte and early embryo are organized, and reorganized, by microtubules and microfilaments during oogenesis and embryogenesis (Dekens et al., 2003; Knaut et al., 2000; Strasser et al., 2008; Theusch et al., 2006; Yabe et al., 2009; reviewed by Lindeman and Pelegri, 2010). To date, the cytoskeletal components of zebrafish oocytes and embryos have generally been analyzed by the application of immunofluorescence light microscopy and/or electron microscopy to fixed preparations. Although methods of fixation to optimize cytoskeletal preservation in embryos have been developed (reviewed by Topczewski and Solnica-Krezel, 2009) and their use has led to important discoveries (reviewed by Lindeman and Pelegri, 2010), research on the function of the cytoskeleton in zebrafish development would benefit enormously from live-cell imaging of fluorescent cytoskeletal proteins. Such studies have revolutionized our understanding of cytoskeleton organization and dynamics in somatic cells, where essentially all cutting-edge cytoskeletal work is now performed using live imaging. Various laboratories have embarked on live-imaging strategies to study cytoskeletal dynamics in zebrafish; examples include microtubule imaging by injection of rhodamine-labeled tubulin into zebrafish zygotes (Li et al., 2006, 2008), the labeling of microfilaments by injection of plasmids that drive the transient expression of the F-actin-binding domain of utrophin fused to mCherry (Andersen et al., 2010), and the creation of a transgenic zebrafish line that expresses a GFP-tagged a-tubulin (Asakawa and Kawakami, 2010). For any experiment aimed at live visualization of the cytoskeleton, the key question is, ‘‘What probe to use?’’’ Useful probes must fulfill multiple criteria: they must not perturb the biology, they must report faithfully on the organization and dynamics of the filament system, they must emit as many photons as possible for as long as possible, and they must provide optimal contrast in the face of background signal from the cytoplasm. The last consideration is often under-appreciated. For all cytoskeleton filaments and their associated binding proteins, there exist at least two protein pools: (1) molecules that are in filaments or binding to filaments and

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(2) molecules that are free in the cytoplasm and often exchange rapidly with the filament-associated pool. In the thick cells of an early embryo, the majority of signal may come from the soluble pool, which lowers the contrast for imaging the filament. For this reason, the best probes for filament visualization in embryos are often not tagged versions of the primary polymer subunits themselves (e.g. tubulin, actin), but rather probes that bind selectively to the polymeric form of the subunit and thus have a lower pool of free proteins. Such polymer-binding probes must be critically evaluated for unwanted interactions; they may tend to stabilize or bundle the polymer if their levels are too high, and they may also bind selectively to certain subsets of the filaments. Despite these caveats, this strategy has been quite successful, and here we discuss its application to microtubule and actin visualization in zebrafish. In this chapter, we describe methods for live imaging of microtubules and microfilaments in cleaving zebrafish embryos, the former by use of a transgenic zebrafish line (W€ uhr et al., 2010) that expresses the GFP-tagged microtubule-binding domain of ensconsin (Faire et al., 1999) and the latter by injection of the actin-binding domain of utrophin bearing a GFP tag (Burkel et al., 2007), respectively. We also compare the quality of images obtained by various optical platforms.

II. Maintaining the Breeding Competence of Zebrafish throughout the Day In the wild, zebrafish spawn at the onset of light in the morning (Detrich et al., 1999). In the lab, this behavior potentially limits the time frame for experimentation on cleavage-stage embryos. Several procedures exist for circumventing this restriction: (1) use of isolation cabinets on light cycles that shift ‘‘morning’’’ for zebrafish mating ensembles to suit the investigator or (2) use of in vitro fertilization, in which females are squeezed and their eggs collected in defined medium or salmon ovarian fluid to prevent activation (Corley-Smith et al., 1999; Sakai et al., 1997). The former technique requires considerable cabinetry, whereas the latter can delay egg activation by at most 6 h (Siripattarapravat et al., 2009) and females require significant time to recover from egg donation. We have found that it is possible to obtain newly fertilized embryos over a large portion of the day (6–8 h) with a simpler routine. We maintain males and females in separate tanks in our zebrafish facility. One day before the experiment, two females and one male are placed together in a crossing cage and allowed to acclimate, which appears to predispose them to mate. The following morning (as defined by the light cycle), males and females are separated immediately after they have spawned sufficient embryos for initiation of experimentation. When more embryos are required, the connubial trio is reunited. We are careful not to separate a threesome for more than 2 h, because they become refractory to further mating that day. Using this method, we typically obtain sufficient embryos to conduct three to four experiments at 2-h intervals in a single day, provided that the fish are at optimal age (4–12 months), well fed, and husbanded.


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III. Mounting Zebrafish Embryos for Live Imaging A. Rationale Proper mounting of cleaving embryos is one of the most important steps for live imaging. To obtain images of high quality, one must immobilize the embryos and place them within the working distance of moderate to high numerical aperture (NA) objectives. In this context, upright and inverted microscopes have different experimental advantages and disadvantages. Mounting of dechorionated embryos on an upright microscope with a water-immersion or air objective is comparatively easy, but one is limited to using objectives of modest NA. This restriction will be reduced as vendors build immersion lenses with increasingly high numerical NA and long working distance. For example, Nikon has introduced a 25 1.1 NA water objective with a 2-mm working distance (Nikon Inc.), which delivers an extraordinary increase in performance at low magnification. Unfortunately, such lenses are extremely expensive and may require special adapters. In contrast, mounting embryos for inverted microscopy is more difficult but permits the use of oil-immersion, high NA objectives. In our experience, the best method to immobilize a dechorionated embryo is to place it in the pocket of an agarose specimen chamber cast on a Petri dish. The lateral dimensions of the pockets are slightly smaller than the diameter of an embryo (Fig. 1A) so that friction between the gently compressed embryo and the walls of the pocket resists embryo movement. To cast a chamber with pockets that can accommodate embryos of differing size, we use a plastic mold that creates squares with sides of 550–700 mm; the depth of each pocket is 400 mm. Mounting of the embryo is performed on a dissecting microscope after which the Petri dish is transferred to the microscope used for imaging. The increased difficulty of mounting an embryo for inverted microscopy arises from its natural tendency to rotate so that the heavy yolk faces downward, that is, opposite to the desired yolk-up orientation in the mounting pocket. In addition, the specimen chamber must be cast on the glass cover slip of a suitable culture dish (Fig. 1B). The key, in our experience, is to use an agarose pocket of optimal dimensions, so the embryo is prevented from rotating but not overly compressed.

B. Methods 1. Machine the embryo mounting mold from a suitable plastic or metal (e.g. plexiglass, aluminum) to the dimensions shown in Fig. 1A. 2. Pour melted 2% agarose in egg water (Westerfield, 2007) into a Petri dish. Insert the mold into the agarose solution and put a weight on top. Allow the agarose to set at 4 C. If mounting for inverted microscopy, use a coverslip-bottom culture dish (e.g. MatTek Corp., Ashland, MA, USA). 3. Remove the mold and add egg water and embryos into the dish. 4. Using the dissecting microscope for visualization, dechorionate embryos with two sharp No. 5 forceps.

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[(Fig._1)TD$IG]

Fig. 1 Mounting of cleavage-stage zebrafish embryos. (A) Mold used to prepare agarose mounting pockets for cleavage-stage embryos. Pockets of differing dimensions provide flexibility in mounting of embryos of heterogeneous size. (B) Configuration of embryos for observation using an upright microscope equipped with a water-immersion (dipping) objective (left panel) or for imaging via inverted microscopy and an oil-immersion objective (right panel).

5. With a blunt glass rod, maneuver an embryo into a square chamber whose dimensions are slightly smaller than the embryo’s diameter. Orient the embryo as desired and, if necessary, add low-melting-point agarose (0.8%) to fix the embryo in position. Position additional embryos in remaining pockets as desired. 6. To reduce swaying when transferring, remove most of the egg water from the dish, leaving just enough to cover the embryo. Transfer the dish to the microscope stage and add egg water to a level sufficient for immersing the objective (Fig. 1B). 7. For inverted microscopy, carefully transfer the culture dish to the microscope stage and bring the objective into oil contact with the dish’s coverslip (Fig. 1B). Add egg water to cover the embryos so that they do not dehydrate during imaging.

IV. Live Imaging of Microtubules in Cleaving Zebrafish Embryos A. Rationale Li et al. (2006, 2008) demonstrated real-time imaging of microtubules in cleaving zebrafish embryos by injection of rhodamine-labeled tubulin at the one-cell stage. Due to the thickness of early zebrafish blastomeres and the large proportion of


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rhodamine–tubulin that remains monomeric, the signal-to-noise ratio of fluorescent microtubule polymer relative to the fluorescent subunit pool is substantially lower than that achieved by comparable injection of thin, adherent tissue culture cells (Zhai et al., 1996). Fig. 2A (right panel) shows an image from our laboratory of a cleaving zebrafish embryo whose microtubules are labeled by incorporation of injected Alexa-647-labeled tubulin. Although the contrast is unusually high, astral microtubules are barely visible over the background from soluble tubulin. To obtain higher contrast images of microtubule dynamics and organization in cleaving zebrafish blastomeres and to circumvent the injection step, we (W€ uhr et al., 2010) generated a transgenic fish line, Tg(bactin2:HsENSCONSIN17-282-3xEGFP)hm1, that expresses the microtubule-binding domain of ensconsin fused to three sequential GFP moieties (EMTB-3GFP). This probe was first tested in tissue culture cells, where it was shown to associate tightly but dynamically with microtubules without

[(Fig._2)TD$IG]

Microtubule imaging in cleaving embryos from the Tg(bactin2:HsENSCONSIN17-282-3xEGFP)hm1 zebrafish line. (A) Transgenic, EMTB-3GFP-expressing embryos were injected with Alexa-647-labeled tubulin. The GFP and Alexa-647 signals were imaged simultaneously as described in Section IV.B.3. (B) Time lapse images of microtubules labeled by EMTB3GFP (enlargements of the boxed region of panel A, left side). The dynamic instability of the ends of individual microtubules can be followed; the green arrows delineate a growing microtubule end, whereas the red arrows show a shortening microtubule. (C) Ratiometric image generated as the composite of the two images shown in panel A. The image has been pseudocolored to differentiate regions labeled preferentially by EMTB-3GFP (red) or by Alexa-647-labeled tubulin (blue). See Section IV.C for further details. (D) Simultaneous labeling of microtubules and chromatin in embryos expressing EMTB-3GFP and H2BmCherry. See Section IV.C for details. (See Plate no. 1 in the Color Plate Section.)

Fig. 2

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perturbing either microtubule organization or dynamics when expressed at levels appropriate for imaging (Faire et al., 1999). Later, EMTB-3GFP was adapted for imaging microtubules in echinoderm embryos, where its increased contrast relative to directly labeled tubulin was a compelling advantage (von Dassow et al., 2009). In that work, the probe was introduced by mRNA injection at the one-cell stage, which precluded live imaging of the first division. We introduced this probe into zebrafish by making a transgenic line. In addition to permitting visualization immediately after egg spawning and fertilization due to maternal expression of the transgene, the transgenic approach has the advantage that we know the probe has not perturbed embryonic development since the fish line is fully fertile. Below we compare the two methods directly by injection of Alexa-647-labeled tubulin into the transgenic zebrafish line. We have also used EMTB fused to a single GFP and expressed in bacteria to visualize microtubules live in Xenopus egg extracts, demonstrating the versatility of ensconsin-based probes.

B. Methods

1. Generation of Transgenic Zebrafish Lines The transgenic line, Tg(bactin2:HsENSCONSIN17-282-3xEGFP)hm1, was created in an unspecified wild-type strain by use of the Tol2Kit (Kawakami, 2004; Kwan et al., 2007; Urasaki et al., 2006; W€ uhr et al., 2010). EMTB-3GFP expression is driven by the beta actin promoter, chosen for its high expression levels. Beginning with eight founders, we selected progeny that gave the highest expression levels without detectable developmental toxicity. The line is now in its third generation, the transgene is mostly stably transmitted, and expression of EMTB-3GFP remains robust. EMTB-3GFP expression levels do tend to decline with generational passage of the transgene, and we compensate for this by selecting adult females that express the brightest eggs for propagation. Using identical methods, we have created a transgenic zebrafish line in wild-type strain AB that expresses human histone H2B fused in frame to mCherry2 for visualization of chromatin dynamics.

2. Preparation of Alexa-647-Labeled Tubulin and Embryo Microinjection Tubulin was purified from calf brain and labeled with Alexa647-succinimideester (Invitrogen) as described by Hyman et al. (1991). The ratio of fluorophore to tubulin dimer was 0.7. Tg(bactin2:HsENSCONSIN17-282-3xEGFP)hm1 zebrafish were mated, and, shortly after fertilization, 5 nL of labeled tubulin (11 mg/ml) were injected through the yolk into the blastodiscs of embryos.

3. Laser-Scanning Confocal Microscopy Images were recorded using a Zeiss LSM 710 inverted microscope equipped with a 63 plan-apochromat objective (NA = 1.4). The pinhole was set at 63 mm, the


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pixel size was 0.11 mm, and pixel dwell time was 0.79 ms. Specimens were illuminated simultaneously by argon (488 nm, 25 mW) and helium–neon (633 nm, 5 mW) lasers. The emission spectra of GFP and Alexa 647 were recorded from 492 to 598 nm and from 637 to 755 nm, respectively. Images of a single focal plane were collected at 7.7-s intervals.

C. Results Fig. 2A compares the labeling of spindle microtubules by EMTB-3GFP and by Alexa-647-tubulin in early anaphase of the second mitosis in an injected, transgenic embryo: the left panel shows the EMTB-3GFP signal, whereas the right panel shows the Alexa-647 signal. Movie S1 shows the same embryo in both imaging modalities as anaphase commences. Spindle microtubules were brightly labeled in the green channel, and their contrast with respect to the background was high. Neither spindle morphology nor function appears to be perturbed by binding of EMTB-3GFP to spindle microtubules (see Movie S1), as expected since the transgenic line is fertile. Furthermore, the dynamic instability (Mitchison and Kirschner, 1984) of individual microtubules at the front of asters as they expanded in telophase was readily detected (see time-lapse imagery of Fig. 2B, which shows enlargements of the boxed region of 2A). In contrast, the same spindle observed in the Alexa-647 channel was less clearly visualized; the central spindle microtubules were bright, but the background fluorescence of the cytoplasm was high and individual microtubule ends in the asters could not be identified with confidence. The clarity of microtubule labeling by the transgenic zebrafish line is striking, but one may ask whether EMTB-3GFP, which interacts with microtubules noncovalently, faithfully delineates all of the microtubules throughout the spindle.1 The ensconsin (GFP) and tubulin (Alexa-647) signals correlate well, but subtle differences are apparent that cannot be explained by lower background fluorescence. To compare microtubule labeling by the two approaches, we generated a composite, ratiometric image from the two images of Fig. 2A. Fig. 2C shows that EMTB-3GFP preferentially labels certain microtubule populations, which are shown in red in the pseudocolored ratio image. These include the distal ends (distal with respect to the centrosome) of astral microtubules and microtubules of the furrow microtubule array [microtubules to the left of the spindle (Danilchik et al., 2003)]. Astral microtubules proximal to the centrosomes label equivalently with the two probes (green pseudocolor). EMTB-3GFP staining of the aster interaction zone (W€ uhr et al., 2010), where the two asters meet, is very low (blue pseudocolor) compared to the signal in the tubulin channel, suggesting the probe is selectively excluded from these microtubules. Possible non-mutually exclusive explanations for the differential labeling of microtubules by EMTB-3GFP could be: (1) EMTB-3GFP’s local 1

In this discussion we make the explicit assumption that labeling of microtubules by Alexa-647-tubulin is uniform throughout the spindle.

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concentration within the asters and the aster interaction zones might not be sufficiently high to saturate its binding sites on the polymer, whereas this condition is met for the dispersed microtubule ends at the astral peripheries; (2) EMTB-3GFP may compete with other microtubule-associated proteins for binding to specific subsets or subregions of spindle microtubules; and/or (3) the affinity of EMTB-3GFP may be altered by regional regulation of posttranslational modification. We are currently working on evaluating these hypotheses, with the aim of engineering a modified probe with less differential binding. However, for most purposes the current probe provides excellent microtubule imaging and is clearly superior to directly labeled tubulin. For visualization of the advancing front of astral microtubules, selective binding of the probe is even advantageous. To observe microtubule and chromatin dynamics simultaneously during embryonic cleavage, we crossed the EMTB-3GFP line (hm1) with a beta-actin:H2BmCherry2 line [Tg(ba:h2b-mCherry2)hm13] (Fig. 2D, Movie S2). Fig. 2D shows that the mCherry2-tagged histone labels the chromosomes at the metaphase plate (red signal), and Movie S2 shows a blastomere undergoing a complete mitotic cycle of chromosome condensation, spindle assembly, and chromosome partition. The high signal-to-noise ratios of the EMTB-3GFP-labeled microtubules and of the mCherry2-H2B-labeled chromosomes in these movies should facilitate quantitative analysis of cleavage in the large blastomeres of the meroblastic zebrafish embryo.

V. Live Imaging of Microfilaments in Cleaving Zebrafish Embryos A. Rationale Live imaging of microfilaments in the large blastomeres of the zebrafish embryo is even more problematic than live imaging of microtubules, most likely because the concentration of soluble, unpolymerized actin is very high compared to polymerized actin in fibers. In a comparable embryo (Xenopus laevis), actin is present at 20 mM, and most is bound to sequestering proteins (Rosenblatt et al., 1995). Sequestered monomer probably contributes to very high background staining if actin is imaged via immunofluorescence of labeled actin monomers. Rhodaminelabeled phalloidin, which binds only to F-actin with extremely high selectivity, has been used to study microfilaments during cleavage of zebrafish embryos (Li et al., 2008; Theusch et al., 2006), but this approach typically requires fixation and restricts fixation methods to those that preserve filament structure (aldehyde fixation works, organic solvent fixation does not). When labeled phalloidin has been used to image microfilaments in living zebrafish embryos (Li et al., 2008), the probe must be restricted to very low concentrations and thus low signal since phalloidin is in fact a toxin derived from the death cap mushroom. Bement and co-workers developed several FP-tagged probes for microfilaments based on the actin-binding calponin homology domain of utrophin (Utr-CH)(Burkel et al., 2007). They showed that


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transient expression of GFP-Utr-CH in Xenopus oocytes by injection of synthetic mRNA or plasmid constructs reports on the distribution of F-actin without perturbing actin dynamics and is less toxic than phalloidin. It is possible, even likely, that this probe binds selectively to a certain population of filaments, but this is difficult to determine when we lack alternative methods for filament visualization. We attempted to generate a transgenic fish line that would express Utr-CH-GFP, but we were unable to establish founder fish, most likely due to probe toxicity during development. As an alternative, we developed a protocol for injection of bacterially expressed Utr-CH-GFP (kindly provided by David Burgess, Boston College) into the one-cell stage of the zebrafish embryo. With this probe we could visualize cortical microfilaments in living embryos with excellent contrast.

B. Method His-tagged Utr-CH-GFP was expressed in Escherichia coli, purified via chromatography on nickel columns, and flash frozen in 150 mM aspartic acid and 10 mM HEPES solution (pH 7.2–7.3) in the laboratory of David Burgess. Shortly after fertilization, zebrafish embryos were injected through the yolk into the blastodisc with 2 nL of utrophin–GFP (1 mg/ml) and then mounted for upright microscopy as described in Section III.B. Images were recorded using a Zeiss LSM 710 upright microscope equipped with a 20 water-immersion objective (plan-apochromat DIC, NA = 1.0). The pinhole was set at 32 mm, the pixel size was 0.59 mm, and pixel dwell time was 1.58 ms. Specimens were illuminated with a 25-mWArgon laser at 488 nm. Emission spectra were recorded from 492 to 598 nm. Images of a single focal plane were collected at 41-s intervals.

C. Results Figure 3 shows the lateral views of a one-cell zebrafish embryo undergoing cytokinesis; the boxed regions are enlarged and shown at higher contrast below. The cortical actin filaments are brightly labeled by Utr-CH-GFP as the cleavage furrow develops. Cleavage appears to be unaffected by the utrophin probe (Movie S3, the same embryo). At t = 0 min, the cell is in late telophase, and Utr-CH-GFP fluorescence marks the aster–aster interaction zone (W€ uhr et al., 2010), where cytokinesis will cleave the cell. By t = 26 min, the daughter cells have re-entered mitosis, and Utr-CH-GFP stains comet tails behind rapidly moving vesicles (see Movie S3 beginning at t = 18 min). These comet tails in mitotic cells presumably represent Arp2/3 nucleated assemblies akin to Listeria comet tails, which have been seen before in live embryos (Taunton et al., 2000; Velarde et al., 2007). Comet tail assembly and vesicle movement were abolished by treatment of embryos with the actin-depolymerizing agent cytochalasin B but were insensitive to the anti-microtubule drug nocodazole (data not shown). Although further validation of our utrophin-based labeling strategy is required, we consider it likely

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[(Fig._3)TD$IG]

Fig. 3

F-actin imaging in an embryo injected with Utr-CH-GFP at the one-cell stage. Viewed from the side, the cortex is brightly labeled. The enlargements (shown at higher contrast below the top panels) illustrate the dynamics of the microfilaments. At t = 0 min, the cell is in late telophase, and Utr-CH-GFP staining marks the aster-aster interaction zone (yellow arrows). As the daughter cells from the first division re-enter mitosis (t = 26 min), Utr-CH-GFP-labeled comets propel vesicles (e.g. red arrow) whose movement is actin-dependent (see Section V.C for details). (See Plate no. 2 in the Color Plate Section.)

to be a useful, non-perturbing method for live imaging of the structure and function of the actin cytoskeleton in zebrafish embryos. One important question for future work is to what extent this probe reports on localization of all actin filaments versus a subset with particular structure or biochemistry.

VI. Comparison of Microscopic Techniques for Imaging the Cytoskeleton of Cleaving Zebrafish Embryos Table I summarizes the advantages and disadvantages of four fluorescence-imaging modalities we tested and provides representative micrographs obtained by each. We note that these comments apply to the instrument we used and may not represent fundamental limitations. For example, new gallium arsenide and avalanche photodiode detectors for scanning microscopes may increase sensitivity and lower noise and photobleaching, albeit at additional cost. In the two-cell embryo imaged by

13


Table I Imaging the zebrafish microtubule cytoskeleton during cleavage: Advantages and disadvantages of four fluorescenceimaging modalities Type

Advantages

Disadvantages

Epifluorescence microscopy

- Least expensive - Fast - Efficient collection of photons (low bleaching and phototoxicity) - CCD camera (low noise, high sensitivity)

- Blurry, especially in thick specimens at high magnification (low contrast) - Low depth penetration - Low contrast of microtubules

Spinning disc confocal microscopy (SDCM)

- Fast - Less expensive than LSCM or 2PM - Faster than LSCM and 2PM - Fairly efficient collection of photons (low bleaching and phototoxicity) - Reasonable background suppression - CCD camera - Can generate optical sections - Can generate optical sections - Good depth penetration - Good background suppression

- Fixed pinhole size, only optimal for certain objectives - Cross-talk between pinholes - Higher background than LSM especially in a thick specimen, leading to lower contrast images

- Can generate optical sections - Excellent depth penetration - Very good background suppression

- Slow scan rate - PMT is noisy (8-bit) - Expensive - Low brightness, fast bleaching, phototoxicity is likely (probably because common fluorophores are optimized for one-photon excitation)

Example

Movie S4

Laser-scanning confocal microscopy (LSCM)

Movie S5 - Slow scan rate - PMT is noisy (8-bit) - Expensive - Very few photons collected (bright signal required, fast bleaching, phototoxicity is likely without care to limit exposure) Movie S6

Two-photon microscopy (2PM)

Movie S6

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conventional epifluorescence, the spindles of each blastomere are quite blurry due to high background signal from out-of-focus fluorescence. Although basic measurements of mitosis, such as rate of spindle assembly, spindle size, and spindle orientation, could be made (see Movie S4), the dynamics of single microtubules, or even bundles, cannot be resolved. Spinning disc confocal microscopy provides greater clarity and single microtubule resolution of early mitotic spindles (Table I, Movie S5) provided that they are relatively close to the surface of the embryo. Spinning disc confocal often seems to provide an advantage over scanning laser confocal for live imaging of the cytoskeleton in tissue culture cells due to lower photobleaching and in some cases superior signal to noise. However, the lack of depth of penetration is a problem for application of current Yokogawa spinning discs to zebrafish embryos. We look forward to development of new spinning disc units with smaller pinholes that are optimized for lower magnification work at depth. For the large, cleaving blastomeres of the zebrafish embryo, laser-scanning confocal microscopy and twophoton microscopy were clearly superior because of their greater depth of penetration. With care to limit exposure, photobleaching and phototoxicity were not a problem with the one-photon modality. Both produced images of mitotic spindles with very high spatial resolution and contrast (Table I, Movie S6). The dynamics of individual microtubules were easily observed. Our current method of choice is laserscanning microscopy with one-photon excitation. This is partly due to the faster bleaching of GFP caused by two-photon excitation, but of greater importance is the higher signal-to-noise that was obtained with one-photon excitation (Movie S6). We do not understand to what extent these factors represent fundamental advantages of one-photon excitation versus limitations of the particular instruments we used.

VII. Discussion and Future Directions In this chapter, we describe methods to image microtubules and actin filaments in the thick cells of living cleavage-stage zebrafish embryos. Our methods make use of FP-tagged filament-binding proteins, EMTB-3GFP for microtubules and UtrCH-GFP for microfilaments, that appear not to affect the dynamics or organization of the respective polymers. These probes yield superior contrast during live imaging when compared to filament labeling by fluorescently derivatized polymer subunits themselves (i.e. tubulin, actin), presumably because the free pools of the binding proteins are much lower than those of the filament subunits. Furthermore, we have successfully developed a transgenic zebrafish line that expresses EMTB-3GFP and shown that it yields valuable information about microtubule organization and function in cleavage-stage zebrafish embryos (W€ uhr et al., 2010). For analysis of microtubule function at later stages of development, we suggest that the EMTB-3GFP probe be introduced into strains lacking pigmentation [e.g. nacre (Lister et al., 1999) and casper (White et al., 2008)]. A disadvantage of EMTB-3GFP is that the probe turns over rapidly on microtubules (Bulinski et al., 2001), which prevents its use to measure microtubule turnover or sliding by photoactivation experiments (Mitchison, 1989). To


15

enable such measurements, we suggest the creation of a zebrafish line that expresses tubulin linked to a photoconvertible FP (McKinney et al., 2009). Although we have not been able to generate a transgenic line that constitutively expresses Utr-CH-GFP, suggesting subtle toxic effects of this probe expressed at high levels, we consider it probable that such a line can be created, perhaps by using a weaker, or inducible, promoter. Indeed, we envision a bright future for live analysis of cellular dynamics of all kinds by use of transgenic zebrafish that express appropriate FP-tagged probes. We chose to study the zebrafish embryo not only for its potential in understanding cytoskeletal function during development, but also because the blastomeres created during cleavage are among the largest of vertebrate cells. One of our goals is to understand how the cytoskeleton scales with cell size to solve the physical challenges of organizing large cells (W€ uhr et al., 2008). To this end, a combination of the unique experimental advantages of Xenopus egg extracts and transgenic zebrafish embryos is likely to yield important experimental synergisms. Xenopus egg extracts provide the opportunity to observe cytoskeletal function in vitro with single molecule resolution (Needleman et al., 2010), and the system can be easily titrated with proteins and drugs. Conversely, the zebrafish embryo provides experimental read out from a truly in vivo system. The two systems, used in combination, are likely to lead to rapid advances in our knowledge of cytoskeletal function and other cellular processes. Acknowledgments We thank David Burgess for generous gift of purified Utr-CH-GFP. H.W.D. was supported by NSF grant ANT-0635470. N.D.O and S.G.M. were supported by NHGRI P50 HG004071 and NIDCD R01 DC010791. This work was supported by the National Institutes of Health (NIH) grant GM39565.

Appendix A. Supplementary Movies Supplementary data associated with this chapter can be found, in the online version, at http://www.elsevierdirect.com/companions/9780123870360. References Amsterdam, A., and Hopkins, N. (2004). Retroviral-mediated insertional mutagenesis in zebrafish. In ‘‘Methods in Cell Biology,’’ (H. W. Detrich III, M. Westerfield, and L. I. Zon, eds.), Vol. 77, pp. 3–20. Elsevier, San Diego. Andersen, E., Asuri, N., Clay, M., and Halloran, M. (2010). Live imaging of cell motility and actin cytoskeleton of individual neurons and neural crest cells in zebrafish embryos. J. Vis. Ex 36 DOI: 10.3791/1726. Asakawa, K., and Kawakami, K. (2010). A transgenic zebrafish for monitoring in vivo microtubule structures. Dev. Dyn. 239, 2695–2699. Bahary, N., Davidson, A., Ransom, D., Shepard, J., Stern, H., Trede, N., Zhou, Y., Barut, B., Zon, L. I. (2004). The Zon laboratory guide to positional cloning in zebrafish. In ‘‘Methods in Cell Biology,’’ (H. W. Detrich III, M. Westerfield, and L. I. Zon, eds.), Vol. 77, pp. 305–329. Elsevier, San Diego.

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M. W€ uhr et al. Beis, D., and Stainier, D. Y. (2006). In vivo cell biology: following the zebrafish trend. Trends Cell. Biol. 16, 105–112. Betchaku, T., and Trinkaus, J. P. (1978). Contact relations, surface activity, and cortical microfilaments of marginal cells of the enveloping layer and of the yolk syncytial and yolk cytoplasmic layers of Fundulus before and during epiboly. J. Exp. Zool. 206, 381–426. Bulinski, J. C., Odde, D. J., Howell, B. J., Salmon, T. D., and Waterman-Storer, C. M. (2001). Rapid dynamics of the microtubule binding of ensconsin in vivo. J. Cell Sci. 114, 3885–3897. Burkel, B. M., von Dassow, G., and Bement, W. M. (2007). Versatile fluorescent probes for actin filaments based on the actin-binding domain of utrophin. Cell Motil. Cytoskeleton 64, 822–832. Corley-Smith, G. E., Brandhorst, B. P., Walker, C., and Postlethwait, J. H. (1999). Production of haploid and diploid androgenetic zebrafish (including methodology for delayed in vitro fertilization). In ‘‘Methods in Cell Biology,’’ (H. W. Detrich III, M. Westerfield, and L. I. Zon, eds.), Vol. 59, pp. 45–60. Elsevier, San Diego. Danilchik, M. V., Bedrick, S. D., Brown, E. E., and Ray, K. (2003). Furrow microtubules and localized exocytosis in cleaving Xenopus laevis embryos. J. Cell Sci. 116, 273–283. Dekens, M. P., Pelegri, F. J., Maischein, H. M., and N€ usslein-Volhard, C. (2003). The maternal-effect gene futile cycle is essential for pronuclear congression and mitotic spindle assembly in the zebrafish zygote. Development 130, 3907–3916. Detrich, H. W., Westerfield, M., and Zon, L. I. (1999). Overview of the zebrafish system. In ‘‘Methods in Cell Biology,’’ (H. W. Detrich III, M. Westerfield, and L. I. Zon, eds.), Vol. 59, pp. 3–10. Elsevier, San Diego. Driever, W., Solnica-Krezel, L., Schier, A. F., Neuhauss, S. C. F., Malicki, J., Stemple, D. L., Stainier, D. Y. R., Zwartkruis, F., Abdelilah, S., Rangini, Z., et al. (1996). A genetic screen for mutations affecting embryogenesis in zebrafish. Development 123, 37–46. Dumont, S., and Mitchison, T. J. (2009). Compression regulates mitotic spindle length by a mechanochemical switch at the poles. Curr. Biol. 19, 1086–1095. Faire, K., Waterman-Storer, C. M., Gruber, D., Masson, D., Salmon, E. D., Bulinski, J. C. (1999). E-MAP115 (ensconsin) associates dynamically with microtubules in vivo and is not a physiological modulator of microtubule dynamics. J. Cell Sci. 112, 4243–4255. Haffter, P., Granato, M., Brand, M., Mullins, M. C., Hammerschmidt, M., Kane, D. C., Odenthal, J., van Eeden, F. J. M., Jiang, Y.-J., Heisenberg, C.-P., et al. (1996). The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio. Development 123, 1–36. Hyman, A., Drechsel, D., Kellogg, D., Salser, S., Sawin, K., Steffen, P., Wordeman, L., Mitchison, T. (1991). Preparation of modified tubulins. Methods Enzymol. 196, 478–485. Jesuthasan, S., and St€ ahle, U. (1997). Dynamic microtubules and specification of the zebrafish embryonic axis. Curr. Biol. 7, 31–42. Kawakami, K. (2004). Transgenesis and gene trap methods in zebrafish by using the Tol2 transposable element. In ‘‘Methods in Cell Biology,’’ (H. W. Detrich III, M. Westerfield, and L. I. Zon, eds.), Vol. 77, pp. 201–222. Elsevier, San Diego. Knaut, H., Pelegri, F., Bohmann, K., Schwarz, H., and N€ usslein-Volhard, C. (2000). Zebrafish vasa RNA but not its protein is a component of the germ plasm and segregates asymmetrically before germline specification. J. Cell Biol. 149, 875–888. Kwan, K. M., Fujimoto, E., Grabher, C., Mangum, B. D., Hardy, M. E., Campbell, D. S., Parant, J. M., Yost, H. J., Kanki, J. P, and Chien, C. B. (2007). The Tol2kit: a multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev. Dyn. 236, 3088–3099. Li, W. M., Webb, S. E., Chan, C. M., and Miller, A. L. (2008). Multiple roles of the furrow deepening Ca2+ transient during cytokinesis in zebrafish embryos. Dev. Biol. 316, 228–248. Li, W. M., Webb, S. E., Lee, K. W., and Miller, A. L. (2006). Recruitment and SNARE-mediated fusion of vesicles in furrow membrane remodeling during cytokinesis in zebrafish embryos. Exp. Cell Res. 312, 3260–3275.


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Lindeman, R. E., and Pelegri, F. (2010). Vertebrate maternal-effect genes: Insights into fertilization, early cleavage divisions, and germ cell determinant localization from studies in the zebrafish. Mol. Reprod. Dev. 77, 299–313. Lister, J. A., Robertson, C. P., Lepage, T., Johnson, S. L., and Raible, D. W. (1999). nacre encodes a zebrafish microphthalmia-related protein that regulates neural-crest-derived pigment cell fate. Development 126, 3757–3767. McKinney, S. A., Murphy, C. S., Hazelwood, K. L., Davidson, M. W., and Looger, L. L. (2009). A bright and photostable photoconvertible fluorescent protein. Nat. Methods 6, 131–133. Mitchison, T. J. (1989). Polewards microtubule flux in the mitotic spindle: evidence from photoactivation of fluorescence. J. Cell Biol. 109, 637–652. Mitchison, T., and Kirschner, M. (1984). Dynamic instability of microtubule growth. Nature 312, 237–242. Needleman, D. J., Groen, A., Ohi, R., Maresca, T., Mirny, L., Mitchison, T. (2010). Fast microtubule dynamics in meiotic spindles measured by single molecule imaging: evidence that the spindle environment does not stabilize microtubules. Mol. Biol. Cell 21, 323–333. Orkin, S. H., and Zon, L. I. (1997). Genetics of erythropoiesis: induced mutations in mice and zebrafish. Annu. Rev. Genet. 31, 33–60. Pelegri, F., and Mullins, M. C. (2004). Genetic screens for maternal-effect mutations. In ‘‘Methods in Cell Biology,’’ (H. W. Detrich III, M. Westerfield, and L. I. Zon, eds.), Vol. 77, pp. 21–51. Elsevier, San Diego. Rosenblatt, J., Peluso, P., and Mitchison, T. J. (1995). The bulk of unpolymerized actin in Xenopus egg extracts is ATP-bound. Mol. Biol. Cell 6, 227–236. Sakai, N., Burgess, S., and Hopkins, N. (1997). Delayed in vitro fertilization of zebrafish eggs in Hank’s saline containing bovine serum albumin. Mol. Mar. Biol. Biotechnol. 6, 84–87. Siripattarapravat, K., Busta, A., Steibel, J. P., and Cibelli, J. (2009). Characterization and in vitro control of MPF activity in zebrafish eggs. Zebrafish 6, 97–105. Skromne, I., and Prince, V. E. (2008). Current perspectives in zebrafish reverse genetics: moving forward. Dev. Dyn. 237, 861–882. Solnica-Krezel, L., and Driever, W. (1994). Microtubule arrays of the zebrafish yolk cell: organization and function during epiboly. Development 120, 2443–2455. Str€ ahle, U., and Jesuthasan, S. (1993). Ultraviolet irradiation impairs epiboly in zebrafish embryos: evidence for a microtubule-dependent mechanism of epiboly. Development 119, 909–919. Strasser, M. J., Mackenzie, N. C., Dumstrei, K., Nakkrasae, L. I., Stebler, J., Raz, E. (2008). Control over the morphology and segregation of zebrafish germ cell granules during embryonic development. BMC. Dev. Biol. 8, 58. Taunton, J., Rowning, B. A., Coughlin, M. L., Wu, M., Moon, R. T., Mitchison, T. J., Larabell, C. A. (2000). Actin-dependent propulsion of endosomes and lysosomes by recruitment of N-WASP. J. Cell Biol. 148, 519–530. Theusch, E. V., Brown, K. J., and Pelegri, F. (2006). Separate pathways of RNA recruitment lead to the compartmentalization of the zebrafish germ plasm. Dev. Biol. 292, 129–141. Topczewski, J., and Solnica-Krezel, L. (2009). Cytoskeletal dynamics of the zebrafish embryo. In ‘‘Essential Zebrafish Methods, Part A: Cell and Developmental Biology,’’ (H. W. Detrich III, M. Westerfield, and L. I. Zon, eds.), pp. 133–157. Elsevier, San Diego. Trinkaus, J. P. (1949). The surface gel layer of Fundulus eggs in relation to epiboly. Proc. Natl. Acad. Sci. U. S. A. 35, 218–225. Trinkaus, J. P. (1951). A study of mechanisms of epiboly in the egg of Fundulus heteroclitus. J. Exp. Zool. 118, 269–319. Urasaki, A., Morvan, G., and Kawakami, K. (2006). Functional dissection of the Tol2 transposable element identified the minimal cis-sequence and a highly repetitive sequence in the subterminal region essential for transposition. Genetics 174, 639–649. Velarde, N., Gunsalus, K. C., and Piano, F. (2007). Diverse roles of actin in C. elegans early embryogenesis. BMC Dev. Biol. 7, 142.

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M. W€ uhr et al. Von Dassow, G., Verbrugghe, K. J., Miller, A. L., Sider, J. R., and Bement, W. M. (2009). Action at a distance during cytokinesis. J. Cell Biol. 187, 831–845. Westerfield, M. (2007). The zebrafish book: A guide for the laboratory use of zebrafish (Danio rerio) University of Oregon Press, Eugene, Oregon. White, R. M., Sessa, A., Burke, C., Bowman, T., LeBlanc, J., Ceol, C., Bourque, C., Dovey, M., Goessling, W., Burns, C. E., et al. (2008). Transparent adult zebrafish as a tool for in vivo transplantation analysis. Cell Stem Cell 2, 183–189. W€ uhr, M., Chen, Y., Dumont, S., Groen, A. C., Needleman, D. J., Salic, A., Mitchison, T. J. (2008). Evidence for an upper limit to mitotic spindle length. Curr. Biol. 18, 1256–1261. W€ uhr, M., Tan, E. S., Parker, S. K., Detrich III, H. W., and Mitchison, T. J. (2010). A model for cleavage plane determination in early amphibian and fish embryos. Curr. Biol. 20, 2040–2045. Yabe, T., Ge, X., Lindeman, R., Nair, S., Runke, G., Mullins, M. C., Pelegri, F. (2009). The maternal-effect gene cellular island encodes aurora B kinase and is essential for furrow formation in the early zebrafish embryo. PLoS. Genet. 5, e1000518. Zhai, Y., Kronebusch, P. J., Simon, P. M., and Borisy, G. G. (1996). Microtubule dynamics at the G2/M transition: abrupt breakdown of cytoplasmic microtubules at nuclear envelope breakdown and implications for spindle morphogenesis. J. Cell Biol. 135, 201–214.

CHAPTER 2

Analysis of Cell Proliferation, Senescence, and Cell Death in Zebrafish Embryos Daniel Verduzco*,y and James F. Amatruda*,y,z *

Departments of Pediatrics, UT Southwestern Medical Center, Dallas, Texas, USA

y

Molecular Biology, UT Southwestern Medical Center, Dallas, Texas, USA

z

Internal Medicine, UT Southwestern Medical Center, Dallas, Texas, USA

Abstract I. Introduction: The Cell Cycle in Zebrafish A. Forward-Genetic Screens II. Zebrafish Embryo Cell-Cycle Protocols A. Analysis of Cell Proliferation and Mitosis B. Analysis of DNA Damage, Senescence, and Apoptosis C. In Situ Hybridization III. Screening for Chemical Suppressors of Zebrafish Cell-Cycle Mutants IV. Conclusions V. Reagents and Supplies Acknowledgments References

Abstract Proper control of cell proliferation is critical for normal development, growth, differentiation, and tissue homeostasis. Dysregulation of cell division and cell death underlies almost all cancers, and contributes to the pathology of birth defects and degenerative diseases. The zebrafish has proved to be an excellent system for elucidating the roles of the cell cycle in normal development, and ways in which dysregulation of cell proliferation contributes to disease. This chapter describes the methods for studying the cell cycle in zebrafish embryos, including protocols to examine cell proliferation, DNA damage, senescence, and cell death.

METHODS IN CELL BIOLOGY, VOL 101 Copyright 2011. Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-387036-0.00002-5

20

Daniel Verduzco and James F. Amatruda

I. Introduction: The Cell Cycle in Zebrafish In multicellular organisms, the cell cycle is a fundamental feature of cellular physiology that is critical for normal development, organogenesis, and tissue homeostasis. Reflecting this central role, the molecular pathways that regulate cell division in eukaryotes are evolutionarily conserved. Aberrations in the control of the cell cycle are common in degenerative diseases and cancer. Therefore, analysis of the cell cycle in nonmammalian organisms can illuminate the processes underlying human development and disease. Forward genetic screens in yeast and Drosophila have been invaluable for gene discovery and have made important contributions to understanding pathways regulating cell proliferation. Importantly, it has been found that the human orthologs of some genes identified in these organisms are misexpressed in human tumors (Hariharan and Haber, 2003). Zebrafish have proven to be an excellent model of early vertebrate development (Driever et al., 1996; Haffter et al., 1996) and also of a wide variety of human diseases such as cancer, anemia, cardiovascular defects, neuromuscular conditions, kidney disease, and host–pathogen interaction, to name a few examples (Ackermann and Paw, 2003; Bassett and Currie, 2003; Drummond, 2005; Goessling et al., 2007; Hsu et al., 2007; Lambrechts and Carmeliet, 2004; Miller and Neely, 2004). The particular advantages that make zebrafish ideal for developmental embryology – including external fertilization of oocytes, transparent embryos, and rapid embryonic development – also provide the opportunity to study early cell divisions, tissue-specific cellular proliferation, and, more broadly, the role of cell-cycle genes in development and disease. A number of methods and markers have been successfully applied to investigate the cell cycle in zebrafish embryos, including video microscopy (Kane, 1999; Kane et al., 1992), histone-green fluorescent protein fusions (Pauls et al., 2001), 5-bromo-2-deoxyuridine (BrdU) labeling (Baye and Link, 2007; Link et al., 2000), proliferating cell nuclear antigen (PCNA) RNA and protein expression (Koudijs et al., 2005; Wullimann and Knipp, 2000), phosphohistone H3 (pH3) immunohistochemistry (Shepard et al., 2005), and minichromosome manintenance protein expression (Ryu and Driever, 2006). Studies of the developing zebrafish embryo have revealed similarities to the early cell divisions of other vertebrates such as Xenopus. In the zebrafish, the first seven cell divisions are synchronous and cycle rapidly between DNA replication (S phase) and mitosis (M phase) without the intervening gap phases, G1 and G2 (Kimmel et al., 1995). The midblastula transition (MBT) ensues during the 10th cell division, which is approximately 3 h postfertilization. MBT is accompanied by loss of division synchrony, increased cell-cycle duration, activation of zygotic transcription, and the onset of cellular motility (Kane and Kimmel, 1993). Embryonic cells first exhibit a G1 gap phase between the M and S phases during MBT. Recently, Dalle Nogare et al. demonstrated that during cycles 11–13, embryonic cells acquire a G2 phase in a transcription-independent fashion, through inhibition of Cdk1 and its activating phosphatase, Cdc25a (Dalle Nogare et al., 2008).

2. Analysis of Cell Proliferation, Senescence, and Cell Death in Zebrafish Embryos

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Further understanding of cell-cycle regulation in zebrafish embryos was obtained by studying their responses to various cell-cycle inhibitors, including aphidocolin, hydroxyurea, etoposide, camptothecin, and nocodazole (Ikegami et al., 1997a, 1997b, 1999). Exposure to these agents after MBT induces cell-cycle arrest, sometimes accompanied by initiation of an apoptotic program. However, prior to MBT, the embryonic cells continue to divide, often with deleterious effects, after exposure to cell-cycle inhibitors. These studies indicate that zebrafish embryos do possess cell-cycle checkpoints, but they are not functional until after MBT. Later developmental stages of zebrafish embryogenesis provide the opportunity to study the cell cycle in distinct tissue types. Studies of cell-cycle regulation in older embryos (10–36 h postfertilization) have focused on the developing eyes and central nervous system. Lineage analysis of CNS progenitor cells revealed a correlation between morphogenesis and cell-cycle number, implying that the nervous system development may be at least partially regulated by the cell cycle (Kimmel et al., 1994). Although most developing vertebrate embryos exhibit a constant lengthening of the cell-cycle duration throughout the development, meticulous analysis of cell number in the developing zebrafish retina revealed a surprising mechanism of modulated cell-cycle control. Li et al. (2000) reported that the retinal cell-cycle duration temporarily slows between 16 and 24 h postfertilization (hpf), followed by an abrupt change to more rapid cell divisions. Several studies have elucidated a role for the zebrafish cell-cycle machinery in tissue differentiation during development and in the regenerative response to injury. Bessa et al. (2008) found that Meis1, a marker of the eye primordium, promotes G1-S progression and a block of differentiation in the zebrafish eye through regulation of cyclin D1 and c-myc expression. Fischer and coworkers showed that loss of caf1b in zebrafish (by mutation or MO injection) leads to an S-phase arrest and eventual apoptosis that can be rescued by p53 deficiency. However, loss of caf1b also leads to a block in differentiation in tissues that express caf1b, implicating caf1b in the switch from proliferation to differentiation (Fischer et al., 2007). The effect of loss of early mitotic inhibitor 1 (emi1) on somite formation was evaluated by Zhang et al. These authors found that cell-cycle progression was required for proper somite morphogenesis, but not for the formation of the segmentation clock (Zhang et al., 2008). The role of the cell cycle in regeneration has also been assessed. Certain traumas result in loss of hair cell precursors, which results in deafness in vertebrates. Hernandez et al. (2007) used BrdU labeling and transgenic green fluorescent protein reporter lines to study hair cell regeneration, identifying proliferation-dependent and -independent mechanisms of hair cell renewal. A. Forward-Genetic Screens Several groups have carried out forward-genetic screens to identify mutations that alter cell proliferation in embryos. Shepard et al. used pH3 as a marker of cell proliferation in a two-generation haploid genetic screen. They identified seven mutant lines with different alterations in pH3 immunoreactivity. At least two of

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these lines demonstrate aneuploidy and increased cancer susceptibility as heterozygotes (Shepard et al., 2005, 2007). Using a similar screening strategy, Pfaff et al. identified a further set of genes required for cell proliferation mutants, among which was Scl-interrupting locus, which was identified as a novel, vertebrate-specific regulator of mitotic spindle assembly (Pfaff et al., 2007). Koudijs et al. (2005) used PCNA expression in the CNS as a readout to identify new mutations in repressors of the hedgehog signaling pathway. Finally, another screen for genes that control eye growth uncovered two zebrafish lines mutant for the anaphase-promoting complex/ cyclosome (APC/C) (Wehman et al., 2006). Loss of APC/C results in a loss of mitotic progression and apoptosis; in this study, co-labeling with BrdU and pH3 revealed cells undergoing mitotic catastrophe. In this chapter, we provide protocols to characterize the various phases of cell division in zebrafish embryos, and protocols to detect DNA damage, senescence, and cell death. Assays discussed in this chapter include DNA content analysis by flow cytometry, whole-mount embryonic antibody staining, mitotic spindle analysis, BrdU incorporation, cell death analysis, and in situ hybridization with cellcycle regulatory genes. Each assay targets different phases of the cell cycle and in total create a detailed picture of zebrafish embryo cell proliferation. Although our studies have focused on embryonic assays for cell-cycle characterization, it is likely that these protocols can be modified to study adult tissues. These protocols can be applied to a variety of experiments such as characterization of the cell-cycle phenotypes of mutants or the analysis of RNA overexpression and morpholino knockdown of cell-cycle regulatory genes. Furthermore, the genetic tractability of the zebrafish system (Patton and Zon, 2001) makes it an excellent organism in which to pursue forward genetic screens for mutations or chemical screens for novel compounds that alter cell division using one or more of these cell-cycle assays.

II. Zebrafish Embryo Cell-Cycle Protocols1 A. Analysis of Cell Proliferation and Mitosis

1. DNA Content Analysis A profile of the cell cycle in disaggregated zebrafish embryos or adult tissue can be obtained through DNA content analysis. In this technique, cells are stained with a dye that fluoresces upon DNA binding, such as Hoechst 33342 or propidium iodide. The intensity of fluorescence is proportional to the amount of DNA in each cell (Krishan, 1975). Analysis by fluorescence-activated cell sorting (FACS) generates a histogram showing the proportion of cells that have an unreplicated complement of DNA (G1 phase), those that have a fully replicated complement of DNA (G2 or M phase), and those that have an intermediate amount of DNA (S phase). 1

Items in boldface indicate reagents and supplies listed in Section V.


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Protocol: All steps are performed on ice except for the dechorionation (step 1) and RNAse incubation (step 9). 1. Dechorionate embryos and wash with E3. Analysis of single embryos is possible, though in practice, we typically pool approximately 40 embryos/tube. 2. Disaggregate embryos (using small pellet pestle) in 500 ml of DMEM (or other tissue culture medium) + 10% fetal calf serum in a matching homogenizing tube. 3. Bring volume to 1 mL with DMEM/serum and remove aggregates by passing cell suspension sequentially through 105 and 40 mm mesh. 4. Count a sample using a hemocytometer. 5. Place volume containing at least 2 106 cells in a 15 mL conical tube, and bring volume to 5 mL with 1 phosphate-buffered saline (PBS). 6. Spin at 1200 rpm for 10 min at 4 C. 7. Carefully aspirate off liquid and gently resuspend cell pellet in 2 mL propidium iodide solution. 8. Add 2 mg of DNAse-free RNAse (Roche). This step is necessary to remove double-stranded RNA, which binds propidium iodide. 9. Incubate in the dark at room temperature for 30 min. 10. Place samples on ice and analyze on FACS machine. Note: Samples can also be fixed in Ethanol, allowing multiple samples or time points to be collected for subsequent analysis. 1. Harvest cells and prepare single cell suspension in DMEM/serum as above, steps 1–4. 2. Wash cells in PBS and resuspend at 1–2 106 cells/mL. 3. To 1 mL cells in a 15 mL polypropylene, V-bottom tube, add 3 mL ice-cold absolute EtOH. To avoid clumping, add the ethanol dropwise while vortexing the sample. 4. Fix cells for at least 1 h at 4 C. Cells may be stored for several weeks at 20 C before undergoing PI staining. 5. Wash cells twice in 1 PBS. We typically increase the speed of centrifugation to 2500 rpm because the cells do not pellet as readily after EtOH fixation. 6. Resuspend the pellet in 1 mL propidium iodide solution. Add 2 mg of DNAsefree RNAse (Roche) and incubate for 3 h at 4 C. 7. Place samples on ice and analyze on FACS machine.

2. Whole Mount Immunohistochemistry with Mitotic Marker pH3 Histone H3 phosphorylation is considered to be a crucial event for the onset of mitosis and this antibody has been widely used in Drosophila and mammalian cell lines as a mitotic marker (Hendzel et al., 1997). Two members of the Aurora/AIK kinase family, Aurora A and Aurora B, phosphorylate histone H3 at the serine 10 residue (Chadee et al., 1999; Crosio et al., 2002). Increased serine 10 phosphorylation

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of histone H3 has been seen in transformed fibroblasts (Chadee et al., 1999), suggesting that this antibody could make an excellent marker for cell proliferation in the zebrafish as well as detecting cell-cycle mutations that may result in transformed phenotypes. In zebrafish, the pH3 antibody stains mitotic cells throughout the embryo (Fig. 1A). pH3 staining in developing organs like the nervous system increases as they undergo proliferation during distinct developmental stages. Protocol: 1. Fix embryos overnight at 4 C in 4% paraformaldehyde (PFA). 2. Permeabilize embryos for 7 min in 20 C acetone.

[(Fig._1)TD$IG]

Fig. 1

Useful techniques for the study of the cell cycle, proliferation, or apoptosis as shown in zebrafish embryos. (A) Antibody staining against phosphorylated histone H3 in wild-type 24 hpf embryos. (B) BrdU incorporation to mark cells in S phase in the tail of a 28 hpf wild-type embryo. (C and D) Apoptotic cells can be visualized by TUNEL (C, wild-type 24 hpf embryo) or acridine orange (D, 24 hpf crash&burn mutant embryo). (E) Anti-a tubulin can be used to examine mitotic spindle formation. (F) DNA content analysis shows the population of embryonic cells present in all phases of the cell cycle.


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3. Wash embryos in H2O followed by two 5-min washes in PBS plus Tween-20 (PBST). 4. Incubate for 30 min at room temperature in block. 5. Incubate overnight at 4 C in rabbit anti-pH3 at a concentration of 1.33 mg/mL in block. Two different sources of antibody have been used: Santa Cruz Biotechnology and an antiphosphopeptide polyclonal antibody to the sequence (ARKS[PO4]TGGKAPRKQLC) made and affinity purified by Genemed synthesis. 6. Wash 4 15 min in PBST. 7. Incubate for 2 h at room temperature in horseradish peroxidase-conjugated secondary goat anti-rabbit IgG (Jackson Immunoresearch) at a concentration of 3 mg/mL in block. 8. Wash 4 15 min in PBST. 9. Develop in the dark for 3–5 min at room temperature in diaminobenzidine (DAB) solution (0.67 mg/mL DAB in 15 mL of PBST to which 12 ml of 30% H2O2 has been added). 10. Wash in PBST and store embryos at 4 C in PFA.

3. Mitotic Spindle/Centrosome Detection Study of the mitotic spindle and centrosomes is an important step in understanding mutants with cell-cycle defects, particularly those whose phenotypes appear to be related to problems in mitosis. Genomic instability is one of the main alterations seen in human cancers and such unequal segregation of chromosomes can be caused by problems in mitotic spindle formation or centrosome number (Kramer et al., 2002). In this protocol, anti-a-tubulin labels the mitotic spindle, anti-b-tubulin the centrosome, and diamidino-2-phenylindole (DAPI) the DNA. Protocol: Fix embryos in PFA for 4 h at room temperature. Dehydrate in methanol at 20 C for at least 30 min. Rehydrate embryos in graded methanol:PBST series (3:1, 1:1, 1:3) for 5 min each. Wash 1 5 min in PBST. Place in 20 C acetone for 7 min. Wash 3 5 min in PBST. Incubate for 1 h at room temperature in block. Incubate in monoclonal mouse a-tubulin antibody (Sigma) at a concentration of 1:500 and in polyclonal rabbit g-tubulin antibody (Sigma) at a concentration of 1:1000 (both diluted in block) at 4 C overnight. 9. Wash 4 15 min in PBST. 10. Incubate in rhodamine-conjugated goat anti-mouse secondary (Molecular Probes) at 1:600 dilution and fluorescein-conjugated goat anti-rabbit secondary (Jackson Immunoresearch) at 1:600 dilution for 2 h room temperature. 1. 2. 3. 4. 5. 6. 7. 8.

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11. 12. 13. 14.

Wash 2 15 min in PBST. Include a 1:500 dilution of 100 mM DAPI during the third wash to stain DNA. Wash 2 15 min in PBST. For observation by epifluorescence microscopy, embryos are mounted on glass slides with VectaShield mounting media (Vector Labs) and coverslipped. To permit the specimen to lie flat, it is helpful to remove the yolk using forceps or a tungsten needle. Alternatively, for embryos >18 hpf, the tail can be cut off from the embryo and mounted on the slide.

4. BrdU Incorporation BrdU is a nucleoside analog that is specifically incorporated into DNA during S phase (Meyn et al., 1973) and can subsequently be detected with an anti-BrdU-specific antibody. This technique has been used to label replicating cells in zebrafish embryos (Larison and Bremiller, 1990) and adults (Rowlerson et al., 1997). The following protocol is designed to label a fraction of proliferating cells in zebrafish embryos, to allow comparison of the replication fraction of different embryos (Fig. 1 B). If the embryos are chased for varying amounts of time after the BrdU pulse, then fixed and stained for both BrdU and pH3 (section B), the transit of cells from S phase into G2/M can be assessed. This is useful in analyzing mutants with mitotic phenotypes. Protocol: 1. Dechorionate embryos and chill 15 min on ice in E3. 2. Prepare cold 10 mM BrdU/15% dimethylsulfoxide in E3 and chill on ice. Place embryos in BrdU solution and incubate for 20 min on ice to allow uptake of BrdU. 3. Change into warm E3 and incubate exactly for 5 min at 28.5 C. Note: longer incubation times will result in more cells being labeled. 4. Fix for 2 h at room temperature in PFA. Longer fixation may decrease the staining. 5. Transfer to methanol at 20 C overnight. All subsequent steps are performed at room temperature unless otherwise noted. 6. Rehydrate in graded methanol:PBST series (3:1, 1:1, 1:3) for 5 min each. 7. Wash 2 in PBST, 5 min. 8. Digest embryos in 10 mg/mL proteinase K, 10 min. 9. Wash in PBST. Refix in PFA for not more than 20 min. 10. Wash quickly 3 in H2O, then 2 in 2N HCl. 11. Incubate for 1 h in 2N HCl. This step denatures the labeled DNA to expose the BrdU epitope. 12. Remove the 2N HCl solution from the embryos and neutralize in 0.1 M borate buffer, pH 8.5, 20 min, room temparature. 13. Rinse several times in PBST. Block for 30 min in BrdU blocking solution. 14. Incubate in monoclonal anti-BrdU antibody at a dilution of 1:100 in BrdU block for 2 h at room temperature or overnight at 4 C. (If carrying out simultaneous BrdU/pH3 staining, add the primary anti-pH3 antibody as described in Section B, except that BrdU block is used.)


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15. Wash 5 10 min in PBST. 16. Incubate for 2 h room temperature with horseradish peroxidase or fluorophoreconjugated anti-mouse secondary antibody. (For simultaneous BrdU/pH3 stain, add a fluorescent anti-rabbit antibody as well.) 17. Wash 5 10 min in PBST. If using fluorescent secondary, mount embryos as described in Section C, step 14. 18. If using HRP-conjugated secondary antibody, develop in the dark for 3–5 min at room temperature in DAB solution (0.67 mg/mL DAB in 15 mL of PBST to which 12 mL of 30% H2O2 has been added). When staining is complete, wash 3 5 min in PBST, then fix in PFA.

B. Analysis of DNA Damage, Senescence, and Apoptosis

1. COMET Assay The Comet Assay, also known as the single cell microgel electrophoresis assay, is a highly sensitive technique that is used to detect DNA damage at the single-cell level (Singh et al., 1988). Cells are embedded into a thin agarose gel, through which a current is run allowing for migration of DNA. Smaller fragments of DNA, resulting from DNA damage, will travel more quickly and appear as a tail to the nucleus ‘‘comet head.’’ The comets can be visualized using a nuclear stain, such as SYBR green, and visualized under a fluorescent microscope. The following protocol is designed to isolate cells from zebrafish embryos and detect any kind of DNA damage. Variations of this technique allows for specific detection of double-stranded break. Protocol: 1. Dechorionate embryos and wash with E3. Typically, about 25–50 embryos are used. 2. Disaggregate embryos (using small pellet pestle) in 500 mL of DMEM (or other tissue culture medium) + 10% fetal calf serum or lamb serum in a matching homogenizing tube. 3. Bring volume to 1 mL with DMEM/serum. 4. Count the samples using a hemocytometer. 5. Spin down cells at 3000 RPM and resuspend in PBS to a concentration of 1 105 cells/mL. 6. Combine 10 mL of cells with 90 mL of molten LMAgarose (Trevigen) prewarmed to 37 C. Pipette 75 mL of the cell agarose mixture onto a CometSlide (Trevigen) prewarmed to 37 C. 7. Incubate the slide flat at 4 C for 30 min in the dark to allow the gel to solidify. 8. Immerse the slide in Lysis solution (Trevigen) containing 9% DMSO. After this point, it is very important to maintain the slide in low-light conditions. 9. Dry off the slide, and immerse it in alkaline solution for 30 min.

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[(Fig._2)TD$IG]

Fig. 2

The COMET assay reveals double-strand DNA breaks. Images of zebrafish embryo cells from a single-cell microgel electrophoresis experiment. (A) Unirradiated control cell. (B) A cell after exposure to 2000 R (20 Gy) g-irradiation. The arrow indicates the comet tail, composed of fragmented DNA (Verduzco and Amatruda, unpublished data).

10. Prepare a large horizontal electrophoresis apparatus by filling the chamber with fresh alkaline electrophoresis buffer and adjusting the volume of the alkaline electrophoresis buffer such that the current is 300 mA when the voltage is set to 25 V. Additionally, the chamber should be prepared and used in a 4 C room. 11. Place the slide in the electrophoresis apparatus. Run for 30 min at 4 C in the dark. 12. Dry off the slide. Rinse by dipping in ddH2O. 13. Incubate the slide in 70% EtOH for 5 min at room temperature in the dark. 14. Air-dry the slide for 1 h. 15. Pipette 50 mL of SYBR green staining solution (Trevigen) onto the microgel on the slide. 16. View the slide using fluorescence microscopy under a fluorescein filter (Fig. 2). 17. Comets can be analyzed using CometScore by Tritek Corp or another similar software program.

2. Detection of Senescence-Associated b-Galactosidase The study of cellular senescence was initiated by Hayflick and Moorhead (1961). Cellular senescence pertains to the cessation of cell replication and certain morphological and transcriptional changes that occur when cells permanently cease dividing. Although it is unclear whether the events that occur during in vitro cellular senescence also occur during organismal aging (Hayflick, 2007; Masoro, 2006), studies have revealed strong connections between cellular senescence, cancer, and age-related diseases (Campisi, 2005). Cellular senescence most likely arose evolutionarily as a mechanism to defend against tumorigenesis (Shay and Roninson, 2004). When a cell is afflicted by stress that may result in transformation (such as oxidative stress, DNA damage, or overepxpression of oncogenes), tumor-suppressor genes such as p53 may force the cell to undergo senescence-induced arrest. Arrested cells are functional but are not a risk for tumor initiation. Senescence also occurs as the ends of chromosomes, the telomeres, shorten. During reach replication cycle, if


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no active telomerase is present (Bodnar et al., 1998), the telomeres shorten, leading eventually to critically short telomeres that may interfere with gene expression and genomic stability (Shay and Wright, 2006). Normal cells senesce before telomeres shorten to the point of causing genomic instability, therefore instilling a counting mechanism that confirms Hayflick’s observation in 1961 (Shay and Wright, 2006). Senescent cells lose sensitivity to mitogens or growth factors, repress cell-cycle genes, such as cdk2, and become insensitive to apoptotic signals. Morphological changes occur resulting in an enlarged shape and flattened body (Ben-Porath and Weinberg, 2005), as well as expression of unique markers, and many of unknown function such as b-galactosidase activity at pH 6.0 (Dimri et al., 1995). Kishi and coworkers have used senescence-associated b-galactosidase staining in several studies to characterize senescence in normal and mutant zebrafish embryos and during aging of zebrafish adults (Kishi, 2004; Kishi et al., 2003, 2008; Tsai et al., 2007). Protocol: We have used the Senescence-Associated b-Galacotsidase Detection Kit from Sigma (CS 0030). The following protocol adapts the manufacturer’s instructions specifically for use with zebrafish embryos, and is kindly provided by Jenny Richardson and Dr. Elizabeth Patton, Edinburgh Cancer Research Center: 1. Dechorionate embryos and add 1.5 mL of 1 fixation buffer (prepared from 10 Sigma Senescence Fixation Buffer). Incubate overnight at 4 C. 2. Wash embryos 4 times in 1 PBS, 1 h each wash. 3. Make up the senescence staining mixture as per the manufacturer’s protocol. Add 1 mL to embryos and incubate for 24 h at 37 C. 4. Wash embryos 3 times in 1 PBS, 10 min each wash. 5. Embryos can be stored at 4 C in 1 PBS and 0.1% NaN3 or in 70% glycerol at 4 C. An alternative protocol was described by Kishi et al. (2008) in a recent paper describing a senescence-based genetic screen. The following protocol is adapted from Kishi et al., ‘‘The identification of zebrafish mutants showing alterations in senescence-associated biomarkers,’’ PLoS Genet. 2008 Aug 15;4(8):e1000152: 1. Fix embryos or adult zebrafish in 4% PFA in 1 PBS at 4 C (for 3 days in adults and overnight in embryos). 2. Wash three times for 1 h in PBS with pH 7.4 and for a further 1 h in PBS with pH 6.0 at 4 C. 3. Stain the samples overnight at 37 C in 5 mM potassium ferrocyanide, 5 mM potassium ferricyanide, 2 mM MgCl2, and 1 mg/ml X-gal in PBS adjusted to pH 6.0.

3. Apoptosis Detection by TUNEL Staining Apoptosis is a form of programmed cell death that eliminates damaged or unneeded cells. It is controlled by multiple signaling pathways that mediate

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responses to growth, survival, or death signals. Cell-cycle checkpoint controls are linked to apoptotic cascades and these connections can be compromised in diseases including cancer. The defining characteristics of apoptosis are membrane blebbing, cell shrinkage, nuclear condensation, segmentation, and division into apoptotic bodies that are phagocytosed (Wyllie, 1987). The DNA strand breaks that occur during apoptosis can be detected by enzymatically labeling the free ends with modified nucleotides, which can then be detected with antibodies (Gavrieli et al., 1992). Protocol: 1. Embryos are fixed overnight at 4 C in PFA. 2. Wash in PBS and transfer to methanol for 30 min at 20 C. 3. Rehydrate embryos in a graded methanol:PBST series (3:1, 1:1, 1:3) for 5 min each. 4. Wash 1 5 min in PBST. 5. Digest embryos in proteinase K (10 mg/mL) at room temperature (1 min for embryos younger than 16 hpf, 2 min for embryos older than 16 hpf). 6. Wash twice in PBST. 7. Postfix in PFA for 20 min at room temperature. 8. Wash 5 5 min in PBST. 9. Postfix for 10 min at 20 C with prechilled ethanol:acetic acid 2:1. 10. Wash 3 5 min in PBST at room temperature. 11. Incubate for 1 h at room temperature in 75 mL equilibration buffer (TdT – ApopTag Peroxidase In Situ Apoptosis Detection Kit from Serologics Corporation). 12. Add small volume of working strength TdT (reaction buffer and TdT at a ratio of 2:1 plus 0.3% Triton) (Serologics Corporation). 13. Incubate overnight at 37 C. 14. Stop reaction by washing in working strength stop/wash buffer (1 mL concentrated buffer from Serologics Kit with 34 mL water) for 3–4 h at 37 C. 15. Wash 3 5 min in PBST. 16. Block with 2 mg/mL BSA, 5% sheep serum in PBST for 1 h at room temperature. 17. Incubate in anti-digoxigenin peroxidase antibody included in kit (full strength). 18. Wash 4 30 min PBST at room temperature. 19. Develop in the dark for 5 min at room temperature in DAB solution (0.67 mg/ mL in 15 mL of PBST) and 12 ml 30% H2O2. 20. Wash in PBST and store embryos at 4 C in PFA.

4. Apoptosis Detection by Acridine Orange Another method of apoptotic cell detection that can be performed on living embryos is acridine orange staining. The basis of this method is that the ATPdependent lysosomal proton pump is preserved in apoptotic but not necrotic cells;


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therefore, apoptotic cells will take up the acridine orange dye whereas living or necrotic cells will not (Darzynkiewicz et al., 1992). This method is useful for identifying mutants based on an apoptotic phenotype in order to further characterize them in living assays. Protocol: 1. Live dechorionated embryos are incubated in a 2 mg/mL solution of acridine orange (Sigma) in 1 E3 for 30 min at room temperature. 2. Embryos are washed 5 quickly in E3 and then visualized on a stereo dissecting microscope equipped for FITC epifluorescence.

C. In Situ Hybridization RNA expression analysis by in situ hybridization of antisense probes in wholemount zebrafish embryos is a commonly used technique to localize expression of developmental regulatory genes. While the technique is not exceptionally quantitative, it can reveal stark differences in gene expression. More quantitative analysis of gene expression, such as Northern blotting, and real-time polymerase chain reaction do not permit the examination of alterations in tissue-specific expression or an expression pattern. Cell division is a highly controlled process that involves regulation at both the transcriptional and posttranslational stages. Cyclins are a class of proteins that play critical roles in guiding cells through the G1, S, G2, and M phases of the cell cycle by regulating the activity of the cyclin-dependent kinases. The name cyclin alludes to the fact that their expression levels oscillate between peaks and nadirs that are coordinated with particular phases of the cell cycle (reviewed in Murray, 2004). The tightly regulated expression of these important cell-cycle genes incorporates transcriptional, translational, and posttranslational controls. Many genes involved in cell-cycle regulation are specifically expressed during the cell-cycle phase in which they act. Zebrafish orthologs of cell-cycle regulatory genes such as PCNA and cyclins have been found to possess similar expression patterns throughout the proliferative tissues of developing zebrafish embryos (C. Thisse, B. Thisse, unpublished data and www. zfin.org). In situ hybridization for cell-cycle regulatory genes can be performed using previously published in situ hybridization protocols (Jowett, 1999; Thisse et al., 1993, 1994).

III. Screening for Chemical Suppressors of Zebrafish Cell-Cycle Mutants Another way to probe the cell cycle is via chemical agents. Chemical screens could identify novel compounds that are useful tools for studying the cell cycle. Furthermore, mutations in cell-cycle genes are commonly found in human cancer.

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Given the need to improve upon current cancer therapy, one approach is to identify small molecule suppressors that bypass the consequences of specific cell-cycle gene mutations. Akin to the use of genetic modifier screens to identify secondary mutations that enhance or suppress a primary defect (St Johnston, 2002), chemical suppressor screens would directly identify small molecules that rescue a genetic phenotype. If the phenotype is disease-related, such compounds might represent lead therapeutic agents. Zebrafish have recently been utilized in chemical screens to identify compounds that perturb specific aspects of development (Anderson et al., 2007; Bayliss et al., 2006; den Hertog, 2005; Khersonsky et al., 2003; Peterson et al., 2000, 2004). The zebrafish system offers several advantages for chemical screens, providing information on tissue specificity and toxicity, and accounting for compound activation via drug metabolism. Furthermore, cells are not transformed and are in their normal physiological milieu of cell–cell and cell–extracellular matrix interactions. Murphey et al. (2006) carried out a high-throughput chemical screen to detect small molecules capable of perturbing the cell cycle during zebrafish development, identifying several compounds that were not previously detected in cell-based screens of the same library. As another application of this technique, Stern et al. screened a 16,000-compound library to identify small molecules capable of suppressing the cell proliferation defect in the crash&burn cell-cycle mutant (Stern et al., 2005). This technology could easily be applied to other cell-cycle mutants and modified to use cell-cycle assays other than pH3 staining. In addition, such chemical suppressor screens could be applied to any zebrafish model of human disease (Dooley and Zon, 2000). For these reasons, we provide a detailed protocol below. The following protocol can be repeated weekly giving a throughput of more than 1000 compounds per week for a recessive lethal mutation. In the case of homozygous viable mutants, the throughput could be improved by using fewer embryos (3–5) per well in 96-well plates. Protocol: 1. For a chemical screen, large numbers of embryos at approximately the same developmental stage need to be generated. Set up 100 heterozygote pairwise matings with fish separated by a divider. The next morning, remove the divider, allow the fish to mate, and collect the embryos. 2. Dilute chemicals into screening medium. The screen is conducted in 48-well plates with a volume of 300 ml per well. Individual chemicals could be added to each well, but to improve throughput, we devised a matrix pooling strategy: The chemical library (courtesy of the Institute of Chemistry and Cell Biology, Harvard Medical School) was arrayed in 384-well plates with the last four columns empty, thus containing 320 compounds per plate. Given this plate geometry, 8 10 matrix pools were created. A hit detected in both a horizontal and a vertical pool identified the individual compound. a. Transfer 80 ml of screening medium to each well of four 384-well plates using a TECAN liquid-handling robot.


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b. Pin transfer 1 ml of each compound (arrayed at 5 mg/mL in DMSO) into each well of screening medium by performing 10 transfers with a 100 nl 384-pin array for each of the four 384-well plates (total of 320 4 = 1280 compounds). c. Pooling was performed with a TECAN liquid-handling robot by pipeting the diluted chemicals from the 384-well plates to 48-well plates. For vertical pools, 30 mL was transferred from each of 8 wells plus an additional 60 mL of screening medium to bring the total volume to 300 ml. For horizontal pools, 30 mL was transferred from each of 10 wells. 3. Aliquot embryos to the 48-well plates at 50% epiboly. a. Prior to aliquoting embryos to wells, examine them under a dissecting microscope and discard all dead, delayed, or deformed embryos. b. Pool embryos in a single 100 mm tissue culture dish or a 50 mL conical tube. c. Decant the embryo medium and remove as much liquid from the embryo suspension as possible with a transfer pipet. Pressing the transfer pipet tip to the bottom of the tube or dish allows most liquid to be removed without aspirating the embryos. d. Add approximately 20 embryos to each well by scooping them with a small chemical weighing spatula. With 20 embryos per well and a Mendelian recessive inheritance, there is a 0.3% chance of a well having no mutants. Since a hit requires detection in both a horizontal and a vertical pool, each with 20 embryos, the false-positive rate for identification of complete suppressors is 0.001%. 4. Place 48-well plates into an incubator at 28.5 C. 5. One to two hours later, clean out any dead embryos from each well using a long glass Pasteur pipet bent at a 90 degree angle. 6. Incubate at 28.5 C overnight. 7. Dechorionate embryos by adding 150 mL of a 5 mg/mL pronase solution to each well. After 10 min, gently shake plates until embryos come out of the chorions. 8. Using a transfer pipet fitted with a 10 mL tip, remove as much of the pronase/ chemical mixture as possible from each well. 9. Rinse the embryos once in fresh embryo medium and remove as in step 8. 10. Add 500 ml of PFA to each well. 11. Parafilm the edges of the plates to prevent evaporation and fix at 4 C at least overnight but not longer than a week. 12. Using a transfer pipet, move embryos to 48-well staining grids made of acetoneresistant plastic with a wire mesh bottom. 13. Perform pH3 staining protocol by placing staining grids into 11 8.5 cm reservoirs containing 20–30 mL of the appropriate solution. To change solutions, the grid can be lifted out of one reservoir and placed into another reservoir with the next solution. For overnight antibody incubations, the reservoir should be sealed with parafilm to prevent evaporation.

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14. After staining is complete, move embryos with a transfer pipet back into 48-well plates that have been precoated with 100 mL of 1% agarose in 1 PBS. The agarose forms a meniscus that keeps embryos in the center of the well where they are easier to score. 15. Score for absence of mutants or for partial suppression without effect on wild types. In addition to suppressors and enhancers, one can identify compounds that affect both wild types and mutants, thus having a more general effect. 16. Deconvolute matrix pool to identify individual chemicals.

IV. Conclusions Given the power of zebrafish in forward vertebrate genetics and organism-based small molecule screens, the system will nicely complement traditional model organisms for studying the cell division cycle. Many of the assays that are commonly used to probe the cell cycle in systems such as yeast, Drosophila, and mammalian cells can be used in the zebrafish. The protocols outlined in this chapter can be utilized to characterize known mutants for alterations in cell proliferation or, alternatively, can be used to screen for more cell-cycle mutants. Given that zebrafish embryos are amenable to gene knockdown via antisense morpholino-modified oligonucleotides and overexpression by mRNA injection, these protocols can also be used to study cell-cycle genes in the zebrafish without generating a mutant.

V. Reagents and Supplies Alkaline solution Buffer Anti-BrdU Block BrdU block

DAPI DMSO E3 Mesh PBS PBST Pellet pestle & tubes PFA Propidium iodide

0.6 g NaOH, 250 mL 200 mM EDTA pH 10, to 50 mL ddH2O Alkaline electrophoresis 12 g NaOH, 2 mL 500 mM EDTA pH 8, to 1 L ddH2O Roche cat # 1170 376 2% Blocking reagent (Roche cat # 1096 176), 10% fetal calf serum, 1% dimethylsulfoxide in PBST 0.2% Blocking reagent (Roche cat # 1096 176), 10% fetal calf serum, 1% dimethylsulfoxide in PBST. The lower concentration of blocking reagent improves detection 40 ,6-Diamidino-2-phenylindole Dimethylsulfoxide 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4 Small Parts, Inc. 105 mm mesh with cat # U-CMN-105D. 40 mm mesh with cat # U-CMN-40D Phosphate-buffered saline, pH 7.5. 1 PBS with 0.1% (v/v) Tween-20 Fisher, cat # K749520-0090 4% PFA buffered with 1 PBS 0.1% Sodium citrate, 0.05 mg/mL propidium iodide, 0.0002% Triton X-100 (added fresh)

2. Analysis of Cell Proliferation, Senescence, and Cell Death in Zebrafish Embryos Sigma senescence fixation buffer 10 (cat # F1797) Sigma senescence staining mixture (prepare just prior to use)

Screening medium

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Contains 20% formaldehyde, 2% glutaraldehyde, 70.4 mM Na2HPO4, 14.7 mM KH2PO4, 1.37 M NaCl, and 26.8 mM KCl Mix the following for preparation of 10 ml of the staining mixture: 1 mL of staining solution 10 buffer (cat # S5818) 125 mL of reagent B (cat # R5272) 125 mL of reagent C (cat # R5147) 0.25 mL of X-gal solution (cat # X3753) 8.50 mL of ultrapure water E3 supplemented with 1% DMSO, 20 mM metronidazole, 50 units/mL penicillin, 50 mg/mL streptomycin, and 1 mM Tris pH 7.4

Acknowledgments We thank Len Zon and members of the Zon laboratory for useful discussions. Original versions of many of these protocols were worked out by Jennifer L. Shepard, Ryan Murphey, Howard M. Stern, Kathryn L. Pfaff, and J.F.A. D.V. was supported by NIH Training Grant 5 T32 GM008203 and J.F.A. was supported by grants from the Lance Armstrong Foundation, the Amon G. Carter Foundation, the Welch Foundation, and NIH/NCI grant 1R01CA135731.

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Daniel Verduzco and James F. Amatruda Dalle Nogare, D. E., Pauerstein, P. T., and Lane, M. E. (2008). G2 acquisition by transcription-independent mechanism at the zebrafish midblastula transition. Dev. Biol. . Darzynkiewicz, Z., Bruno, S., Del Bino, G., Gorczyca, W., Hotz, M. A., Lassota, P., Traganos, F. (1992). Features of apoptotic cells measured by flow cytometry. Cytometry 13(8), 795–808. den Hertog, J. (2005). Chemical genetics: drug screens in Zebrafish. Biosci. Rep. 25(5–6), 289–297. Dimri, G. P., Lee, X., Basile, G., Acosta, M., Scott, G., Roskelley, C., Medrano, E. E., Linskens, M., Rubelj, I., Pereira-Smith, O., et al. (1995). A biomarker that identifies senescent human cells in culture and in aging skin in vivo. Proc. Natl. Acad. Sci. U.S.A. 92(20), 9363–9367. Dooley, K., and Zon, L. I. (2000). Zebrafish: a model system for the study of human disease. Curr. Opin. Genet. Dev. 10(3), 252–256. Driever, W., Solnica-Krezel, L., Schier, A. F., Neuhauss, S. C., Malicki, J., Stemple, D. L., Stainier, D. Y., Zwartkruis, F., Abdelilah, S., Rangini, Z., Belak, J., Boggs, C. (1996). A genetic screen for mutations affecting embryogenesis in zebrafish. Development 123, 37–46. Drummond, I. A. (2005). Kidney development and disease in the zebrafish. J. Am. Soc. Nephrol. 16(2), 299–304. Fischer, S., Prykhozhij, S., Rau, M. J., and Neumann, C. J. (2007). Mutation of zebrafish caf-1b results in S phase arrest, defective differentiation, and p53-mediated apoptosis during organogenesis. Cell Cycle 6 (23), 2962–2969. Gavrieli, Y., Sherman, Y., and Ben-Sasson, S. A. (1992). Identification of programmed cell death in situ via specific labeling of nuclear DNA fragmentation. J. Cell Biol. 119(3), 493–501. Goessling, W., North, T. E., and Zon, L. I. (2007). New waves of discovery: modeling cancer in zebrafish. J. Clin. Oncol. 25(17), 2473–2479. Haffter, P., Granato, M., Brand, M., Mullins, M. C., Hammerschmidt, M., Kane, D. A., Odenthal, J., van Eeden, F. J., Jiang, Y. J., Heisenberg, C. P., Kelsh, R. N., Furutani-Seiki, M., et al. (1996). The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio. Development 123, 1–36. Hariharan, I. K., and Haber, D. A. (2003). Yeast, flies, worms, and fish in the study of human disease. N. Engl. J. Med. 348(24), 2457–2463. Hayflick, L. (2007). Biological aging is no longer an unsolved problem. Ann. N.Y. Acad. Sci. 1100, 1–13. Hayflick, L., and Moorhead, P. S. (1961). The serial cultivation of human diploid cell strains. Exp. Cell. Res. 25, 585–621. Hendzel, M. J., Wei, Y., Mancini, M. A., Van Hooser, A., Ranalli, T., Brinkley, B. R., Bazett-Jones, D. P., Allis, C. D. (1997). Mitosis-specific phosphorylation of histone H3 initiates primarily within pericentromeric heterochromatin during G2 and spreads in an ordered fashion coincident with mitotic chromosome condensation. Chromosoma 106(6), 348–360. Hernandez, P. P., Olivari, F. A., Sarrazin, A. F., Sandoval, P. C., and Allende, M. L. (2007). Regeneration in zebrafish lateral line neuromasts: expression of the neural progenitor cell marker sox2 and proliferation-dependent and-independent mechanisms of hair cell renewal. Dev. Neurobiol. 67(5), 637–654. Hsu, C. H., Wen, Z. H., Lin, C. S., and Chakraborty, C. (2007). The zebrafish model: use in studying cellular mechanisms for a spectrum of clinical disease entities. Curr. Neurovasc. Res. 4(2), 111–120. Ikegami, R., Hunter, P., and Yager, T. D. (1999). Developmental activation of the capability to undergo checkpoint-induced apoptosis in the early zebrafish embryo. Dev. Biol. 209(2), 409–433. Ikegami, R., Rivera-Bennetts, A. K., Brooker, D. L., and Yager, T. D. (1997a). Effect of inhibitors of DNA replication on early zebrafish embryos: evidence for coordinate activation of multiple intrinsic cellcycle checkpoints at the mid-blastula transition. Zygote 5(2), 153–175. Ikegami, R., Zhang, J., Rivera-Bennetts, A. K., and Yager, T. D. (1997b). Activation of the metaphase checkpoint and an apoptosis programme in the early zebrafish embryo, by treatment with the spindledestabilising agent nocodazole. Zygote 5(4), 329–350. Jowett, T. (1999). Analysis of protein and gene expression. Methods Cell Biol. 59, 63–85. Kane, D. A. (1999). Cell cycles and development in the embryonic zebrafish. Methods Cell Biol. 59, 11–26.


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Kane, D. A., and Kimmel, C. B. (1993). The zebrafish midblastula transition. Development 119(2), 447–456. Kane, D. A., Warga, R. M., and Kimmel, C. B. (1992). Mitotic domains in the early embryo of the zebrafish. Nature 360(6406), 735–737. Khersonsky, S. M., Jung, D. W., Kang, T. W., Walsh, D. P., Moon, H. S., Jo, H., Jacobson, E. M., Shetty, V., Neubert, T. A., Chang, Y. T. (2003). Facilitated forward chemical genetics using a tagged triazine library and zebrafish embryo screening. J. Am. Chem. Soc. 125(39), 11804–77805. Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., and Schilling, T. F. (1995). Stages of embryonic development of the zebrafish. Dev. Dyn. 203(3), 253–310. Kimmel, C. B., Warga, R. M., and Kane, D. A. (1994). Cell cycles and clonal strings during formation of the zebrafish central nervous system. Development 120(2), 265–276. Kishi, S. (2004). Functional aging and gradual senescence in zebrafish. Ann. N. Y. Acad. Sci. 1019, 521–526. Kishi, S., Bayliss, P. E., Uchiyama, J., Koshimizu, E., Qi, J., Nanjappa, P., Imamura, S., Islam, A., Neuberg, D., Amsterdam, A., Roberts, T. M. (2008). The identification of zebrafish mutants showing alterations in senescence-associated biomarkers. PLoS Genet 4(8), e1000152. Kishi, S., Uchiyama, J., Baughman, A. M., Goto, T., Lin, M. C., Tsai, S. B. (2003). The zebrafish as a vertebrate model of functional aging and very gradual senescence. Exp. Gerontol 38(7), 777–786. Koudijs, M. J., den Broeder, M. J., Keijser, A., Wienholds, E., Houwing, S., van Rooijen, E. M., Geisler, R., van Eeden, F. J. (2005). The zebrafish mutants dre, uki, and lep encode negative regulators of the hedgehog signaling pathway. PLoS Genet. 1(2), e19. Kramer, A., Neben, K., and Ho, A. D. (2002). Centrosome replication, genomic instability and cancer. Leukemia 16(5), 767–775. Krishan, A. (1975). Rapid flow cytofluorometric analysis of mammalian cell cycle by propidium iodide staining. J. Cell. Biol. 66(1), 188–193. Lambrechts, D., and Carmeliet, P. (2004). Genetics in zebrafish, mice, and humans to dissect congenital heart disease: insights in the role of VEGF. Curr. Top. Dev. Biol. 62, 189–224. Larison, K. D., and Bremiller, R. (1990). Early onset of phenotype and cell patterning in the embryonic zebrafish retina. Development 109(3), 567–576. Li, Z., Hu, M., Ochocinska, M. J., Joseph, N. M., and Easter Jr., S. S. (2000). Modulation of cell proliferation in the embryonic retina of zebrafish (Danio rerio). Dev. Dyn. 219(3), 391–401. Link, B. A., Fadool, J. M., Malicki, J., and Dowling, J. E. (2000). The zebrafish young mutation acts noncell-autonomously to uncouple differentiation from specification for all retinal cells. Development 127 (10), 2177–2188. Masoro, E. J. (2006). Dietary restriction-induced life extension: a broadly based biological phenomenon. Biogerontology 7(3), 153–155. Meyn, R. E., Hewitt, R. R., and Humphrey, R. M. (1973). Evaluation of S phase synchronization by analysis of DNA replication in 5-bromodeoxyuridine. Exp. Cell. Res. 82(1), 137–142. Miller, J. D., and Neely, M. N. (2004). Zebrafish as a model host for streptococcal pathogenesis. Acta Trop. 91(1), 53–68. Murphey, R. D., Stern, H. M., Straub, C. T., and Zon, L. I. (2006). A chemical genetic screen for cell cycle inhibitors in zebrafish embryos. Chem. Biol. Drug Des. 68(4), 213–219. Murray, A. W. (2004). Recycling the cell cycle: cyclins revisited. Cell 116(2), 221–234. Patton, E. E., and Zon, L. I. (2001). The art and design of genetic screens: zebrafish. Nat. Rev. Genet 2(12), 956–966. Pauls, S., Geldmacher-Voss, B., and Campos-Ortega, J. A. (2001). A zebrafish histone variant H2A.F/Z and a transgenic H2A.F/Z:GFP fusion protein for in vivo studies of embryonic development. Dev. Genes Evol. 211(12), 603–610. Peterson, R. T., Link, B. A., Dowling, J. E., and Schreiber, S. L. (2000). Small molecule developmental screens reveal the logic and timing of vertebrate development. Proc. Natl. Acad. Sci. U.S.A. 97(24), 12965–12969.

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Daniel Verduzco and James F. Amatruda Peterson, R. T., Shaw, S. Y., Peterson, T. A., Milan, D. J., Zhong, T. P., Schreiber, S. L., MacRae, C. A., Fishman, M. C. (2004). Chemical suppression of a genetic mutation in a zebrafish model of aortic coarctation. Nat. Biotechnol. 22(5), 595–599. Pfaff, K. L., Straub, C. T., Chiang, K., Bear, D. M., Zhou, Y., Zon, L. I. (2007). The zebra fish cassiopeia mutant reveals that SIL is required for mitotic spindle organization. Mol. Cell Biol. 27(16), 5887–5897. Rowlerson, A., Radaelli, G., Mascarello, F., and Veggetti, A. (1997). Regeneration of skeletal muscle in two teleost fish: Sparus aurata and Brachydanio rerio. Cell Tissue Res. 289(2), 311–322. Ryu, S., and Driever, W. (2006). Minichromosome maintenance proteins as markers for proliferation zones during embryogenesis. Cell Cycle 5(11), 1140–1142. Shay, J. W., and Roninson, I. B. (2004). Hallmarks of senescence in carcinogenesis and cancer therapy. Oncogene 23(16), 2919–2933. Shay, J. W., and Wright, W. E. (2006). Telomerase therapeutics for cancer: challenges and new directions. Nat. Rev. Drug Discov. 5(7), 577–584. Shepard, J. L., Amatruda, J. F., Finkelstein, D., Ziai, J., Finley, K. R., Stern, H. M., Chiang, K., Hersey, C., Barut, B., Freeman, J. L., Lee, C., Glickman, J. N., et al. (2007). A mutation in separase causes genome instability and increased susceptibility to epithelial cancer. Genes Dev. 21(1), 55–59. Shepard, J. L., Amatruda, J. F., Stern, H. M., Subramanian, A., Finkelstein, D., Ziai, J., Finley, K. R., Pfaff, K. L., Hersey, C., Zhou, Y., Barut, B., Freedman, M., et al. (2005). A zebrafish bmyb mutation causes genome instability and increased cancer susceptibility. Proc. Natl. Acad. Sci. U.S.A. 102(37), 13194–13199. Singh, N. P., McCoy, M. T., Tice, R. R., and Schneider, E. L. (1988). A simple technique for quantitation of low levels of DNA damage in individual cells. Exp. Cell Res. 175(1), 184–191. St Johnston, D. (2002). The art and design of genetic screens: Drosophila melanogaster. Nat. Rev. Genet 3 (3), 176–188. Stern, H. M., Murphey, R. D., Shepard, J. L., Amatruda, J. F., Straub, C. T., Pfaff, K. L., Weber, G., Tallarico, J. A., King, R. W., Zon, L. I. (2005). Small molecules that delay S phase suppress a zebrafish bmyb mutant. Nat. Chem. Biol. 1(7), 366–370. Thisse, C., Thisse, B., Halpern, M. E., and Postlethwait, J. H. (1994). Goosecoid expression in neurectoderm and mesendoderm is disrupted in zebrafish cyclops gastrulas. Dev. Biol. 164(2), 420–449. Thisse, C., Thisse, B., Schilling, T. F., and Postlethwait, J. H. (1993). Structure of the zebrafish snail1 gene and its expression in wild-type, spadetail and no tail mutant embryos. Development 119(4), 1203–1215. Tsai, S. B., Tucci, V., Uchiyama, J., Fabian, N. J., Lin, M. C., Bayliss, P. E., Neuberg, D. S., Zhdanova, I. V., Kishi, S. (2007). Differential effects of genotoxic stress on both concurrent body growth and gradual senescence in the adult zebrafish. Aging Cell 6(2), 209–224. Wehman, A. M., Staub, W., and Baier, H. (2006). The anaphase-promoting complex is required in both dividing and quiescent cells during zebrafish development. Dev. Biol. . Wullimann, M. F., and Knipp, S. (2000). Proliferation pattern changes in the zebrafish brain from embryonic through early postembryonic stages. Anat. Embryol. (Berl.) 202(5), 385–400. Wyllie, A. H. (1987). Apoptosis: cell death in tissue regulation. J. Pathol. 153(4), 313–316. Zhang, L., Kendrick, C., Julich, D., and Holley, S. A. (2008). Cell cycle progression is required for zebrafish somite morphogenesis but not segmentation clock function. Development 135(12), 2065–2070.

CHAPTER 3

Analysis of Cilia Structure and Function in Zebrafish Jarema Malicki,* Andrei Avanesov,* Jade Li,y Shiaulou Yuany and Zhaoxia Suny *

Division of Craniofacial and Molecular Genetics, Tufts University, Massachusetts, USA

y

Departments of Genetics, Yale University School of Medicine, New Haven, Connecticut, USA

Abstract I. Introduction II. Cilia in Zebrafish Embryos and Larvae A. Kupffer’s Vesicle B. Pronephros C. Sensory Organs D. Neural Tube III. Analytical Tools for Cilia Morphology and Motility A. Method 1: Imaging of Kupffer’s Vesicle Cilia Following arl13b/sco-eGFP mRNA Overexpression B. Method 2: Modified Methods for Imaging of the Pronephric Duct and Spinal Central Canal Using arl13b/sco-eGFP Over-Expressants C. Alternative Method: Imaging of Cilia Using arl13b/scorpion-eGFP Transgenic Fish D. Method 3. Detection of Protein Expression in Cilia Using Immunohistochemistry IV. Analysis of Cilia-related Mutant Phenotypes in Zebrafish A. Left–Right Asymmetry Defects B. Method 4: Evaluation of Heart Positioning in Live Zebrafish Embryos C. Alternative Method: Analysis of Left–Right Asymmetry Defects Using in situ Hybridization D. Kidney Cysts E. Method 5: Detection of Kidney Cysts in Zebrafish F. Degeneration of Sensory Neurons G. Method 6: Evaluation of Photoreceptor Cell Layer Morphology on Plastic Sections H. Method 7: Analysis of Photoreceptor Cells on Cryosections via Immunohistochemistry I. Method 8: Whole-Mount Staining and Imaging of Hair Cells in the Inner Ear J. Method 9: Detection of Basal Bodies in Hair Cells K. Method 10: Detection of Neuromast Hair Cells in a Living Specimen METHODS IN CELL BIOLOGY, VOL 101 Copyright 2011. Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-387036-0.00003-7

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Jarema Malicki et al.

L. Method 11: Labeling of Olfactory Neurons by DiI Incorporation V. Future Directions Acknowledgments References

Abstract The cilium, a previously little studied cell surface protrusion, has emerged as an important organelle in vertebrate cells. This tiny structure is essential for normal embryonic development, including the formation of left-right asymmetry, limb morphogenesis, and the differentiation of sensory cells. In the adult, cilia also function in a variety of processes, such as the survival of photoreceptor cells, and the homeostasis in several tissues, including the epithelia of nephric ducts. Human ciliary malfunction is associated with situs inversus, kidney cysts, polydactyly, blindness, mental retardation, obesity, and many other abnormalities. The genetic accessibility and optical transparency of the zebrafish make it an excellent vertebrate model system to study cilia biology. In this chapter, we describe the morphology and distribution of cilia in zebrafish embryonic and larval organs. We also provide essential protocols to analyze cilia formation and function.

I. Introduction Cilia are thin cell surface protrusions of varying length supported by a characteristic arrangement of microtubules. On cross sections, the vast majority of cilia feature nine evenly spaced peripheral microtubule doublets. Two additional microtubule singlets are frequently present in the center of the cilium. The microtublebased core cytoskeleton of the cilium is known as the axoneme. The arrangement of axonemal microtubules is exceptionally well conserved in evolution, remaining essentially unchanged in mammals, fish, and green algae (Avidor-Reiss et al., 2004; Rosenbaum and Witman, 2002; Satir et al., 2008). Cilia are associated with a wide spectrum of processes in embryonic development and in adult physiology. In the embryo, cilia are necessary for the determination of left–right (LR) asymmetry, limb morphogenesis, the formation of ventricular spaces in the brain, the differentiation of the craniofacial skeleton, and finally the differentiation of sensory neurons, such as photoreceptor cells of the retina (Basu and Brueckner, 2008; Haycraft et al., 2005; Tissir et al., 2010; Tsujikawa and Malicki, 2004; Walczak-Sztulpa et al., 2010). In adult mammals, cilia are required for the function of photoreceptor and olfactory sensory neurons, the removal of foreign particles from the respiratory track, the transport of oocytes in the oviduct, the motility of the sperm, the maintenance of kidney duct morphology, and the deposition of fat (Afzelius, 2004; Cardenas-Rodriguez and Badano, 2009). On the cellular level, cilia appear to play two major roles. First, they function as mechanical drivers of movement. Cilia can propagate the movement of cells or small

3. Analysis of Cilia Structure and Function in Zebrafish

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multicellular organisms. Similarly, apical cilia drive fluid flow over the surface of epithelial sheets. Second, cilia provide a subcellular compartment for signal transduction cascades that range from phototransduction to hedgehog signaling. This second mode of action underlies the role of cilia in signal detection and signal transduction (reviewed in, Green and Mykytyn, 2010). Given the diversity of ciliary functions, it is to be expected that they have received considerable attention in organisms that range from unicellular algae to humans. The zebrafish is a vertebrate animal model exceptionally well suited for the studies of cilia formation and function for several reasons. First, the zebrafish embryo develops outside the maternal organism, is transparent, and features several organs that differentiate prominent cilia early in embryogenesis. These characteristics make it easy to monitor the length and movement of the ciliary axoneme (Sun et al., 2004; Tsujikawa and Malicki, 2004). Second, zebrafish cilia mutants frequently feature the curved body axis, a phenotype, that is very easy to detect in genetic screens. Consequently an impressive collection of ciliary mutants is available in zebrafish (Brand et al., 1996; Doerre and Malicki, 2002; Drummond et al., 1998; Malicki et al., 1996a; Sun et al., 2004). In addition to forward genetic strategies, the zebrafish embryonic development can be analyzed using fast and inexpensive antisense approaches (Eisen and Smith, 2008; Nasevicius and Ekker, 2000). One interesting application of these antisense methods is the use of zebrafish as a tool for testing human mutant alleles associated with ciliary diseases (Khanna et al., 2009; Leitch et al., 2008; Zaghloul et al., 2010). In this chapter, we briefly describe cilia in several organs of wild-type zebrafish and point out defects that are associated with their malfunction in mutant strains. Methods to study the ciliary structure and function in the zebrafish model are presented.

II. Cilia in Zebrafish Embryos and Larvae Cilia are abundant and can be easily visualized in a number of organs in zebrafish embryos (Fig. 1). For example, the length, density, and motility of cilia have been analyzed in the Kupffer’s vesicle (KV), the pronephric duct, neurons of sensory systems, and the neural tube (Essner et al., 2005; Kramer-Zucker et al., 2005; Tsujikawa and Malicki, 2004). In the following we describe the distribution of cilia and the developmental timing of their differentiation in several organs of zebrafish. As the vast majority of zebrafish research is performed on embryos and early larvae, the descriptions of ciliogenesis that we provide in this section focus on these stages. A. Kupffer’s Vesicle KV is a closed, spherical transient structure embedded on the ventral side of the tailbud. It contains a liquid-filled lumen lined with ciliated epithelial cells that are derived from dorsal forerunner cells (Cooper and D’Amico, 1996; D’Amico and Cooper, 1997; Oteiza et al., 2008). Although in terms of its developmental origin

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[(Fig._1)TD$IG]

Fig. 1

Cilia in zebrafish. (A) The zebrafish larva at approximately 120 hpf. The locations of tissues that feature prominent cilia are indicated. (B) Confocal image of olfactory cilia stained with anti-acetylated tubulin antibody (green) at 3 dpf. F-actin is labeled with fluorophore-conjugated phalloidin (in red) to visualize the morphology of the olfactory pit. (C) Transverse cryosection through the retina at 3 dpf. Cilia are stained with anti-acetylated tubulin (green) and anti-Ift88 (red) antibodies, and imaged using confocal microscopy. The photoreceptor cell layer is indicated with a bracket. IFT88 signal is enriched at the base of cilia (red). (D) Confocal image of the anterior macula in the ear of a whole embryo at 3 dpf. Hair cells and their cilia are


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and structure the KV is fairly distinct from the mouse node, it plays a similar role in the establishment of the LR asymmetry of the body plan (Bisgrove et al., 2005; Essner et al., 2005). It is thought that the generation and sensing of directional fluid flow by cilia in the KV and the node alike are the initial symmetry-breaking events along the LR axis (McGrath et al., 2003; Nonaka et al., 2002, 1998). As a result, the KV represents an excellent system to evaluate the morphology and motility of cilia (Fig. 1J, 2B), as well as the functional consequences of ciliary defects. Cilia in the KV are best imaged during the four to eight-somite stage (around 11–13 hour postfertilization (hpf) at 28.5 C), when this structure is most prominent. Both motile and immotile cilia can be found in the KV, and most KV cells contain a single cilium (Borovina et al., 2010; Kramer-Zucker et al., 2005; Kreiling et al., 2007; Okabe et al., 2008). Since zebrafish development is very rapid, maternally deposited gene products frequently persist to quite advanced developmental stages. This needs to be taken into consideration during the analysis of zygotic mutants, since they start their life with a robust dose of maternal gene products, which frequently mask the early developmental function of analyzed genes (Cao et al., 2010; Huang and Schier, 2009).

B. Pronephros The functional kidney in the zebrafish embryo is the pronephros. During development, the pronephros originates from the intermediate mesoderm. By the 20somite stage, a pair of pronephric ducts are already formed, one on each side of the body axis (Drummond et al., 1998). The differentiation of the pronephric glomerulus takes place slightly later, and glomerular filtration commences between 36–48 hpf (Drummond et al., 1998). Ciliary defects in renal epithelial cells almost inevitably lead to the formation of kidney cysts in both zebrafish and mammals visualized using antibodies to acetylated tubulin (green). The apical cell surface marker Crumbs is also visualized via antibody staining (in red). (E) Confocal image of a crista in the ear of a whole embryo at 4 dpf. Hair cells are visualized as in (D). Phalloidin is used to visualize tissue morphology (red). (F) Transverse cryosection through the trunk exposes the proneprhic duct at 4 dpf. Its apical surface differentiates actin-rich microvilli and is easy to visualize with phalloidin (blue). A bundle of ciliary axonemes on the apical surface of a multiciliated cell is visualized with anti-acetylated tubulin antibodies (green). The apical surface of this cell is visualized with anti-Crumbs antibodies (red). (G) Confocal image of a lateral line neuromast in the whole embryo. Hair cells and their cilia are visualized with anti-acetylated tubulin staining. (H) Confocal en face image of a neuromast stained with phalloidin (green) and anti-g -tubulin antibody (red). Arrows in the inset indicate planar orientation of neuromast hair cells. (I) Spinal canal cilia visualized via the expression of a mouse Arl13b-GFP transgene. To visualize cell membranes, embryos were injected with mRNA encoding membrane-bound RFP. (J) Kupffer’s vesicle cilia labeled using antibodies to acetylated alpha tubulin in a whole embryo. (K) Transverse section through the central retina at 5 dpf. An electron micrograph showing retinal photoreceptors. Note differentiated outer segment (OS). Asterisk marks the photoreceptor cell soma. (L) An electron micrograph showing numerous cilia and villae in the lumen of the pronephric duct. A transverse section through the zebrafish trunk at 4 dpf. (M) A close-up view of panel (L) showing a cross-section through the ciliary shaft. The 9 + 2 arrangement of microtubules is evident. (N) A cross-section of a hair cell kinocilium in the ear of 3 dpf. An electron micrograph showing the 9 + 2 arrangement of microtubules. (O) Section through the ear macula showing the kinocilium (arrowheads) and a nearby bundle of sterocilia. In (C–E, and G) apical is up. Image in panel (I) courtesy of Brian Ciruna. B–D, F–H, J, and K–N are courtesy of the present and former Malicki lab members: Motokazu Tsujikawa, Yoshihiro Omori, Chengtian Zhao, and Peter Kovach. (E, O) reprinted with permission from (Tsujikawa and Malicki, 2004). (See Plate no. 3 in the Color Plate Section.)

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(Fig. 3B–F) (Pazour et al., 2002; Sun et al., 2004; Yoder et al., 2002). The zebrafish pronephric duct features two populations of ciliated cells: multiciliated cells (MCC) and single ciliated cells (SCC), which form clustered cilia bundles and single cilia, respectively (Liu et al., 2007; Ma and Jiang, 2007). Cilia on both cell types display the ‘‘9 + 2’’ configuration of microtubules, i.e., they contain a central pair of microtubules. While both MCCs and SCCs can be found in the anterior and middle segment of the pronephric duct, the posterior portion of the duct consists of only SCCs. Conveniently, cilia formation in SCCs and MCCs takes place at different time points; although single cilia can be detected as early as 20 hpf, bundles of cilia from MCCs are observed starting only at around 36 hpf. Due to the presence of maternal contribution, some cilia mutants initially display normal SCC cilia on the apical surface of pronephric duct cells at earlier time points but do not form cilia bundles later on (Cao et al., 2010). Therefore, it is useful to investigate the pronephric cilia both at an early stage (e.g., 20 hpf) and a later time point (e.g., 5 dpf). C. Sensory Organs Similar to other vertebrates, including mammals, zebrafish sensory organs feature prominent populations of cilia. These are present on the apical surface of sensory neurons, including photoreceptors, as well as olfactory and mechanosensory cells (Fig. 1B–E, G, H). It is well established that in the case of photoreceptors and olfactory cells, sensory signal transduction cascades are compartmentalized to cilia (Kennedy and Malicki, 2009; McEwen et al., 2007; Pugh and Lamb, 2000). In the following we briefly comment on the major features of cilia in the visual, auditory, and olfactory systems.

1. Photoreceptors Photoreceptor cells of the vertebrate retina acquire nearly all visual information available to the organism. Outer segments, specialized compartments that project from the apical surface of photoreceptor cells, mediate this function. Interestingly, the outer segment is a highly modified cilium that features dramatically enlarged membranes forming hundreds of parallel folds (Fig. 1K) (Rodieck, 1973; Kennedy and Malicki, 2009). Rod outer segments harbor approximately 108–109 light-sensitive opsin molecules and similarly impressive quantities of other components of the visual transduction cascade (Pugh and Lamb, 2000). As found in other types of cilia, the outer segment is supported by microtubules, which originate from the basal body located at the apical terminus of the photoreceptor soma and span its entire length (Insinna et al., 2008; Wen et al., 1982). The axoneme of the photoreceptor outer segment lacks the central microtubule pair and appears to fulfill two roles: it supports the elaborate folding of the outer segment membrane, and provides a transport route for outer segment proteins, such as opsins. Photoreceptors of the zebrafish retina first become postmitotic at 40–48 hpf, start to express opsins at about 50 hpf, and begin to form outer segments by 54 hpf (Hu and Easter, 1999; Raymond et al., 1995; Schmitt and Dowling, 1999). Based on behavioral tests, outer segments are functional by 3 dpf


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(Easter and Nicola, 1996) but continue to elongate for at least the next three weeks (Branchek and Bremiller, 1984).

2. Mechanosensory Hair Cells Similar to other vertebrates, the inner ear of zebrafish consists of interconnected canals and chambers that in several locations feature patches of specialized cells known as mechanosensory hair cells (Fig. 1D, E) (Bang et al., 2001; Haddon and Lewis, 1996; Whitfield et al., 1996). Similar to photoreceptors, hair cells also differentiate a prominent apical cilium, known as the kinocilium. The first cells that feature hair cell morphology differentiate in the otic vesicle at two locations and become first detectable around 19.5 hpf (Haddon and Lewis, 1996). They are positioned underneath otoliths, deposits of protein and inorganic salts, which appear as two round structures, one anterior to the other inside the otic vesicle (Haddon and Lewis, 1996; Malicki et al., 1996b; Riley et al., 1997). As the animal develops, new hair cells are gradually added next to the previous ones, forming sensory patches, known as maculae (Bang et al., 2001; Haddon and Lewis, 1996). Later in development, around 21–25 dpf, a third macula differentiates in the third chamber of the inner ear, the lagena (Bang et al., 2001). In addition to maculae, three other locations known as the anterior, the lateral, and the posterior crista differentiate hair cell populations at about 3 dpf (Haddon and Lewis, 1996). The kinocilia of each crista project into the lumen of a different semicircular canal, and are substantially longer than those of maculae (Haddon and Lewis, 1996). Although they seem to be immotile, they feature the 9+2 configuration of microtubules. Adjacent to the kinocilium, hair cells feature a bundle of apical protrusions, known as stereocilia. The term ‘‘stereocilia’’ is a misnomer, as these structures are supported by actin cytoskeleton and thus are related to microvilli, not cilia. The detection of sound waves by hair cells is thought to be mediated by a physical displacement of stereociliary bundles (Hudspeth, 1989). The role of kinocilium in this process is, however, unclear. In zebrafish, hair cell kinocilia are maintained throughout the adulthood and thus could play a role in hearing (Bang et al., 2001; Haddon and Lewis, 1996). In contrast, the hair cells in the adult mammalian auditory organ lose their kinocilia (Kikuchi and Hilding, 1965; Kimura, 1966). In vitro studies using explants from the adult mammalian ear show that after an induced mechanical stress, the subsequent self-repair of the damaged tissue is accompanied by a regeneration of the kinociliary axoneme, which suggests a role for kinocilia in hair cell morphogenesis (Sobkowicz et al., 1995). In line with this idea, vertebrate cilia have been linked to cell–cell signaling pathways, one of which is the planar cell polarity pathway (Cao et al., 2010; Huangfu et al., 2003; Jones et al., 2008). One characteristic feature found in aquatic vertebrates, including fish, is the presence of hair cells on the body surface, in sensory organs known as neuromasts (Fig. 1G, H). Similar to those in the ear, hair cells of neuromasts help to detect hydrodynamic movements (reviewed in, Montgomery et al., 2000). Neuromasts differentiate in a stereotypical pattern on both the trunk and the head. Neuromasts

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of the trunk and the tail belong to the posterior lateral line, while the ones on the head form the anterior lateral line (reviewed in, Dambly-Chaudiere et al., 2003; Metcalfe, 1985; Metcalfe et al., 1985). The anterior lateral line differentiates the first neuromast anterior to the ear by 30 hpf (Metcalfe, 1985). Shortly thereafter, at 36 hpf, the posterior lateral line begins to differentiate neuromasts in the rostal–caudal sequence: the anterior trunk neuromasts are the first to appear, whereas the tail ones appear last. The posterior lateral line system reaches the tail by 48 hpf and includes about six neuromasts (Metcalfe, 1985). As the larva grows, the posterior lateral line continues to differentiate additional neuromasts (Ledent, 2002), and by 5 dpf, 10–11 neuromasts are found along the myoseptum on each side of the fish (Metcalfe et al., 1985). As the organism matures, additional lateral line branches form and new neuromasts are added to preexisting branches, so that eventually over a thousand neuromasts are found in the adult zebrafish (Ledent, 2002). Interestingly, the neuromasts of the lateral line are polarized in the plane of the skin surface, a feature particularly evident in the localization of hair cell kinocilia (Lopez-Schier et al., 2004; Lopez-Schier and Hudspeth, 2006). This polarity is already prominent by 3 dpf, and is characterized by several topographical features (Fig. 1H) (Lopez-Schier and Hudspeth, 2006). First, within each neuromast the basal bodies of hair cell kinocilia are displaced toward the edge of the apical surface along a single axis in either direction (Fig. 1H, arrows in the inset). Second, hair cells of each neuromast are subdivided into two groups of approximately equal size: in one area, their kinocilia are displaced in one direction, while in the remaining area, in the opposite direction. Finally, the axis of polarization varies in neuromasts; while early differentiating neuromasts polarize along the anterior–posterior axis, the ones differentiating later display dorsoventral polarity (Lopez-Schier et al., 2004). Given the ease of neuromast identification, their early differentiation, and accessibility, the lateral line neuromasts provide an attractive opportunity to study the genetic bases of planar cell polarity (Lopez-Schier and Hudspeth, 2006; Lopez-Schier et al., 2004; McDermott et al., 2010; Raible and Kruse, 2000)

3. Olfactory Sensory Neurons The olfactory system is also ciliated in both zebrafish and other vertebrate embryos (Hansen and Zeiske, 1993; Kimura et al., 2009; Menco and Farbman, 1985). In zebrafish, the openings of olfactory pits appear between 34–36 hpf, and their inside surfaces harbor ciliated dendritic endings of olfactory sensory neurons (Hansen and Zeiske, 1993). As olfactory pits gradually enlarge during embryogenesis, the number of cilia increases (Fig. 1B) (Hansen and Zeiske, 1993). On the structural level, zebrafish olfactory cilia feature both 9 + 0 and 9 + 2 microtubule configurations (Wloga et al., 2009). Similar to photoreceptor outer segments as well as some invertebrate cilia, the tips of zebrafish olfactory cilia contain microtubule singlets (Ward et al., 1975; Wloga et al., 2009). Consistent with the 9 + 0 and 9 + 2 configurations, the olfactory systems of both frog and zebrafish contain motile and nonmotile cilia (Drummond, 2009; Mair et al., 1982). Both types are easily detectable using antibody staining of whole animals.


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D. Neural Tube By about 18 hpf, the zebrafish neural keel differentiates the central canal and begins to acquire the form of a tube (Kimmel et al., 1995). In adult vertebrates, the inside lining of the spinal canal is provided by the ciliated ependymal epithelium (Nakayama and Kohno, 1974; Worthington and Cathcart, 1963). Also in zebrafish embryos and early larvae, the presence of cilia has been well documented on the apical surfaces of cells that line the spinal canal lumen (Fig. 1I) (Borovina et al., 2010; Kramer-Zucker et al., 2005). Despite their 9 + 0 microtubule configuration, these cilia are motile and are required to propagate the flow of the cerebrospinal fluid inside brain ventricles and the spinal canal (Kramer-Zucker et al., 2005; Worthington and Cathcart, 1966). The spinal canal cilia have been proposed to play developmental roles (Sawamoto et al., 2006). In zebrafish, the disruption of spinal canal cilia leads to the abnormal expansion of brain ventricles that resembles hydrocephalus (Kramer-Zucker et al., 2005). Finally, it is worth noting that basal bodies of zebrafish neuroepithelial cells are displaced to the posterior region of the apical surface and thus display planar polarization (Borovina et al., 2010).

III. Analytical Tools for Cilia Morphology and Motility In this section, we describe general approaches used to detect cilia. Labeling of the ciliary axoneme using antibodies to microtubules has been the most common approach. There are several commercially available monoclonal markers that recognize various posttranslational modifications of tubulin (Table I). The most commonly used antibody recognizes acetylated a-tubulin (Fig. 1) (Duldulao et al., 2009; Kishimoto et al., 2008; Omori and Malicki, 2006; Tsujikawa and Malicki, 2004; Zhao and Malicki, 2007). Although this marker has been used extensively, it is not entirely specific to cilia as acetylated a-tubulin is also present in the cytoplasm of some cells, such as hair cells (Fig. 1D, E) (Omori and Malicki, 2006). Likewise, axons of retinal ganglion cells also contain richly acetylated microtubules. Antibodies that recognize other modifications of tubulin, such as glutamylation or glycosylation, can also be used to label cilia (Table I, Wloga et al., 2009). In addition, we have developed a rabbit polyclonal anti-Scorpion/Arl13b (anti-Sco) antibody that is also a robust and specific marker for cilia (Duldulao et al., 2009) (Fig. 2A). Small aliquots of anti-Sco can be requested from the authors and a detailed protocol for immuno-staining and imaging has been described elsewhere (Drummond, 2009). Detection of the basal body is another way to reveal the position of cilia (Fig. 1H). A number of monoclonal antibodies to g -tubulin recognize this structure. A transgenic line expressing GFP-tagged centrin has been used as well (Borovina et al., 2010; Zolessi et al., 2006). Although not useful in visualizing the axoneme, these methods help to survey for the potential displacement of basal bodies in cilia mutants. Antibodies and transgenic lines that have been used in the analysis of zebrafish cilia are listed in Table I.

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[(Fig._2)TD$IG]

Fig. 2 Localization of proteins in cilia. (A) Sco/Arl13b is highly enriched in the cilia of the pronephric duct. A confocal image of a whole embryo stained with anti-Sco/Arl13b antibody (in green), and an antiacetylated tubulin antibody (a-tub, in red). (B) tg(actin::sco-eGFP) embryos show cilia-specific eGFP signal (green). The left panel shows a fixed embryo counterstained with DAPI (in blue) to visualize nuclei. Inset shows an enlargement. The middle and right panels show snapshots of a live embryo featuring both motile (red arrows, note the change of position in the two pictures) and immotile (yellow arrow, note the same position in the two images) cilia in the KV. (C-D) Seahorse localization shown by immunolabeling with anti-Seahorse (Sea, in green) and anti-acetylated tubulin (a-tub, in red) antibodies. C shows results from co-incubation of the two antibodies, while D shows results from sequential incubations. (See Plate no. 4 in the Color Plate Section.)


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Motility is an important aspect of ciliary function. High-speed videomicroscopy techniques allow for qualitative and quantitative analysis of cilia beat frequency, amplitude and waveform in a variety of ciliated organs (Kramer-Zucker et al., 2005; Okabe et al., 2008; Zhao and Malicki, 2007). These approaches utilize brightfield or DIC microsocopy and a high-speed camera to visualize cilia movement in the KV, the pronephric duct, the olfactory pit, and the spinal canal. It has been, however, challenging to visualize immotile cilia via these imaging methods. A slight modification to these approaches allows for the visualization of both motile and immotile cilia by high-speed fluorescence microscopy (Fig. 2B). An essential component of this method is the use of embryos expressing GFP-tagged cilia markers, such as Scorpion/Arl13b (Fig. 2B) (Duldulao et al., 2009). We outline two protocols for live imaging of fluorescently labeled cilia that can be applied to various ciliated organs such as the KV, the pronephric duct, and the spinal central canal.

A. Method 1: Imaging of Kupffer’s Vesicle Cilia Following arl13b/sco-eGFP mRNA Overexpression Microinjection of in vitro transcribed scorpion/arl13b-eGFP mRNA allows for robust and versatile GFP-labeling of cilia as the amount and concentration of transcript can be adjusted as desired. This method is particularly useful for imaging cilia at early developmental stages. The following describes the method for imaging motile and immotile cilia in KV.

1. Materials

Standard microinjection reagents (borosilicate micropipettes, microinjection apparatus, agar injection molds, embryo medium, phenol red, etc.) (Yuan and Sun, 2009). Methods 1 and 2: scorpion/Arl13b eGFP (zebrafish scorpion/arl13b, eGFPtagged) RNA, in vitro transcribed from linearized DNA template as described previously (Yuan and Sun, 2009). Alternative Method: Tg(bact::arl13b-GFP) transgenic fish line. 1.5% low-melting point agarose solution (in embryo medium) in small glass petri dish on a 50 C heat block. Viewing chamber made from a large microscope slide (25 60 mm) and a large coverslip (22 40 mm) separated by two ‘‘spacer’’stacks of 3 small #1 coverslips (22 22 mm); glued together with ‘‘Super Glue’’ (cyanoacrylate). The space between the slide and the coverslip fits a living zebrafish embryo without crushing it, thus enabling in vivo imaging using florescence microscopy. Prepare the chambers ahead of time by gluing together two stacks of three small coverslips to a large microscope slide; position them on the slide 3 cm apart from each other. During the experiment, embryos will be embedded in agar directly on the surface of the large the coverslip, which is then inverted and placed between the small coverslip spacers and the slide (for detailed diagrams, refer to Westerfield, 2000).

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Fine-tipped Dumont forceps or tungsten needles (Fine Science Tools, 11252-40 or 10130-05). Clear nail polish. An inverted microscope equipped for imaging of fluorescently labeled samples and a Coolsnap HQ2 (Photometrics) cooled-CCD camera or equivalent. MetaMorph (Molecular Devices) or ImageJ (NIH) software.

2. Protocol 1. Following the previously described method (Yuan and Sun, 2009), inject wildtype zebrafish embryos at the one-cell stage with 250 pg of in vitro transcribed scorpion-eGFP RNA. It is possible to inject embryos as late as the four-cell stage. We found, however, that the earlier the injection the more evenly distributed the injected mRNA will be. Varying the amount of injected transcript allows one to control expression levels. 2. At the 4–6 somite stage (11–12 hpf), dechorionate embryos manually using fine-tipped Dumont forceps. 3. Using a glass, briefly immerse 5–10 embryos in a 1.5% low-melting point agarose solution for 5–10 s on a 50 C heat block. 4. Using the same pipette, take up the embryos and a small amount of agarose and place them in the middle of a large glass coverslip (22 40 mm). Quickly move to the next step. 5. Gently adjust embryos in the agarose using a pair of fine-tipped Dumont forceps or a tungsten needle so that the tailbud touches the coverslip. We recommend positioning the embryo in such a way that the anterior–posterior axis of the KV is parallel to the surface of the coverslip. This method positions the KV as closely as possible to the surface of the coverslip, thereby minimizing the optical depth required to image it. Work quickly as the agarose will begin to solidify rapidly at room temperature (RT). 6. Wait 5 min for the agarose to completely solidify at RT. 7. Using forceps, pick up the coverslip. Invert it so that the side with the agarembedded embryos is facing down. 8. Mount the coverslip onto the viewing chamber slide, so that it spans the 3 cm space created between two stacks of coverslips as described in materials above. Embryos should be facing the interior of the chamber. 9. Apply a small amount of nail polish to the corners of the coverslip to prevent it from moving. Let the nail polish dry for 5 min. 10. Use a Pipetteman to inject 100 to 200 mL of embryo medium into the interior of the chamber. 11. Image embryos on an inverted microscope with a 25 or 40 water immersion objective and a Coolsnap HQ2 (Photometrics) camera using GFP fluorescence. The KV will be located at the tip of the tailbud and should appear as a spherical structure posterior to the notochord and somites. Using this mounting method, the KV should be easily visible as it will be located very close to the coverslip surface.


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12. Once you locate the KV, use the fine focus to determine an optimal Z-section. Acquire 3 to 5 time-series of immunofluorescence images per each embryo at different Z-planes within the KV. Collect all time series at the rate of 50 fps (frames per second) with a 5 ms exposure time and a 320 320 pixel resolution (fine adjustments to these settings may be required depending on your microscope setup). In our experience, a length of 20 s for each time-series is optimal. It is worth noting that this acquisition speed is insufficient for measuring beat frequency, but will allow one to distinguish motile from immotile cilia. 13. After the acquisition of each time-series, record a short time-series in DIC or brightfield illumination to serve as an overlay for data analysis. Be sure to mark the antero–posterior orientation of each KV. This will aid in further analyses. 14. View each time-series at 10 fps for analysis using MetaMorph (Molecular Devices) or ImageJ (NIH) software. Be sure to distinguish between immotile and motile populations of cilia. This protocol can be modified to visualize other ciliated organs such as the pronephric duct and spinal central canal, as described below.

B. Method 2: Modified Methods for Imaging of the Pronephric Duct and Spinal Central Canal Using arl13b/sco-eGFP Over-Expressants

1. Additional Materials

0.003% Phenylthiourea (PTU) in embryo medium; 3% Methycellulose: add 3 g methycellulose (Sigma, M7027) to 100 ml embryo medium, stir overnight at 4 C, spin in a benchtop centrifuge for 10 min, store at 4 C; 25 MESAB solution: dissolve 0.4 g of ethyl 3-aminobenzoate methanesulfonate salt (Sigma, A5040) in 100 ml of 10 mM Tris (pH 7.0), store frozen.

2. Protocol 1. Following the previously described method, inject wild-type zebrafish embryos at the one-cell stage with 250 pg of in vitro transcribed arl13b/sco-eGFP RNA. 2. To prevent the appearance of pigmentation at later stages of development, raise embryos in 0.003% PTU in embryo medium after 20–24 hpf. 3. At 2–5 dpf, dechorionate embryos manually by using fine-tipped Dumont forceps. 4. Anesthetize embryos in 1 MESAB in embryo medium for 1 to 3 min. 5. Using a pipette tip, place a small drop of 3% methylcellulose in embryo medium at the center of viewing chamber slide between the stacks of small coverslips. 6. Using a glass Pasteur pipette, remove 3 to 5 embryos from the embryo medium/ MESAB solution and place them onto the drop of methylcellulose on the viewing chamber slide.

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7. Gently adjust embryos/larvae in methylcellulose using a pair of fine-tipped Dumont forceps or a tungsten needle so that the side of each embryo is parallel to the coverslip. 8. Using forceps, mount a medium coverslip (22 40 mm) onto the viewing chamber. Make sure that the embryos are well positioned in the interior of the chamber. 9. If the embryo is not satisfactorily oriented after coverslip placement, the coverslip can be gently pushed to improve the orientation. This can be done without damage to the embryo due to the viscosity of the methylcellulose solution. 10. Apply a small amount of nail polish to the corners of the coverslip to prevent it from moving. Let the nail polish dry for 5 min. 11. Image embryos on an inverted fluorescence microscope equipped with a 25 or 40 water immersion objective and a Coolsnap HQ2 (Photometrics) camera using GFP fluorescence. We recommend locating the pronephric duct by looking directly posterior to the otic vesicle. It will appear as a tubule lined with motile cilia. The pronephric duct continues posteriorly along the body axis following closely the gut to the cloaca. The spinal cord can easily be located by searching through the dorsal portion of the embryo. It will appear as a tube lined with cells featuring single motile cilium. The spinal cord extends from the head of the embryo down to the tail, following the notochord closely. This mounting method enables the visualization of the pronephric duct and the spinal cord, since they will be located very close to the coverslip. 12. Follow Steps 12–14 of the previous protocol for further imaging and analysis. C. Alternative Method: Imaging of Cilia Using arl13b/scorpion-eGFP Transgenic Fish An alternative method is to utilize a transgenic line that constitutively expresses Arl13b/Scorpion-eGFP. Since injected RNA transcripts gradually degrade as the embryo develops, the level of expression may fade by 3 dpf depending on the stability of that particular transcript and the translated product. To circumvent this difficulty and enable analysis at later stages in embryonic development, we generated a transgenic line. A similar line using mouse Arl13b was also generated by the Ciruna group (Borovina et al., 2010). Similar to microinjection of arl13b/sco-eGFP RNA, arl13b/scorpion-eGFP transgenic embryos can be used to image a variety of ciliated organs including the KV (Fig. 2B), the pronephric duct and spinal central canal. The two protocols described above can be adapted to the use of the scorpion/arl13b-GFP transgenic line by simply omitting the microinjection step. D. Method 3. Detection of Protein Expression in Cilia Using Immunohistochemistry An increasing number of proteins have been found in the cilium. To examine whether a protein of interest localizes to the cilium, co-immunostaining with anti-acetylated tubulin (clone 6-11B-1, Sigma) and an antibody against the protein


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of interest, followed by confocal imaging, is often the method of choice. We found, however, that the extreme high affinity of the anti-acetylated tubulin antibody for cilia may lead to artifacts. An example is shown in Figure 2C. When zebrafish embryos fixed with methanol were probed with combined primary antibodies (affinity purified anti-Seahorse, 1:100; anti-acetylated tubulin, 1:20,000), Seahorse appears to be enriched on pronephric ductcilia. However, when antibody incubations are carried out sequentially, the ciliary localization of Seahorse disappears (Fig. 2C). Further control experiments using anti-Seahorse antibody in the absence of antiacetylated tubulin antibody detected no enrichment of this protein on the cilium (not shown), suggesting that the ciliary enrichment of Seahorse shown in Figure 2C is a false positive result. A similar false positive signal was detected for a number of other antibodies when used in combination with an anti-acetylated tubulin antibody (Sun lab, unpublished). Co-immunolabeling for acetylated tubulin thus needs to be performed keeping the above considerations in mind. Appropriate dilution of antibodies, sequential staining, and careful controls are essential to circumvent this problem. Below, we describe a protocol for sequential labeling.

1. Materials

Anti-acetylated tubulin, clone 6-11B-1, Sigma T6793 (Table I). Dent’s fixative: one part of DMSO mixed with four parts of methanol. 30% hydrogen peroxide. PBT: PBS with 0.5% Tween-20. Blocking solution: PBT + 10% serum. Use serum that matches the species in which secondary antibodies were generated. Secondary antibodies: Jackson ImmunoResearch (West Grove, PA). Mounting medium: we use Vectorshield Hardset Mounting Medium (Vector Laboratories, H-1400).

2. Protocol 1. Fix embryos with Dent’s fixative overnight. For embryos younger than 30 hpf, use precooled Dent’s fixative and incubate at –20 C. For older embryos, use Dent’s fixative at RT and incubate at RT. 2. Bleach pigment by incubating embryos in 1:2 hydrogen peroxide/methanol solution for 2 hours to overnight at RT. 3. Wash with methanol three times, 5 min each. 4. Rehydrate embryos by incubating in 50:50 PBT/methanol and PBT sequentially for 5 min each. 5. Block embryos in the blocking solution for 30 min. 6. Incubate embryos in the blocking solution and a 1:20,000 dilution of antiacetylated tubulin antibody at RT for 2 h or at 4 C overnight. 7. Wash embryos with PBT five times, 20 min each. 8. Incubate embryos with a fluorophore-conjugated anti-mouse IgG at the concentration recommended by the manufacturer for 2 h at RT.

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9. Wash embryos with PBT five times, 20 min each. 10. Incubate embryos with an antibody developed in a non-mouse host (because the cilia marker 6-11-B anti-acetylated tubulin antibody is a mouse monoclonal antibody) against the protein of interest in blocking solution at RT for 2 h or at 4 C overnight. 11. Wash embryos with PBT five times, 20 min each. 12. Incubate embryos with an appropriate secondary antibody. 13. Wash embryos with PBT five times, 20 min each. 14. Place stained embryos in mounting medium. Using Dumont forceps, carefully remove as much yolk and muscle as possible. For pronephric duct staining, add coverslip and press the coverslip firmly against the slide. The embryos will be crushed, but usually the pronephric duct will stay intact. You may need to perform tests to determine the proper way to dissect the tissue of your choice. Allow the mounting medium to harden by leaving slides at RT for 30 min. The specificity of an antibody against the protein of interest needs to be tested rigorously. One of the most convincing controls is the absence or decrease of signal in mutants or morphants. The optimal dilution of the antibody should also be tested empirically, since too high a concentration will lead to high background whereas too low a concentration will not provide sufficient signal intensity. IgG purification or affinity purification frequently improves the signal to noise ratio. Many vendors for custom antibody production provide purification services. Alternatively, one can purify antibodies in house following well-established protocols (Harlow and Lane, 1988).

IV. Analysis of Cilia-related Mutant Phenotypes in Zebrafish A. Left–Right Asymmetry Defects The transparency and external development of the zebrafish embryo allow for direct inspection of morphological phenotypes in live embryos. In addition, tools such as in situ hybridization, immunohistochemistry, and histology can also be utilized to analyze mutant phenotypes in depth. One common phenotype associated with cilia defects is abnormal left-right asymmetry. The breaking of bilateral symmetry is believed to be triggered by cilia-driven fluid flow in the KV, similar to that described in the mouse node (Essner et al., 2002). Defects in nodal or KV cilia disrupt the establishment of the LR asymmetry of the body plan, leading to misplacement of multiple internal organs, such the heart (Nonaka et al., 1998). B. Method 4: Evaluation of Heart Positioning in Live Zebrafish Embryos

1. Materials

Zebrafish embryos at 30 hpf. Dissecting microscope. Dumont forceps.


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[(Fig._3)TD$IG]

Phenotypes of zebrafish cilia mutants. (A) Side view of an arl13b/scohi459 mutant embryo (sco) and its wild-type sibling (wt) at 36 hpf. (B) A scohi459 mutant at 3 dpf. Arrow points to a kidney cyst. (C–F) Plastic transverse sections through the zebrafish trunk at 50 hpf. Tissue architecture is visualized via hematoxylin/eosin staining (Method 5). (C, D) Sections through the region of the glomerulus (arrowhead) and the pronephric tubule (arrow) in the wild-type (C) and scohi459 mutant (D) animals. (E, F) Sections through the duct region (arrows) in the wild-type (E) and scohi459 mutants (F). (G–I) Ventral views of embryos at 30 hpf that display heart position (red circles) at the left side (G, L), middle (H, M), and the right side (I, R) of the body axis. Arrowheads indicate the midline. (J, K) Confocal images showing cilia (stained with anti-acetylated tubulin antibody, in red) in the pronephric duct (stained with anti-Cdh17 antibody in green) in an arl13b/scohi459 mutant embryo (K, sco), compared to a wide-type sibling (J, wt) at 33 hpf. Scale bar: 10 mm. Arrow in (J) points to cilia bundle from MCCs while the arrowhead points to cilia from SCCs. (L-N) Plastic (JB4) sections of wild-type and IFT mutant retinae stained using Methylene Blue/Azure II (Method 6). (L) The wild-type retina features elongated photoreceptor cell nuclei (arrow). The black tissue next to photoreceptor cells is the retinal pigmented epithelium. Lightly stained features (arrowhead) are outer segments. (M, N) The photoreceptor cell layer of animals mutant for IFT-related genes, flr and ovl, display defective photoreceptor morphology and the lack of outer segments. Cell death in the photoreceptor cell layer results in the appearance of empty spaces (arrows in M, N). The asterisk shows optic nerve. (O, P) Confocal images of transverse cryosections though the retina. Double cones are visualized with the Zpr-1 antibody in a wild-type (O) and elitp49 mutant (P) retinae (green signal). Mueller glia are identified using anti-carbonic anhydrase antibody (red). (P) Mutations in the gene encoding an IFT particle component, elipsa (eli), lead to a degeneration of photoreceptor cells. Older photoreceptors in the central retina are lost first. (Q, R) Transverse plastic sections through the ear of the wild type (Q) and its oval mutant sibling (R) at 5 dpf. Larvae were embedded in JB4, sectioned, and stained with Methylene Blue/Azure II. Arrow points to hair cell nuclei, which are absent in ovl mutant individuals. (S, T) DiI labeling (Method 11, red signal) of whole embryos stains olfactory pits of a wild-type (S) but not ovl mutant (T) animals. A–F, J, and K are reprinted with permission from Duldulao et al., 2009; G-I from Cao et al., 2009; L-P from Doerre and Malicki, 2002; and Q-T from Tsujikawa and Malicki, 2004. (See Plate no. 5 in the Color Plate Section.)

Fig. 3

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2. Protocol Owing to the transparency of the zebrafish embryo at 30 hpf, the heart is detectable using a dissecting microscope (Fig. 3G-I). Use a pair of tweezers to turn the embryo to a ventral side up position, which allows for easier visualization of the beating heart. The heart of most wild-type embryos at this stage is located on the left side of the embryo, whereas cilia mutants often feature hearts that are either positioned at the center or on the right side (Fig. 3G–I). If mutants develop other obvious morphological phenotypes, such as body curvature, separate mutant and wild-type individuals based on these phenotypes first, then count and record the heart position of every embryo. Perform statistical analysis using results from multiple crosses to determine whether abnormal heart position is observed in significantly higher percentages in mutant embryos, compared to their wild-type siblings. It is worth noting that some zebrafish strains have a high incidence of spontaneous LR defects. Therefore, it is critical to inspect background LR frequency in a particular strain so that experiments are properly interpreted. In zebrafish, the products of many genes associated with cilia formation and/or function are maternally deposited, which can mask the early functions of these genes (Cao et al.). As a result, some cilia mutants will not show a LR asymmetry defect. It is possible to use a morpholino oligo targeting the translation initiation site to block expression of maternally deposited mRNA. In such an experiment proper controls need to be included, such as a rescue of the morphant phenotype, a second nonoverlapping morpholino, a control morpholino with mismatched bases to rule out toxicity or off-target effects. Alternatively, a maternal-zygotic mutant can be generated to study a true null mutant by genetically removing both the maternal and the zygotic contribution of the analyzed gene (Ciruna et al., 2002).

C. Alternative Method: Analysis of Left–Right Asymmetry Defects Using in situ Hybridization In addition to observing the position of internal organs in zebrafish embryos, one can also analyze the distribution of molecular markers important in the establishment of LR asymmetry using in situ hybridization. Southpaw (spaw) is one of the earliest markers of LR asymmetry in zebrafish embryos (Long et al., 2003). In wildtype embryos, spaw is expressed in the left lateral plate mesoderm, while embryos with LR asymmetry defects show left-sided, right-sided, bilateral, or absent spaw expression (Long et al., 2003). Note that the expression of spaw is dynamic and its asymmetric expression is most obvious between the 18–22-somite stages. Other useful markers for LR asymmetry include pitx2, lefty1, and lefty2 (Bisgrove et al., 1999; Campione et al., 1999; Zhao and Malicki, 2007). A detailed protocol for in situ hybridization in zebrafish embryos can be found at: http://zfin.org/ZFIN/Methods/ ThisseProtocol.html (see also Westerfield, 2000).


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D. Kidney Cysts Cilia mutants almost inevitably develop kidney cysts and ventral body curvature (Kramer-Zucker et al., 2005; Sun et al., 2004) (Fig. 3A-F, J, K). Here we describe a method to analyze kidney cysts formation in zebrafish embryos. E. Method 5: Detection of Kidney Cysts in Zebrafish

1. Materials

Dissecting microscope. Day 2–5 zebrafish embryos. Embryo medium. 3% methycellulose (as in Method 2). 25 MESAB solution (as in Method 2). Bouin’s fixative: Polysciences, 16045-1. JB-4 plus embedding kit: Polysciences, 18570. JB-4 infiltration and embedding solutions: prepare following manufacturer’s instructions. Harris’ hematoxylin: Sigma, HHS16. ammonia water: mix 500 ml water with 1.5 ml ammonium hydroxide (VWR, VW3571-1), make fresh before use. stock solution of Eosin Y: dissolve 1 g Eosin Y (Mallinckrodt, 15086-94-9) in 100 ml deionized H2O. stock solution of Phloxine-B: dissolve 1 g Phloxine-B (Sigma, 18472-87-2) in 100 ml deionized H2O. Eosin-Phloxine solution: add 100 ml stock Eosin and 10 ml stock Phloxine to 780 ml deionized H2O, add 4 ml glacial acetic acid, bring volume to 1000 ml with deionized H2O.

2. Protocol Kidney cysts are frequently located in the glomerular–tubular region. Given the transparency of both the embryo and the pronephros, the latter is usually invisible in the wild type when using a dissecting scope. The cyst, on the other hand, has a bubblelike appearance, slightly anterior and medial to the pectoral fin (Fig. 3B, arrow). The cysts are often first visible by 2 dpf, and they tend to enlarge as the embryo develops. The kidney duct is also often dilated. The presence of kidney cysts and dilated ducts can be verified via histological sections as detailed below (Fig. 3C-F).

Fixing, embedding and sectioning 1. 2. 3. 4.

Anaesthetize embryos in embryo medium with 1 MESAB. Fix embryos in Bouin’s fixative at RT overnight. Wash embryos with PBT five times, 5 min each. Incubate embryos in JB-4 infiltration solution at RT for 1 h.

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5. Replace JB-4 infiltration solution with a fresh aliquot; incubate at RT for 1 h to overnight. 6. Replace with fresh embedding solution. 7. Transfer embryos into a mold. Do not use a silicone-coated mold, as it might inhibit the polymerization of JB4 resin. Fill the mold with embedding solution. Orient embryos with a pipette tip or a needle. 8. To facilitate the polymerization of the resin, cover the mold with parafilm. 9. The embedding solution will polymerize in approximately 40 min at RT. 10. Section embryos at 4 mm with a microtome. Typically, 8–10 sections can be collected on each slide.

Hematoxylin/Eosin staining 1. Stain slides in Harris’ hematoxylin for 10 min. For even staining, quickly dip slides in the solution several times and then let them sit in hematoxylin for 10 min. 2. Dip slides in deionized H2O briefly. 3. Dip slides in ammonia water 2–3 times. Sections should turn bright blue. 4. Dip slides in deionized H2O. 5. Stain with Eosin-Phloxine for 5 min at RT. 6. Dip in deionized H2O. 7. Check with a dissecting scope to make sure that staining is sufficiently intense. Repeat the staining procedure if necessary. 8. Air dry at RT or dry in an incubator at 65 C. 9. Add mounting medium (Permount, Fisher, SP15-100) and coverslip. 10. Observe sections using a microscope; nuclei should appear blue while cytoplasm should have a pink appearance.

F. Degeneration of Sensory Neurons Mutations of cilia-related genes, such as those encoding IFT proteins, lead to a degeneration of neurons in the visual, auditory, and olfactory systems (Doerre and Malicki, 2002; Gross et al., 2005; Krock and Perkins, 2008; Omori et al., 2008; Tsujikawa and Malicki, 2004). Below we describe several methods used to monitor ciliated cells in sense organs.

Visual System Cilia-related defects in photoreceptor cells can be monitored using several approaches. The simplest one is to evaluate the gross appearance of mutant retina by analyzing plastic histological sections (Fig. 3L-N). Using a microscope equipped with Nomarski optics to view sections stained to highlight cell nuclei, one can distinguish photoreceptor outer segments (arrowhead in Fig. 3L), a feature


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frequently abnormal in cilia mutants (Doerre and Malicki, 2001). In severely affected mutant retinae, the loss of photoreceptors results in the appearance of obvious holes in the outer retina (arrows in Fig. 3M–N). Immunohistochemical analysis of protein expression is an informative way to follow histological studies. The Zpr-1 antibody stains the entire cell body of red– green double cone photoreceptor cells. This antibody can be used both on whole embryos and on frozen sections (Larison and Bremiller, 1990). Zpr-1-positive cells are found throughout the entire photoreceptor cell layer by 3 dpf (Fig. 3O) (Larison and Bremiller, 1990). In addition, staining of retinae with anti-opsin antibodies is very informative, as the transport of opsin is almost invariably affected in cilia mutants. Defects in IFT-dependent processes, for example, lead to opsin accumulation in the cell body (Doerre and Malicki, 2002; Tsujikawa and Malicki, 2004). Several anti-opsin antibodies are available for zebrafish (Table I, see also Avanesov and Malicki, 2010). Lastly, another useful approach to the analysis of ciliary (i.e., outer segment) defects in zebrafish photoreceptors is electron microscopy. Outer segments differentiate a very distinctive array of parallel membrane folds (Fig. 1K), which can be visualized on electron micrographs in exquisite detail. This membrane architecture is often disrupted or absent in cilia mutants (Doerre and Malicki, 2002; Krock and Perkins, 2008). Electron microscopy of photoreceptors is performed using standard approaches (Doerre and Malicki, 2001; Schmitt and Dowling, 1999). In rare but interesting cases, cilia-related mutations may affect a subset of photoreceptors (Zhao and Malicki, unpublished). For example, rods may degenerate but not cones, or vice versa. To distinguish subtypes of photoreceptors, one can use the opsin antibodies mentioned above. In situ hybridization with antisense opsin RNA antiopsin probes can also be used for the same purpose. Alternatively, transgenes that express GFP in a subset of photoreceptors are available (see, for example, Fadool, 2003). More information on photoreceptor cell markers, including transgenes, has been provided elsewhere (Avanesov and Malicki, 2010).

G. Method 6: Evaluation of Photoreceptor Cell Layer Morphology on Plastic Sections

1. Materials

PBST: PBS + 0.1% Tween-20. Fixative: 4% PFA (Sigma, P6148) in PBST. Ethanol: Sigma, 279741-1L. JB4 embedding kit (as in Method 5). Methylene Blue/Azure II staining solution: Methylene blue (Sigma, MB-1), 0.13%; Azure II (Sigma, A2507), 0.02%; Glycerol, 10%; Methanol, 10%; 45 mM phosphate buffer, pH 6.9). Deionized H2O. Frosted microscope glass slides: Fisher, 12-544-3. Permount: Fisher, SP15-500.

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Table I Markers of cilia and ciliated cells in zebrafish Name/specificity

Marker type/ concentration

Localization

References/Sources

CILIARY AXONEME Acetylated tubulin

Ab, mono 1:1000–1:20,000

Ubiquitous cilia marker

AXO49 (polyglycylated tubulin)

Ab, mono 1:2000

GT335 (glutamylated tubulin)

Ab, mono 1:400

TAP952 (monoglycylated tubulin)

Ab, mono 1:2000

Ab, poly 1:200–1:2000 arl13b/sco-eGFP mRNA bact::arl13b(hennin)-GFP Transgene

Cilia in the nasal epithelium, neuromast hair cells, medial section of pronephros Cilia in olfactory placode, PND, hair cells, cilia in other organs Cilia in olfactory placode, otic vesicle, PND, spinal cord, hypochord, neuromast hair cells Cilia in olfactory placode, CNS, PND, KV Cilia in PND, and other organs Cilia in PND, CNS, KV

Gamma tubulin

Ab, mono 1:200

Ubiquitous basal body marker

centrina-GFP GFP-centrinb

mRNA mRNA

CNS, possibly other organs Retinal neuroepithelium, possibly in other organs

opn1sw1::GFP UV opsin

Transgene Ab, poly 1:1000 RNA probe

UV cones UV cone outer segments

Ab, mono 1:250 Ab, poly 1:200 RNA probe

Red–green double cones

Scorpion (sco)/Arl13b

(Duldulao et al., 2009; Tsujikawa and Malicki, 2004) Sigma, clone 6-11B-1 (Callen et al., 1994; Wloga et al., 2009) (Pathak et al., 2007; Zhao and Malicki, unpublished, Wloga et al., 2009; Wolff et al., 1992) (Callen et al., 1994; Wloga et al., 2009)

(Duldulao et al., 2009) (Duldulao et al., 2009) (Borovina et al., 2010)

BASAL BODIES (Duldulao et al., 2009; Tsujikawa and Malicki, 2004) Sigma, clone GTU88 (Borovina et al., 2010) (Zolessi et al., 2006)

PHOTORECEPTORS

UV opsin Zpr1 (FRet 43) Red opsin Red opsin

Blue opsin Blue opsin Green opsin

UV cones

Red cone outer segments Red cones

Ab, poly 1:200 RNA probe

Blue cone outer segments

Ab, poly 1:500

Green cone outer segments

Blue cones

(Takechi et al., 2003) (Doerre and Malicki, 2001; Vihtelic et al., 1999) (Hisatomi et al., 1996; Takechi et al., 2003) (Larison and Bremiller, 1990) ZIRC (Doerre and Malicki, 2001; Vihtelic et al., 1999) (Chinen et al., 2003; Raymond et al., 1995; Takechi and Kawamura, 2005) (Doerre and Malicki, 2001; Vihtelic et al., 1999) (Chinen et al., 2003; Raymond et al., 1995; Vihtelic et al., 1999) (Doerre and Malicki, 2001; Vihtelic et al., 1999) (Continued)

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Table I (Continued) Name/specificity

Marker type/ concentration

Localization

References/Sources

Green opsin

RNA probe

Green cones

Zpr3 (FRet11)

Ab, mono 1:300 Ab, poly 1:1000 RNA probe

Rods

(Chinen et al., 2003; Raymond et al., 1995; Vihtelic et al., 1999) (Schmitt and Dowling, 1996) ZIRC (Doerre and Malicki, 2001; Vihtelic et al., 1999) (Chinen et al., 2003; Raymond et al., 1995) (Perkins et al., 2002) (Fadool, 2003)

Rod opsin Rod opsin

Rod outer segments Rods

xops1.3::opsin-GFP-CT44 Transgene Transgene xops::EGFPa

Rod outer segments Rods MECHANOSENSORY HAIR CELLS

Acetylated tubulin

Ab, mono 1:1000 - 1:20 000

Cilia and somata of hair cells

HCS-1

Ab, mono 1:20 - 1:250

Somata of hair cells

brn3c::GFP

Transgene

Somata of hair cells

Olfactory marker protein

RNA probe

DiI

Lipophilic Tracer

OLFACTORY SENSORY NEURONS Olfactory sensory neurons Olfactory sensory neurons

(Jing and Malicki, 2009; Tsujikawa and Malicki, 2004; Zhao and Malicki, 2007) Sigma, clone 6-11B-1 (Gale et al., 2000; Lopez-Schier and Hudspeth, 2006; Schibler and Malicki, 2007) (Xiao et al., 2005) (Tsujikawa and Malicki, 2004; Yoshida et al., 2002) (Dynes and Ngai, 1998; Tsujikawa and Malicki, 2004), Invitrogen N22880

KV, Kupffer’s vesicle; PND, pronephric duct. a b

derived from xenopus. derived from zebrafish.

2. Protocol Fixing, embedding and sectioning Use JB-4 embedding and sectioning protocol as described in Method 5. We prefer transverse sections. Ideally, they should contain the optic nerve, which can serve as a point of reference to facilitate comparisons among samples (asterisk in Fig. 3N).

Methylene blue-Azure II staining 1. Dip slides with sections into staining solution for 10 s at RT. 2. Remove and rinse immediately with a generous amount of tap water. If staining is on the faint side, then restain as above. It is suggested to test the timing of this staining technique on dispensable sections first. 3. Allow slides to dry completely. Apply 20–40 mL of Permount and add a coverslip. 4. Slides may be stored at RT indefinitely.

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H. Method 7: Analysis of Photoreceptor Cells on Cryosections via Immunohistochemistry

1. Materials

Anti-opsin antibodies, Zpr-1, see also Table I. PTU solution (as in Method 2). PBST: PBS + 0.1% Tween-20. Fixative: 4% PFA (Sigma, P6148) in PBST. 25 MESAB (as in Method 2). 30% Sucrose/PBST solution. Add 0.05% (w/v) of Sodium Azide and store at 4 C. Frozen Section Medium NEG50: Richard-Allan Scientific, 6502. 3 mL dispensable transfer polyethylene pipettes: e.g., BD Falcon, 357524 Dissecting needle. Superfrost plus microscope slides precleaned: Fisher, 12-550-15. The choice of slides is essential to avoid losing sections during washes. Grease pen: Electron Microscopy Sciences, 71319. Empty tip Boxes with the perforated holding surface: e.g., USA Scientific, 11111800. Blocking solution: PBS + 0.5% Triton X-100 + 10% serum. Use serum that matches the species in which secondary antibodies were generated. VectaShield Mounting Media: Vector Labs, H-1000. Optional: antigen retrieval solution: 100 mM Sodium Citrate in deionized H2O. Optional: Alexa Fluor 546 Phalloidin: Invitrogen, A22283. Prepare stock solution in methanol according to manufacturer’s instructions.

Fixing, embedding and sectioning 1. To visualize outer segments, pigmentation needs to be eliminated. To block pigment formation, raise embryos in the presence of PTU starting at 24 hpf or earlier (Method 2, Step 2) 2. When embryos/larvae reach the desired stage, sacrifice them by over anesthetization in MESAB, and immediately transfer to fixative. 3. Fix embryos for 2 h at RT or overnight at 4 C. Use an orbital shaker. Subsequent steps are at RT unless stated otherwise. 4. Remove fix and wash three times in PBST, 5 min each. 5. Cryoprotect in sucrose solution overnight at 4 C, or at RT until specimens sink. 6. Replace sucrose with Frozen Section Medium. This is a very viscous substance. Use Pasteur pipette. 7. Cut polyethylene pipette perpendicular to its long axis into 6–10 mm wide rings. Place the rings on a glass slide and fill them halfway with Frozen Section Medium. 8. Using Pasteur pipette transfer specimen into the ring ‘‘mold’’ described in the previous step. Some Frozen Section Medium will transfer too, which will fill up the mold completely.


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9. Use dissecting needle to orient specimen. To obtain transverse sections, heads of specimen must point straight down. Cool slides with specimen molds to -20 C in the cryostat chamber. Medium will solidify and turn white in about 5–10 min. 10. Prior to sectioning, remove polyethylene molds by cutting through their walls with a razorblade. 11. Mount frozen blocks on a cryostat specimen holder using additional Frozen Section Medium and collect 12–30 mm thick sections on a prechilled slide inside the cryostat chamber set to 20 C. When brought to RT, sections will thaw and adhere to the slide. 12. Allow slides to dry at RT for 4 h. Then proceed to the next step.

Immunohistochemistry 1. Optional Step: Detection of some antigens may require special treatment such as placing slides into boiling-hot 10% sodium citrate solution for 10 min (this procedure is known as antigen retrieval). 2. Using a grease pen draw a hydrophobic border around at the edge of the slide. Allow the grease to dry for 2–3 min. 3. Briefly rehydrate slides in PBST. This will also dissolve the Frozen Section Medium. Make sure to be gentle, as it is easy to lose sections during this step. 4. In the meantime prepare humidified chamber. For this purpose, we use a tip rack with a cover (see Materials section above for product information). Fill the bottom compartment of the tip rack with tap water. This will serve as a homemade humidifying chamber for antibody incubations. 5. Open the tip rack cover. Place slides on the top of the perforated, tip holding surface. 6. Overlay the slide with 100–200 mL of the blocking solution. The grease border will keep the blocking solution on the slide. Close the rack cover. Incubate the slide with the blocking solution for 30–60 min at RT. 7. Dilute primary antibody(ies) to appropriate concentration(s) in blocking solution. 8. Remove the blocking solution by positioning slides vertically and collecting excess solution with a Kimwipe. Apply the primary antibody solution (as in Step 6). Incubate overnight at 4 C. 9. Next day wash slides three times in PBST, 5 min each. 10. Incubate with secondary antibody(ies) at RT for 2 h. Optional: Add phalloidin to counterstain F-actin (dilute stock solution 1:40). 11. Wash slides several times in PBST. 12. Use Kimwipe to remove excess PBST and apply mounting medium. Place a coverslip and seal with nail polish. 13. Image using confocal or conventional fluorescence microsocopy. 14. The preparations may be stored at 4 C for at least a week, although the quality of signal deteriorates over time.

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Auditory System Mutations of zebrafish IFT genes, and presumably also other cilia-related loci, lead to the loss of kinocilia and subsequently the degeneration of mechanosensory hair cells (Tsujikawa and Malicki, 2004). This can be monitored using similar approaches to those described for photoreceptor cells. The overall health of hair cells is easy to evaluate on transverse sections of maculae. On plastic sections, maculae appear as thickenings of the epithelium that lines the otic vesicle (Fig. 3Q). The maculae contain two major cell types: the supporting cells and hair cells. The latter are oval in shape and positioned next to the apical surface (Bang et al., 2001; Schibler and Malicki, 2007). To evaluate the ear by simple histology on plastic sections, we use the protocol described in Method 6. Prior to embedding, otoliths have to be removed as described in Method 8 below. Anti-acetylated tubulin antibodies stain hair cell bodies in addition to their kinocilia (Fig. 1 D, E, G). The signal is particularly prominent at the apical surface of the cell. Therefore, these antibodies can also be used to evaluate hair cell morphology. In addition, the HCS-1 antibody, which has recently been shown to recognize otoferlin, is a very good marker of hair cell somata in zebrafish (Gale et al., 2002; Goodyear et al., 2010; Schibler and Malicki, 2007). Immunohistochemistry can be conveniently performed on whole animals through at least 5 dpf either alone or in combination with phalloidin staining. Phalloidin is commonly used to stain filamentous actin-rich structures. As hair cell sterocilia are supported by dense actin bundles, phalloidin is very useful to visualize the morphology of stereociliary bundles. Double staining with phalloidin and anti-tubulin antibodies makes it possible to evaluate the planar polarity of hair cells. This is done by comparing the relative positions of the kinocilium and the stereociliary bundle. The directionality of each hair cell can be visualized on en face images of maculae by drawing a line connecting the kinocilium and the center of the stereociliary bundle. In a polarized epithelium, all lines will have the same direction. As in the case of plastic sections, additional steps must be taken to remove otoliths in order to properly visualize hair cells (see Method 8 below). In addition to whole-mount staining, auditory cells and their features can be analyzed on frozen sections. The use of frozen sections is a necessity for reagents that produce high background in whole-mount applications, such as anti-g tubulin antibodies. Hair cells can also be visualized with a brn3c::GFP reporter transgene (Xiao et al., 2005). This transgenic line produces a robust signal in whole animals, which obviates the need to the section. Below we describe a method for staining of hair cells in whole embryos. To analyze the staining of hair cells on frozen sections, follow the protocol described for photoreceptors (Method 7). I. Method 8: Whole-Mount Staining and Imaging of Hair Cells in the Inner Ear

1. Materials

Mouse anti-acetylated tubulin or mouse anti-HCS-1 (Table I). PBST (PBS + 0.1% Tween-20).


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25 MESAB (as in Method 2). Fixative: 4% PFA (Sigma, P6148) in PBST. PBST + 1% Triton X-100. Blocking solution: PBS + 0.5% Triton X-100 + 10% serum. Use serum that matches the species in which secondary antibodies were generated. Water dipping objectives: e.g., Leica 40 dipping lens, working dist. 3.3 mm, NA 0.8; or Leica 63 dipping lens, working dist. 2.2 mm, NA 0.9. Optional: Alexa Fluor 546 Phalloidin: Invitrogen, A22283.

2. Additional Materials Agarose embedding supplies

Embryo medium (as in Method 5). Low melting point-agarose (Sigma, A9414-10G), 1% in embryo medium (keep liquid on 50 C heat block). Regular agarose (Invitrogen, 16500–500), 1% in embryo medium. Custom made molds to imprint the surface of agarose. Blastomere transplantation molds can be used here. They produce wells with the square, 1 1 mm opening, and 30 degree tilted bottom. Molds can be made by local machine shops or purchased from Adaptive Science Tools. Large (90 mm) and small (60 mm) disposable petri dishes. Dissecting needle. Razor blade.

3. Protocol Fixing and staining 1. Sacrifice embryos at desired stage by over anesthetization in MESAB, and immediately transfer to fixative. 2. Fix embryos or larvae for 4 h at RT or overnight at 4 C in freshly prepared fixative on an orbital shaker. Subsequent steps are at RT on the orbital shaker unless stated otherwise. 3. Wash once in PBST for 5 min. 4. Incubate in 1% Triton X-100 in PBST for 8 h or longer. This treatment dissolves otoliths. Alternatively, otoliths can be dissolved by incubating larvae in 120 mM EDTA in 0.1 M phosphate buffer (up to 3 d for dissected adult ears). 5. Wash once in PBST for 5 min. 6. Incubate in the blocking solution for 30 min. 7. Incubate specimen overnight at 4 C in the blocking solution containing appropriate concentration of the primary antibody (Table I). 8. Wash four times in PBST, 30 min each.

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9. Incubate specimen overnight at 4 C in the blocking solution containing appropriate concentration of the secondary antibody. Optional: Add phalloidin to counterstain F-actin (dilute stock solution 1:40). 10. Wash three times in PBST, 30 min each. Wash once in PBS.

Embedding in agarose for imaging 1. Pour 1% regular agarose into a large petri dish and place a mold on a top of it. The mold will float on agarose surface. Let the agarose harden completely. 2. Removing the mold from the agarose will leave imprinted wells on the agarose. Using a razor blade, cut and gently retrieve the agarose block that contains the wells. Place it into a 60 mm petri dish in the orientation that exposes well openings. 3. To properly orient the embryo, place it on its lateral side over the well opening so that the yolk mass fits inside the well. This will position the embryo flat on the agarose surface. 4. Once embryos are positioned properly, immobilize them using 1% low melting point agarose. If necessary, adjust embryo orientation before the agarose hardens. 5. Flood embryos in agarose with egg water and view using a water-dipping objective. Embryos can be stored for several days prior to imaging, although the quality of signal will gradually deteriorate. 6. This procedure can be also used in the analysis of living specimen. For this application, lower heat block temperature to 40 C or less. J. Method 9: Detection of Basal Bodies in Hair Cells

1. Materials

Mouse anti-g tubulin: Sigma, GTU88, T6557 PBST: PBS + 0.1% Tween-20. Fixative: 4% PFA (Sigma, P6148) in PBST. 25 MESAB (as in Method 2) PBD: PBST + 1% v/v DMSO + 1% w/v BSA. Serum that matches the species in which secondary antibodies were generated. 0.25% Trypsin (Fisher, T-360) in PBS. Make fresh each time. Agarose embedding supplies (as in Method 8). Optional – Alexa Fluor 546 Phalloidin: Invitrogen, A22283. Prepare stock solution in methanol according to manufacturer’s instructions.

2. Protocol 1. Sacrifice embryos at desired stage by anesthetization in MESAB, and immediately transfer to fixative. 2. Fix embryos in fresh fixative for 2 h at RT or overnight at 4 C. Use the orbital shaker. Subsequent steps at RT unless stated otherwise. 3. Wash twice in PBST, 5 min each.


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4. Permeabilize using 0.25% Trypsin in PBS (15 min for embryos at 30 hpf, 30 min at 4 dpf). 5. Wash twice in PBST, 5 min each. 6. Incubate in PBST containing 0.2% Triton X-100 for 30 min. 7. Wash twice in PBST, 5 min each. 8. Block for 30 min by incubating in PBD containing 10% serum. 9. Incubate in the primary antibody diluted in PBD (1:300) overnight at 4 C. 10. Wash four times in PBD, 30 min each. 11. Incubate in the secondary antibody (diluted 1:300 in PBD) overnight at 4 C. Optional: Add phalloidin to counterstain F-actin (dilute stock solution 1:40). 12. Wash three times in PBD, 30 min each. 13. Wash once in PBS to remove detergent. 14. Proceed to agarose mounting (see Method 8)

Lateral Line Neuromast hair cells can be labeled with the same reagents as used to stain cilia in the ear. Since neuromasts are protruding from the surface of the skin, sectioning of the tissue may not be necessary. Some protocols, such as anti-g -tubulin staining described above, work very well for the lateral line. Neuromasts are easily identified using the vital dye DASPEI, which stains hair cells. We describe a DASPEI staining method below. K. Method 10: Detection of Neuromast Hair Cells in a Living Specimen

1. Materials

DASPEI: Invitrogen, D426. E3 medium: 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4. MESAB (as in Method 2). Methylcellulose (as in Method 2). Agarose mounting implements (as in Method 8).

2. Protocol 1. 2. 3. 4. 5.

Transfer larvae to 1 mM DASPEI solution in E3 medium. incubate for 20 min. rinse thoroughly in E3. anaesthetize in MESAB, and mount in methylcellulose or agarose. view using fluorescence microscopy.

Olfactory System As olfactory cilia are located on the surface of the animal, they are easily accessible to staining reagents. The length of cilia can be evaluated by antibody staining

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and confocal microscopy in whole animals (Omori et al., 2008). Phalloidin is useful as a counterstain in such experiments to highlight the appearance of the surrounding tissues. Similar to photoreceptors and mechanosensory hair cells, olfactory sensory neurons degenerate following the loss of cilia in IFT mutants (Tsujikawa and Malicki, 2004). The survival of these neurons can be monitored using in situ hybridization with probes to the olfactory marker protein gene or via DiI labeling (Dynes and Ngai, 1998; Hansen and Zeiske, 1993; Tsujikawa and Malicki, 2004). Below, we describe DiI labeling.

L. Method 11: Labeling of Olfactory Neurons by DiI Incorporation

1. Materials

Neuro Trace DiI Tissue Labeling Paste: Invitrogen, N22880. DMSO: Sigma, D5879. Ethanol: Sigma, 279741-1L. Embryo medium (recipe as in Method 5). Incubator set at 30 C. Regular agarose, 1% (as in Method 8).

2. Protocol 1. Prepare DiI/DMSO stock by mixing ca. 60 mg of DiI paste (one scoop with a 200 mL tip) in 500–1000 mL of DMSO. 2. Prepare embryo medium containing 7% DiI/DMSO stock and 1% ethanol. Warm up to 30 C. 3. Incubate live embryos in the solution from Step 2 at 30 C for 10 min. 4. Wash embryos in embryo medium at 28 C. Embryos will not look healthy but will remain alive. 5. To observe olfactory pits, larvae can be positioned vertically, heads up, in agarose wells. In this case, wells can be made by punching holes in an agarose bed with a Pasteur pipette. Mounted embryos can be imaged by confocal or conventional fluorescence microscopy.

V. Future Directions The importance of cilia function in the KV, the kidney duct, and sensory organs is now well appreciated. However, cilia not restricted to these organs and many basic questions remain unanswered. For example, when and where do cilia first start to form in the context of organogenesis? What are the distribution patterns of motile and immotile cilia in different organs during development? What is the normal frequency and waveform of cilia movement in different tissues? Are the morphology, length, and the motility of cilia regulated by physiological parameters? Furthermore,


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although IFT trafficking and calcium influx are intimately associated with cilia formation and function, they are still challenging to image in live embryos. This is an important technical issue that merits further attention. With the development of more advanced imaging tools, we will gain a deeper and more comprehensive understanding of the function of this miniature cell surface organelle in development and adult physiology. Acknowledgments We thank Dr. Chengtian Zhao, and Peter Kovach of the Malicki lab, as well as members of Yale Center for PKD research for helpful discussions, Nicole Semanchik for superb technical assistance and SueAnn Mentone for assistance on histology. This work was supported by NIDDK (RO1 DK069528 and P50 DK057328 (project #3) awards (to ZS), and NEI research grant RO1 EY018176 (to JM).

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CHAPTER 4

Cellular Dissection of Zebrafish Hematopoiesis David L. Stachura*,y and David Traver*,y *

Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego, La Jolla, California, USA y

Department of Cellular and Molecular Medicine, University of California San Diego School of Medicine, La Jolla, California, USA

Abstract I. Introduction II. Zebrafish Hematopoiesis A. Primitive Hematopoiesis B. Definitive Hematopoiesis C. Adult Hematopoiesis III. Hematopoietic Cell Transplantation A. Embryonic Donor Cells B. Adult Donor Cells IV. Enrichment of Hematopoietic Stem Cells V. In vitro Culture and Differentiation of Hematopoietic Progenitors A. Stromal Cell Culture Assays B. Clonal Methylcellulose-based Assays VI. Conclusions References

Abstract The zebrafish is an excellent model system to study vertebrate blood cell development due to a highly conserved hematopoietic system, optical transparency, and amenability to both forward and reverse genetic approaches. The development of functional assays to analyze the biology of hematopoietic mutants and diseased animals remains a work in progress. Here we discuss recent advances in zebrafish hematology, prospective isolation techniques, cellular transplantation, and culturebased assays that now provide more rigorous tests of hematopoietic stem and METHODS IN CELL BIOLOGY, VOL 101 Copyright 2011. Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-387036-0.00004-9

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David L. Stachura and David Traver

progenitor cell function. Together with the proven strengths of the zebrafish, the development and refinement of these assays further enable efforts to better understand the development and evolution of the vertebrate hematopoietic system.

I. Introduction Over the past two decades, the development of forward genetic approaches in the zebrafish has provided unprecedented power in understanding the molecular basis of vertebrate blood development. Establishment of cellular and hematological approaches to better understand the biology of resulting blood mutants, however, has lagged behind these efforts. In this chapter, recent advances in zebrafish hematology will be reviewed, with an emphasis on prospective strategies for isolation of both embryonic and adult hematopoietic stem cells (HSCs) and the development of assays to rigorously test their function.

II. Zebrafish Hematopoiesis Developmental hematopoiesis, in both mammals and teleosts, occurs in four sequential waves (Fig. 1). The first two waves are termed ‘‘primitive,’’ and each generates transient precursors that respectively give rise to embryonic macrophages and erythrocytes (Keller et al., 1999; Palis et al., 1999). The next two waves consist of definitive hematopoietic precursors, defined as multipotent progenitors of adult cell types. The first to arise are erythromyeloid progenitors (EMPs), which give rise to erythroid and myeloid lineages (Bertrand et al., 2005b, 2007; Palis et al., 1999, 2001), followed by multipotent HSCs, which are endowed with the potential to both self-renew and generate all adult hematopoietic cell types [reviewed in Cumano and Godin (2007)]. A. Primitive Hematopoiesis Primitive hematopoiesis has been extensively studied in the mouse, where primitive macrophages and erythroid cells are generated in the extraembryonic yolk sac (YS) (Bertrand et al., 2005b; Palis et al., 1999). In the zebrafish, primitive macrophages develop in an anatomically distinct area known as the rostral blood island (RBI) (Fig. 1A). Transcripts for tal1 (also known as scl), lmo2, gata2a, and fli1a are found in the RBI between the three- and five-somite stages (ss) (Brown et al., 2000; Liao et al., 1998; Thompson et al., 1998). This is quickly followed by expression of spi1 (also known as pu.1) in a subset of these precursors (Bennett et al., 2001; Lieschke et al., 2002). Between 11 and 15 somites, spi1+ macrophages are detectable, and they migrate toward the head midline (Herbomel et al., 1999; Lieschke et al., 2002; Ward et al., 2003) and across the yolk ball (Fig. 1C). Some of these precursors enter circulation, while others migrate into the head (Herbomel et al.,

4. Cellular Dissection of Zebrafish Hematopoiesis

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Fig. 1

Model of hematopoietic ontogeny in the developing zebrafish embryo. (A) Different regions of lateral plate mesoderm give rise to anatomically distinct regions of blood cell precursors. Anatomical regions of embryo responsible for generation of hematopoietic precursors (red), vasculature (blue), and pre-HSCs (green) are highlighted. Cartoon is a five-somite stage embryo, dorsal view. (B) Timing of mouse and zebrafish hematopoietic development. In mouse (left), primitive hematopoiesis initiates in the yolk sac (YS; yellow), producing primitive erythroid cells and macrophages. Later, definitive EMPs emerge in the YS. HSCs are specified in the aorta, gonad, and mesonephros (AGM, teal) region. These HSCs eventually seed the fetal liver (orange), the main site of embryonic hematopoiesis. Adult hematopoiesis occurs in the thymus (blue), spleen (green), and bone marrow (red). Zebrafish hematopoiesis is similar: temporal analogy to mouse hematopoiesis shown in (B, right), spatial locations shown in (C). Numbers in (C) correspond to timing of distinct precursor waves. (C) Embryonic hematopoiesis occurs through four independent waves of precursor production. First, primitive macrophages arise in cephalic mesoderm, migrate onto the yolk ball, and spread throughout the embryo (purple, 1). Then, primitive erythrocytes develop in the intermediate cell mass (ICM; yellow, 2). The first definitive progenitors are EMPs, which develop in the posterior blood island (PBI; orange, 3). Later, HSCs arise in the AGM region (teal, 4), migrate to the CHT (later name for the PBI, orange), and eventually seed the thymus and kidney (blue; red). Similar hematopoietic events in mouse and fish are color-matched between right and left panels of (B). Hematopoietic sites (B) and locations (C) are also color matched. (hpf: hours postfertilization, dpf: days postfertilization, wpf: weeks postfertilization, E: embryonic day, RBI: rostral blood island.) (See Plate no. 6 in the Color Plate Section.)

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1999). By 28–32 h postfertilization (hpf), macrophages are found in circulation and dispersed throughout the embryo. Primitive erythroid cell generation begins in the murine YS blood islands at day 7.5 postcoitum (E7.5) (Fig. 1B) [reviewed in Palis et al. (2010)]. These blood islands consist of nucleated erythroid cells that express embryonic globin genes surrounded by endothelial cells. Although it was previously believed that mammalian primitive erythroid cells uniquely remained nucleated (similar to the nucleated erythrocytes of birds, fish, and amphibians), it is now accepted that mammalian primitive red blood cells do, in fact, enucleate into reticulocytes and prenocytes (Fraser et al., 2007; Kingsley et al., 2004; McGrath et al., 2008). The zebrafish has an equivalent anatomical site to mammalian blood islands, known as the intermediate cell mass (ICM), where two stripes of mesodermal cells expressing tal1, lmo2, and gata1a converge to the midline of the zebrafish embryo and are surrounded by endothelial cells that become the cardinal vein. (Al-Adhami and Kunz, 1977; Detrich et al., 1995) (Fig. 1A and B). Although the ICM is intraembryonic in zebrafish, it has a cellular architecture similar to the mammalian YS blood islands (Al-Adhami and Kunz, 1977; Willett et al., 1999). The development of transgenic zebrafish expressing fluorescent markers under the control of early mesodermal, prehematopoietic promoters (see Table I) now allow testing of primitive fate potentials by prospective isolation strategies and the functional assays outlined in this chapter.

B. Definitive Hematopoiesis Similar to primitive hematopoiesis, definitive hematopoiesis initiates through two distinct precursor subsets. In the mouse, multilineage hematopoiesis is first evident in the YS (Bertrand et al., 2005b; Palis et al., 1999, 2001; Yoder et al., 1997a, 1997b) and placenta (Gekas et al., 2005; Ottersbach and Dzierzak, 2005) by E9.5. Multilineage precursors in both tissues can be isolated and distinguished by the expression of CD41, an integrin molecule that labels early hematopoietic progenitors. CD41+ cells differentiate into both myeloid and erythroid lineages, but conspicuously lack lymphoid potential (Bertrand et al., 2005a; Yokota et al., 2006). These studies suggest that the definitive hematopoietic program in the developing mouse begins with committed EMPs. Recent studies in the zebrafish have demonstrated evolutionary conservation of EMPs as the first definitive precursor formed, and expanded upon findings in the mouse YS. EMPs can be isolated from the zebrafish posterior blood island (PBI) between 26 and 36 hpf (Fig. 1C) by their co-expression of fluorescent transgenes driven by the lmo2 and gata1a promoters (Fig. 2A) (Bertrand et al., 2007). In vitro differentiation experiments (Fig. 2B and C) and in vivo transplantation assays have shown these cells capable of only erythroid and myeloid differentiation (Bertrand et al., 2007). Studies performed in mindbomb mutant zebrafish lacking Notch signaling showed that EMP specification and differentiation are not affected by loss of the Notch pathway (Bertrand et al., 2010b),

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whereas HSCs are absent (Burns et al., 2005), further distinguishing these two definitive progenitors. The zebrafish allows the discrimination of EMPs from (HSCs, both of which have similar cell-surface markers and differ only in their differentiation and self-renewal potentials (Bertrand et al., 2005a,; 2005b). Unlike the murine system, these two progenitors arise in separate anatomical locations (Bertrand et al., 2007), and are therefore easily distinguishable. Importantly, fate-mapping studies in the zebrafish

Table I List of relevant transgenic zebrafish lines currently available for hematopoietic studies, indicating the promoter:gene expressed and the cell population(s) identified Transgene

Tissue

Transgene

Tissue

lmo2:GFP (Zhu et al., 2005) lmo2:mCherry*

Prehematopoietic, vasculature

kdrl:EGFP (Cross et al., 2003)


kdrl:DsRed (Jin et al., 2007b)

lmo2:DsRed (Lin et al., 2005) itga2b:GFP (Lin et al., 2005) itga2b:mCherry*


fli1a:EGFP (Lawson and Weinstein, 2002) fli1a:DsRed (Jin et al., 2007b)

EMPs, HSCs, thromobocytes

Pre-hematopoietic, vasculature Pre-hematopoietic, vasculature Pre-hematopoietic, vasculature Pre-hematopoietic, vasculature Kidney

itga2b:CFP*


ptprc:DsRed (Bertrand et al., 2008) ptprc:CFP* ptprc:AmCyan*

Pan-leukocyte

gata1a:GFP (Long et al., 1997) gata1a:DsRed (Traver et al., 2003b) cmyb:GFP (Bertrand et al., 2008) mpx:EGFP (Renshaw et al., 2006) gata2a:EGFP (Traver et al., 2003b) Ighm:EGFP* Ighz:EGFP*


gata3:AmCyan (Bertrand et al., 2008) rag2:EGFP (Langenau et al., 2003) lck:EGFP (Langenau et al., 2004)

Immature B and T cells Mature T cells

Red blood cells

il7r:mCherry* mhcII:GFP* (Wittamer et al., 2011) mhcII:AmCyan*

Red blood cells

lyz:EGFP (Hall et al., 2007)

Lymphoid precursors, T cells B cells, macrophages, dendritic cells B cells, macrophages, dendritic cells Neutrophils

HSCs, neural

lyz:DsRed (Hall et al., 2007)

Neutrophils

Neutrophils

runx1P1:GFP (Lam et al., 2008) EMPs

Eosinophils

runx1P2:GFP (Lam et al., 2008) HSCs

B cells B cells

ccr9a:cfp* Lymphoid precursors mpeg1:GAL4 (Ellett et al., 2010) Embryonic Macrophages

Pan-leukocyte Pan-leukocyte

This list is not comprehensive; other transgenic animals are being constantly generated, but these are a few of the essential tools currently being used in zebrafish hematopoiesis laboratories. *

Unpublished transgenic animals generated in the Traver laboratory.

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Fig. 2 Functional in vitro differentiation studies demonstrate that gata1+lmo2+ cells are committed erythromyeloid progenitors (EMP). (A) Purified EMP at 30 hpf (lmo2+gata1a+, black gate) have the immature morphology of early hematopoietic progenitors. As a comparison, purified primitive erythroblasts are shown (lmo2lowgata1a+, red gate). Magnification, 1000. (B) Short-term in vitro culture of lmo2+gata1a+ cells atop ZKS cells demonstrate erythroid (E), granulocytic (G), and monocytic/macrophage (M) differentiation potentials. Cultured cells were stained with May-Gr€ unwald/Giemsa and for myeloperoxidase (MPX) activity. lmo2lowgata1a+ cells only differentiated into erythroid cells (not shown).


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have demonstrated EMPs to arise from posterior mesodermal derivatives that express the lmo2 gene. This finding, in combination with lineage tracing studies demonstrating EMPs to completely lack T lymphoid potential, indicate that EMPs and HSCs are unique populations and independently derived during development. The final wave of hematopoiesis culminates with the formation of HSCs, which self-renew and give rise to all definitive blood cell lineages, including lymphocytes. It has been demonstrated that HSCs arise in an area of the mid-gestation mouse bounded by the aorta, gonads, and mesonephros (AGM) at E10–10.5 (Fig. 1B) (Cumano and Godin, 2007; Dzierzak, 2005). Many studies have also suggested that transplantable HSCs are present in the YS on E9 (Lux et al., 2008; Weissman et al., 1978; Yoder et al., 1997a, 1997b) and later in the placenta by E11 (Gekas et al., 2005; Ottersbach and Dzierzak, 2005). While these results suggest that HSCs may arise in distinctly different locations in the developing mouse embryo, it is now clear that HSCs originate from arterial endothelium. Recent studies in the E10.5 mouse (Boisset et al., 2010) and 36–52 hpf zebrafish embryo (Bertrand et al., 2010a; Kissa and Herbomel, 2010) demonstrated directly the birth of HSCs from aortic endothelium via confocal imaging. A commonality among all vertebrate embryos thus seems to be the generation of HSCs from hemogenic endothelium lining the aortic floor (Ciau-Uitz et al., 2000; de Bruijn et al., 2002; Jaffredo et al., 1998; North et al., 2002; Oberlin et al., 2002). Similar studies will need to be performed to determine whether or not additional embryonic sites can likewise generate HSCs autonomously, including the YS and placenta. In all locations, however, it is clear that HSCs are present only transiently; by E11, the fetal liver (FL) is populated by circulating HSCs (Houssaint, 1981; Johnson and Moore, 1975) and becomes the predominant site of blood production during midgestation, producing the first full complement of definitive, adult-type effector cells. Shortly afterward, hematopoiesis is evident in the fetal spleen, and occurs in bone marrow throughout adulthood (Keller et al., 1999). The zebrafish possesses an anatomical site that closely resembles the mammalian AGM (Fig. 1B and C). Between the dorsal aorta and cardinal vein between 28 and 48 hpf, cmyb+ and runx1+ blood cells appear in intimate contact with the dorsal aorta (Burns et al., 2002; Kalev-Zylinska et al., 2002; Thompson et al., 1998). Lineage tracing of CD41+ HSCs derived from this ventral aortic region show their ability to colonize the thymus (Bertrand et al., 2007; Kissa et al., 2008) and pronephros (Bertrand et al., 2008; Murayama et al., 2006), which are the sites of adult hematopoiesis (Jin et al., 2007a; Murayama et al., 2006). After 48 hpf, blood production appears to shift to the caudal hematopoietic tissue (CHT) (Fig. 1C), and later to the pronephros, which serves as the definitive hematopoietic organ for the remainder of life. The development of transgenic zebrafish expressing fluorescent markers under the control of definitive hematopoietic promoters such as itga2b (also known as cd41), cmyb, and runx1 (see Table I) now allows testing of fate potentials by prospective isolation strategies and functional assays outlined later in this chapter.

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C. Adult Hematopoiesis Previous genetic screens in zebrafish were successful in identifying mutants that affected primitive erythropoiesis. The screening criteria used in these screens scored visual defects in circulating blood cells during early embryogenesis; mutants defective in definitive hematopoiesis but displaying normal primitive blood cell development were therefore likely missed. Current screens aimed at identifying mutants with defects in the generation of definitive HSCs in the AGM should reveal new genetic pathways required for multilineage hematopoiesis. Recent studies in zebrafish show that nearly all adult hematopoietic cells derive from HSCs born from aortic endothelium (Bertrand et al., 2010a), consistent with findings in the murine system (Chen et al., 2009; Zovein et al., 2008). Therefore, mutational screens designed to identify defects in hemogenic endothelium may yield information about the full repertoire of hematopoietic regulation, specification, maintenance, and differentiation over the organism’s lifespan. Understanding the biology of mutants isolated using these approaches, however, first requires the characterization of normal, definitive hematopoiesis and the development of assays to study the biology of zebrafish blood cells more precisely. To this end, we have established several tools to characterize the definitive blood-forming system of adult zebrafish. Blood production in adult zebrafish, like other teleosts, occurs in the kidney, which supports both renal functions and multilineage hematopoiesis (Zapata, 1979). Similar to mammals, T lymphocytes develop in the thymus (Trede and Zon, 1998; Willett et al., 1999) (Fig. 3A), which exists in two bilateral sites in zebrafish (Hansen and Zapata, 1998; Willett et al., 1997). The teleostean kidney is a sheath of tissue that runs along the spine (Fig. 3B, E); the anterior portion, or head kidney, shows a higher ratio of blood cells to renal tubules than does the posterior portion (Zapata, 1979), termed the trunk kidney (Fig. 3B, C). All mature blood cell types are found in the kidney and morphologically resemble their mammalian counterparts (Fig. 3G, Fig. 4), with the exceptions that erythrocytes remain nucleated and thrombocytes perform the clotting functions of platelets (Jagadeeswaran et al., 1999). Histologically, the zebrafish spleen (Fig. 3D) has a simpler structure than its mammalian counterpart in that germinal centers have not been observed (Zapata and Amemiya, 2000). The absence of immature precursors in the spleen, or any other adult tissue, suggests that the kidney is the predominant hematopoietic site in adult zebrafish. The cellular compositions of whole kidney marrow (WKM), spleen, and blood are shown in Fig. 3F-H. Morphological examples of all kidney cell types are presented in Fig. 4. Analysis of WKM by fluorescence activated cell sorting (FACS) showed that several distinct populations could be resolved by light scatter characteristics (Fig. 5A). Forward scatter (FSC) is directly proportional to cell size, and side scatter (SSC) proportional to cellular granularity (Shapiro, 2002). Using combined scatter profiles, the major blood lineages can be isolated to purity from WKM following two rounds of cell sorting (Traver et al., 2003b). Mature erythroid cells were found exclusively within two FSClow fractions (Populations R1 and R2,


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Fig. 3

Histological analyses of adult hematopoietic sites. (A) Sagittal section showing location of the thymus (T), which is dorsal to the gills (G). (B) Midline sagittal section showing location of the kidney, which is divided into the head kidney (HK), and trunk kidney (TK), and spleen (S). The head kidney shows a higher ratio of blood cells to renal tubules (black arrows), as shown in a close up view of the HK in (C). (D) Close up view of the spleen, which is positioned between the liver (L) and the intestine (I). (E) Light microscopic view of the kidney (K), over which passes the dorsal aorta (DA, white arrow). (F) Cytospin preparation of splenic cells, showing erythrocytes (E), lymphocytes (L), and an eosinophil (Eo). (G) Cytospin preparation of kidney cells showing cell types as noted above plus neutrophils (N) and erythroid precursors (O, orthochromic erythroblast). (H) Peripheral blood smear showing occasional lymphocytes and thrombocytes (T) clusters amongst mature erythrocytes. (A–D) Hematoxylin and eosin stains, (F–H) May-Gr€ unwald/Giemsa stains.

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Fig. 4 Proposed model of zebrafish definitive hematopoietic differentiation. Isolated, cytospun, and stained blood cells from the zebrafish kidney, thymus, and peripheral tissues and their proposed upstream progenitors. Proposed lineage relationships are based on those demonstrated in clonogenic murine studies. Multipotent and lineage restricted progenitors likely reside in the kidney marrow, but their existence has never been experimentally proven due to a paucity of in vitro assays.

Fig. 5A, D), lymphoid cells within a FSCint SSClow subset (Population R3, Fig. 5A, E), immature precursors within a FSChigh SSCint subset (Population R4, Fig. 5A, F), and myelomonocytic cells within only a FSChigh SSChigh population (Population R5, Fig. 5A, G). Interestingly, two distinct populations of mature erythroid cells exist (Fig. 5A, R1, R2 gates). Attempts at sorting either of these subsets reproducibly resulted in approximately equal recovery of both (Fig. 5D). This likely resulted due to the elliptical nature of zebrafish red blood cells, because sorting of all other populations yielded cells that fell within the original sorting gates upon re-analysis. Examination of splenic (Fig. 5B) and peripheral blood (Fig. 5C) suspensions showed each to have distinct profiles from WKM, each being predominantly erythroid. It should be noted that, due to differences in the fluidics and beam size, erythroid cells are not discretely detectable on BD FACScan, FACS Caliber, or FACS Aria I and II flow cytometers. However, FACS Vantage and LSR-II flow cytometers have a different fluidics system and are well suited to these analyses. Sorting of each scatter population from spleen and blood showed each to contain only erythrocytes, lymphocytes, or myelomonocytes in a manner identical to those in the kidney. Immature precursors were not observed in either tissue. Percentages of cells within each scatter population closely matched those obtained by morphological cell counts,


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Fig. 5 Each major blood lineage can be isolated by size and granularity using FACS. (A) Scatter profile for WKM. Mature erythrocytes are found within R1 and R2 gates, lymphocytes within the R3 gate, immature precursors within the R4 gate, and myeloid cells within the R5 gate. Mean percentages of each population within WKM are shown. Scatter profiling can also be utilized for analyzing spleen (B) and peripheral blood (C). Purification of each WKM fraction by FACS (D–G). (D) Sorting of populations R1 or R2 yields both upon re-analysis. This appears to be due to the elliptical shape of erythrocytes (right panel). (E) Isolation of lymphoid cells. (F) Isolation of precursor fraction. (G) Isolation of myeloid cells. FACS profiles following one round of sorting are shown in left panels, after two rounds in middle panels, and morphology of double-sorted cells shown in right panels (E–G).

demonstrating that this flow cytometric assay is accurate in measuring the relative percentages of each of the major blood lineages. Many transgenic zebrafish lines have been created using proximal promoter elements from genes that demonstrate lineage-affiliated expression patterns in the mouse. These include gata1a:GFP (Long et al., 1997), gata2a:EGFP (Jessen et al., 1998; Traver et al., 2003b), rag2:EGFP (Langenau et al., 2003), lck:EGFP (Langenau et al., 2004), spi1:EGFP (Hsu et al., 2004; Ward et al., 2003), and itga2b:EGFP (Lin et al., 2005; Traver et al., 2003b) stable transgenic lines. In the adult kidney, we have demonstrated that each of these animals expresses green

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fluorescent protein (GFP) in the expected kidney scatter fractions (Traver et al., 2003b). For example, all mature erythrocytes express GFP in gata1a:GFP transgenic animals, as do erythroid progenitors within the precursor population. High expression levels of Gata2 are seen only within eosinophils, Rag2 and Lck only within cells in the lymphoid fraction, and Spi1 in both myeloid cells and rare lymphoid cells. The development of itga2b:EGFP transgenic animals has demonstrated that rare thrombocytic cells are found within the kidney, with thrombocyte precursors appearing in the precursor scatter fraction and mature thrombocytes in the lymphoid fraction. Without fluorescent reporter genes, rare populations such as thrombocytes cannot be resolved by light scatter characteristics alone. By combining the simple technique of scatter separation with fluorescent transgenesis, specific hematopoietic cell subpopulations can now be isolated to a relatively high degree of purity for further analyses. FACS profiling can also serve as a diagnostic tool in the examination of zebrafish blood mutants. The majority of blood mutants identified to date are those displaying defects in embryonic erythrocyte production (Traver et al., 2003a). Most of these mutants are recessive and many are embryonic lethal when homozygous. Most have not been examined for subtle defects as heterozygotes. Several heterozygous mutants such as retsina, riesling, and merlot showed haploinsufficiency as evidenced by aberrant kidney erythropoiesis (Traver et al., 2003b). All mutants displayed anemia with concomitant increases in erythroid precursors. These findings suggest that many of the gene functions required to make embryonic erythrocytes are similarly required in their adult counterparts at full gene dosage for normal function.

III. Hematopoietic Cell Transplantation In mammals, cellular transplantation has been used extensively to functionally test putative hematopoietic stem and progenitor cell populations, precursor/progeny relationships, and cell autonomy of mutant gene function. To address similar issues in zebrafish, several different varieties of hematopoietic cell transplantation (HCT) have been developed (Fig. 6). A. Embryonic Donor Cells Although scatter profiling has proven very useful in analyzing and isolating specific blood lineages from the adult kidney, it cannot be used to enrich blood cells from the developing embryo. To study the biology of the earliest blood-forming cells in the embryo, we have made use of transgenic zebrafish expressing fluorescent proteins. As discussed above, hematopoietic precursors appear to be specified from mesodermal derivatives that express lmo2, kdrl (also known as flk1), and gata2a. The proximal promoter elements from each of these genes have been shown to be sufficient to recapitulate their endogenous expression patterns. Using germline transgenic animals expressing GFP under the control of each of these promoters,


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Fig. 6

Methods of hematopoietic cell transplantation in the zebrafish. See the text for experimental

details.

blood cell precursors can be isolated by flow cytometry from embryonic and larval animals for transplantation into wild-type recipients. For example, GFP+ cells in lmo2:EGFP embryos can be visualized by FACS by 8–10 somites (Traver, 2004). These cells can be sorted to purity and tested for functional potential in a variety of transplantation (Fig. 6) or in vitro culture assays (see Fig. 2). We have used two types of heterochronic transplantation strategies to address two fundamental questions in developmental hematopoiesis. The first is whether cells that express Lmo2 at 8–12 somites have hemangioblastic potential, i.e., can generate both blood and vascular cells. We reasoned that purified cells should be placed into a relatively naive environment to provide the most permissive conditions to assess their full fate potentials. Therefore, we attempted transplantation into 1000 cell stage blastulae recipients. Transplanted cells appear to survive this procedure well and GFP+ cells could be found over several days later in developing embryos and larvae. By isolating GFP+ cells from lmo2:EGFP animals also carrying a gata1a:DsRed transgene, both donor-derived endothelial and erythroid cells can be independently visualized in green and red, respectively. Using this approach we have shown that Lmo2+ cells from 8 to 12 ss embryos can generate robust regions of donor

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endothelium and intermediate levels of circulating erythrocytes (D. Traver, C. E. Burns, H. Zhu, and L. I. Zon, unpublished results). We are currently generating additional transgenic lines that express DsRed or mCherry under ubiquitous promoters to test the full fate potentials of Lmo2+ cells upon transplantation. Additionally, although these studies demonstrate that Lmo2+ cells can generate at least blood and endothelial cells at the population level, single-cell fate-mapping studies need to be performed to assess whether clonogenic hemangioblasts can be identified in vivo. The second question addressed through transplantation is whether the earliest identifiable primitive blood precursors can generate the definitive hematopoietic cells that arise later in embryogenesis. It has been previously reported that the embryonic lethal vlad tepes mutant dies from erythropoietic failure due to a defect in the gata1a gene (Lyons et al., 2002). This lethality can be rescued by transplantation of WKM from wild-type adults into mutant recipients at 48 hpf (Traver et al., 2003b). We therefore tested whether cells isolated from 8 to 12ss lmo2:EGFP embryos could give rise to definitive cell types and rescue embryonic lethality in vlad tepes recipients. Following transplantation of GFP+ cells at 48 hpf, approximately half of the cells in circulation were GFP+ and the other half were DsRed+ 1 day posttransplantation. Three days later, analyses of the same animals showed that the vast majority of cells in circulation were DsRed+, apparently due to the differentiation of Lmo2+ precursors to the erythroid fates. Compared to untransplanted control animals which all died by 12 days postfertilization (dpf), some mutant recipients survived for 1–2 months following transplantation. We observed no proliferation of donor cells at any time point following transplantation, however, and survivors analyzed over 1 month posttransplantation showed no remaining cells in circulation (D. Traver, C. E. Burns, H. Zhu, and L. I. Zon, unpublished results). Therefore, these data indicate that mutant survivors were only transiently rescued by short-lived, donor-derived erythrocytes. Thus, within the context of this transplantation setting, it does not appear that Lmo2 + hematopoietic precursors can seed definitive hematopoietic organs to give rise to enduring repopulation of the host blood forming system.

1. Protocol for Isolating Hematopoietic Cells from Embryos This simple physical dissociation procedure is effective in producing single cell suspensions from early embryos (8–12 ss) as well as from embryos as late as 48 hpf. 1. Stage and collect embryos. We estimate that approximately 200 cells can be isolated per 10–12 ss lmo2:EGFP embryo. It is recommended that as many embryos are collected as possible since subsequent transplantation efficiency depends largely upon cell concentration. At least 500–1000 embryos are recommended. 2. Transfer embryos into 1.5 ml eppendorf centrifuge tubes. Add embryos until they sediment to the 0.5 ml mark. Remove embryo medium since it is not optimal for cellular viability.


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3. Wash 2X with 0.9X Dulbecco’s PBS (Gibco; 500 ml 1X Dulbecco’s PBS + 55 ml ddH2O). 4. Remove 0.9X PBS and add 750 ml ice-cold staining medium (SM; 0.9X Dulbecco’s PBS + 5% FCS). Keep cells on ice from this point onward. 5. Homogenize with blue plastic pestle and pipette a few times with a p1000 tip. 6. Strain resulting cellular slurry through a 40 mm nylon cell strainer (Falcon 2340) atop a 50 ml conical tube. Rinse with additional SM to flush cells through the filter. 7. Gently mash remaining debris atop strainer with a plunger removed from a 28gauge syringe. 8. Rinse with more SM until the conical tube is filled to the 25 ml mark (this helps to remove the yolk). 9. Centrifuge for 5 min @ 200g at 4 C. Remove supernatant until 1–2 ml remain. 10. Add 2–3 ml SM; resuspend by pipetting. 11. Strain again through 40 mm nylon mesh into a 5 ml Falcon 2054 tube. It is important to filter the cell suspension at least twice before running the sample by FACS. Embryonic cells are sticky and will clog the nozzle if clumps are not properly removed beforehand. 12. Centrifuge again for 5 min @ 200g at 4 C. Repeat steps 10–12 if necessary. 13. Remove supernatant and resuspend with 1–2 ml SM depending upon number of embryos used. 14. Propidium iodide (PI) may be added at this point to 1 mg/ml to exclude dead cells and debris on the flow cytometer. When using, however, bring samples having PI only and GFP only to set compensations properly. Otherwise, the signal from PI may bleed into the GFP channel resulting in false positives. Alternatively, add 1:1000 Sytox Red (excited by 633 nm laser), or Sytox Blue (excited by the 405 nm laser) for dead cell discrimination, as they have no spectral overlap with GFP. Embryonic cells are now ready for analysis or sorting by flow cytometry. It is often difficult to visualize GFP+ cells when the expression is low or the target population is rare, so one should always prepare age-matched GFP-negative embryos in parallel with transgenic embryos. It is then apparent where the sorting gates should be drawn to sort bona fide GFP+ cells. If highly purified cells are desired, one must perform two successive rounds of sorting. In general, sorting GFP+ cells once yields populations of approximately 50–70% purity. Two rounds of cell sorting generally yields >90% purity as observed with 10ss lmo2:EGFP cells (Traver, 2004). Cells should be kept ice-cold during the sorting procedure.

2. Transplanting Purified Cells into Embryonic Recipients After sorting, centrifuge cells for 5 min at 200g and 4 C. Carefully remove all supernatant. Resuspend cell pellet in 5–10 ml of ice-cold SM containing 3U heparin and 1U DnaseI to prevent coagulation and lessen aggregation. Preventing the cells

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from aggregating or adhering to the glass capillary needle used for transplantation is critical. Mix the cells by gently pipetting with a 10 ml pipette tip. Keep on ice. For transplantation, we use the same needle-pulling parameters used to make needles for nucleic acid injections, the only difference being the use of filament-free capillaries to maintain cell viability. We also use the standard air-powered injection stations used for nucleic acid injections.

3. Transplanting Cells into Blastula Recipients 1. Stage embryonic recipients to reach the 500–1000 cell stage at the time of transplantation. 2. Prepare plates for transplantation by pouring a thin layer of 2% agarose made in E3 embryo medium into a 6 cm Petri dish. Drop transplantation mold [similar to the embryo injection mold described in Chapter 5 of The Zebrafish Book (Westerfield, 2000) but having individual depressions rather than troughs] atop molten agarose and let solidify. 3. Dechorionate blastulae in 1–2% agarose-coated Petri dishes by light pronase treatment or manually with watchmaker’s forceps. 4. Transfer individual blastulae into individual wells of transplantation plate that has been immersed in 1X HBSS (Gibco). Position the animal pole upward. 5. Using glass, filament-free, fine-pulled capillary needles (1.0 mm OD) backload 3–6 ml of cell suspension after breaking needle on a bevel to an opening of 20 mm. Load into needle holder and force cells to injection end by positive pressure using a pressurized air injection station. 6. Gently insert needle into the center of the embryo and expel cells using either very gentle pressure bursts or slight positive pressure. Transplanting cells near the marginal zone of the blastula leads to higher blood cell yields since embryonic fate maps show blood cells to derive from this region in later gastrula stage embryos. 7. Carefully transfer embryos to agarose-coated Petri dishes using glass transfer pipettes. 8. Place into E3 embryo medium and incubate at 28.5 C. Many embryos will not survive the transplantation procedure, so clean periodically to prevent microbial outgrowth. 9. Monitor by fluorescence microscopy for donor cell types.

4. Transplanting Cells into 48 hpf Embryos 1. All procedures are performed as above except that dechorionated 48 hpf embryos are staged and used as transplant recipients. 2. Fill transplantation plate with 1X HBSS containing 1X penicillin/streptomycin and 1X buffered tricaine, pH 7.0. Do not use E3, as it is suboptimal for cellular viability. Anesthetize recipients in tricaine and then array individual embryos into


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individual wells of transplantation plate. Position head at bottom of well, yolk side up. 3. Load cells as above. Insert injection needle into the sinus venosus/duct of Cuvier and gently expel cells by positive pressure or gentle pressure bursts. Take care not to rupture the YS membrane. A very limited volume can be injected into each recipient. It is thus important to use very concentrated cell suspensions in order to reconstitute the host blood-forming system. If using WKM as donor cells, concentrations of 5 105 cells/ml can be achieved if care is taken to filter and anticoagulate the sample. 4. Allow animals to recover at 28.5 C in E3. Keep clean and visualize daily by microscopy for the presence of donor-derived cells.

B. Adult Donor Cells Whereas the first HSCs transdifferentiate from embryonic aortic endothelium, multilineage hematopoiesis is not fully apparent until the kidney becomes the site of blood cell production. The kidney appears to be the only site of adult hematopoiesis, and we have previously demonstrated that it contains HSCs capable of the long-term repopulation of embryonic (Traver et al., 2003b) and adult (Langenau et al., 2004) recipients. For HSC-enrichment strategies, both high-dose transplants and limiting dilution assays are required to gauge the purity of input cell populations. In embryonic recipients, we estimate that the maximum number of cells that can be transplanted is approximately 5 103, and the precise quantitation of transplanted cell numbers is difficult. To circumvent both issues, we have developed HCT into adult recipients. For transplantation into adult recipients, myeloablation is necessary for successful engraftment of donor cells. We have found g-irradiation to be the most consistent way to deplete zebrafish hematopoietic cells. The minimum lethal dose (MLD) of 40 Gy specifically ablates cells of the blood-forming system and can be rescued by transplantation of one kidney equivalent (106 WKM cells). Thirty-day survival of transplanted recipients is approximately 75% (Traver et al., 2004). An irradiation dose of 20–25 Gy is sublethal, and the vast majority of animals survive this treatment despite having nearly total depletion of all leukocyte subsets 1 week following irradiation (Traver et al., 2004). We have shown that this dose is necessary and sufficient for transfer of a lethal T cell leukemia (Traver et al., 2004), and for longterm (>6 month) engraftment of thymus repopulating cells (Langenau et al., 2004). We do not yet know the average relative chimerism of donor to host cells when transplantation is performed following 20–25 Gy. That this dose is sufficient for robust engraftment, for long-term repopulation, and yields extremely high survival suggests that 20–25 Gy may be the optimal dose for myeloablative conditioning prior to transplantation. Improvement in short-term engraftment and long-term survival of transplant recipients will also likely require matching of MHC loci between donor and host genotypes.

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1. Protocols for Isolating Hematopoietic Cells from Adult Zebrafish Anesthetize adult animals in 0.02% tricaine in fish water. For blood collection, dry animal briefly on tissue then place on a flat surface with head to the left, dorsal side up. Coat a 10 ml pipette tip with heparin (3 U/ml) then insert tip just behind the pectoral fin and puncture the skin. Direct the tip into the heart cavity, puncture the heart, and aspirate up to 10 ml blood by gentle suction. Immediately perform blood smears or place into 0.9X PBS containing 5% FCS and 1 U/ml heparin. Mix immediately to prevent clotting. Blood from several animals may be pooled in this manner for later use by flow cytometry. Red cells may be removed using a red blood cell hypotonic lysis solution (Sigma; 8.3 g/l ammonium chloride in 0.01 M Tris–HCl, pH 7.5) on ice for 5 min. Add 10 volumes of ice-cold SM then centrifuge at 200g for 5 min at 4 C. Resuspended blood leukocytes can then be analyzed by flow cytometry or cytocentrifuge preparations. For collection of other hematopoietic tissues, place fish on ice for several minutes following tricaine. Make a ventral, midline incision using fine scissors under a dissection microscope. For spleen collection, locate spleen just dorsal to the major intestinal loops and tease out with watchmaker’s forceps. Place into ice cold SM. Dissect any nonsplenic tissue away and place on a 40 mm nylon cell strainer (Falcon 2340) atop a 50 ml conical tube. Gently mash the spleen using a plunger removed from a 28-gauge insulin syringe and rinse with SM to flush cells through the filter. Up to 10 spleens can be processed through each filter. Centrifuge at 200g for 5 min at 4 C. Filter again through 40 mm nylon mesh if using for FACS. For kidney collection, remove all internal organs using forceps and a dissection microscope. Take care during dissection because ruptured intestines or gonads will contaminate the kidney preparation. Using watchmaker’s forceps, tease the entire kidney away from the body wall starting at the head kidney and working toward the rear. Place into ice-cold SM. Aspirate vigorously with a 1 ml pipetteman to separate hematopoietic cells (WKM) from renal cells. Filter through 40 mm nylon mesh, wash, centrifuge, and repeat. Perform last filtration step into a Falcon 2054 tube if using for FACS. It is important to filter the WKM cell suspension at least twice before running the sample. PI may be added at this point to 1 mg/ml to exclude dead cells and debris on the flow cytometer. When using, however, compare to samples having PI only and GFP only (if using) to set compensations properly. Otherwise, the signal from PI may bleed into the GFP channel resulting in false positives. Alternatively, add 1:1000 Sytox Red (excited by the 633 nm laser) or Sytox Blue (excited by the 405 nm laser) for dead cell discrimination, as they have no spectral overlap with GFP.

2. Transplanting Whole Kidney Marrow After filtering and washing the WKM suspension three times, centrifuge cells for 5 min at 200g and 4 C. Carefully remove all supernatant. Resuspend cell pellet in 5– 10 ml of ice-cold SM containing 3U heparin and 1U DnaseI to prevent coagulation


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and lessen aggregation. Preventing the cells from aggregating or adhering to the glass capillary needle used for transplantation is critical. Mix the cells by gently pipetting with a 10 ml pipette tip. Keep on ice. For blastulae and embryo transplantation, perform following previous protocols. Between 5 102 and 5 103 cells can be injected into each 48 hpf embryo if the final cell concentration is approximately 5 105 cells/ml.

3. Transplanting Cells into Irradiated Adult Recipients For irradiation of adult zebrafish, we have used a 137Cesium source irradiator typically used for the irradiation of cultured cells (Gammacell 1000). We lightly anaesthetize five animals at a time and then irradiate in sealed Petri dishes filled with fish water (without tricaine). We performed careful calibration of the irradiator using calibration microchips to obtain the dose rate at the height within the irradiation chamber nearest to the 137Cesium point source. We found the dose rate to be uniform among calibration chips placed within euthanized animals in Petri dishes under water, under water alone, or in air alone, verifying that the tissue dosage via total body irradiation (TBI) was accurate. Transplantation into circulation is most efficiently performed by injecting cells directly into the heart. We perform intracardiac transplantation using pulled filament-free capillary needles as above, but we break the needles at a larger bore size of approximately 50 mm. The needle assembly can be handheld and used with a standard gas-powered microinjection station. We have also had limited success transplanting cells intraperitoneally using a 10 ml Hamilton syringe. Engraftment efficiency for WKM is only marginal using this method, but transplantation of T cell leukemia or solid tumor suspensions is highly efficient following irradiation at 20 Gy (Traver et al., 2004).

4. Irradiation 1. Briefly anaesthetize adult zebrafish in 0.02% tricaine in fish water. 2. Place five fish at a time into 60 mm 15 mm Petri dishes (Falcon) containing fish water. Wrap dish with Parafilm and irradiate for length of time necessary to achieve desired dose. 3. Return irradiated animals to clean tanks containing fish water. We have successfully transplanted irradiated animals from 12 to 72 h following irradiation. Using a 20 Gy dose, the nadir of host hematopoietic cell numbers occurs at approximately 72 h postirradiation.

5. Transplantation 1. Prepare cells to be transplanted as above, taking care to remove particulates/ contaminants by multiple filtration and washes. When using WKM as donor cells, we typically make final cell suspensions at 2 105 cells/ml. Keep cells on ice.

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2. Anaesthetize an irradiated animal in 0.02% tricaine in fish water. 3. Transfer ventral side up into a well cut into a sponge wetted with fish water. Under a dissection microscope, remove scales covering the pericardial region with fine forceps. 4. Fill injection needle with 20 ml of cell suspension. Force cells to end of needle with positive pressure and adjust pressure balance to be neutral. Hold needle assembly in one hand while placing gentle pressure on the abdomen of the recipient with the index finger of the other hand. This will position the heart adjacent to the skin and allow visualization of the heartbeat. Insert needle through the skin and into the heart. If the needle is positioned within the heart, and the pressure balance is neutral, blood from the heart will enter the needle and the meniscus will rise and fall with the heartbeat. Inject approximately 5–10 ml by gentle pressure bursts. 5. Return recipient to fresh fish water. Repeat for each additional recipient. Do not feed until the next day to lessen chance of infection.

IV. Enrichment of Hematopoietic Stem Cells The development of many different transplantation techniques now permits the testing of cell autonomy of mutant gene function, oncogenic transformation, and stem cell enrichment strategies in the zebrafish. For HSC enrichment strategies, fractionation techniques can be used to divide WKM into distinct subsets for functional testing via transplantation. The most successful means of HSC enrichment in the mouse has resulted from the subfractionation of whole bone marrow cells with monoclonal antibodies (mAbs) and flow cytometry (Spangrude et al., 1988). We have attempted to generate mAbs against zebrafish leukocytes by repeated mouse immunizations using both live WKM and purified membrane fractions followed by standard fusion techniques. Many resulting hybridoma supernatants showed affinity to zebrafish WKM cells in FACS analyses (Fig. 7). All antibodies showed one of two patterns, however. The first showed binding to all WKM cells at similar levels. The second showed binding to all kidney leukocyte subsets but not to kidney erythrocytes, similar to the pattern shown in the left panel of Fig. 7A. We found no mAbs that specifically bound only to myeloid cells, lymphoid cells, etc when analyzing positive cells by their scatter profiles. We reasoned that these nonspecific binding affinities might be due to different oligosaccharide groups present on zebrafish blood cells. If the glycosylation of zebrafish membrane proteins were different from the mouse, then the murine immune system would likely mount an immune response against these epitopes. To test this hypothesis, we removed both O-linked and Nlinked sugars from WKM using a deglycosylation kit (Prozyme), and then incubated these cells with previously positive mAbs. All mAbs tested in this way showed a time-dependent decrease in binding, with nearly all binding disappearing following 2 h of deglycosylation (Fig. 7A). It thus appears that standard immunization approaches using zebrafish WKM cells elicit a strong immune response against


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[(Fig._7)TD$IG]

Fig. 7

Potential methods of stem cell enrichment. (A) Mouse monoclonal antibodies generated against zebrafish WKM cells react against oligosaccharide epitopes. De-glycosylation enzymes result in time-dependent loss of antibody binding (bold histograms) compared to no enzyme control (left panel and grey histograms). (B) Differential binding of lectins to WKM scatter fractions. Peanut agglutinin splits both the lymphoid and precursor fraction into positive and negative populations (left panels). Potato lectin shows a minor positive fraction only within the lymphoid fraction (right panels). (C) Hoechst 33342 dye reveals a side population (SP) within WKM. 0.4% of WKM cells appear within the verapamil-sensitive SP gate (left panel). Only the lymphoid fraction, where kidney HSCs reside, contains appreciable numbers of SP cells (right panels).

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oligosaccharide epitopes. This response is likely to be extremely robust, because we did not recover any mAbs that reacted with specific blood cell lineages. Similar approaches by other investigators using blood cells from frogs or other teleost species have yielded similar results (L. du Pasquier, M. Flajnik, personal communications). In an attempt to circumvent the glycoprotein issue, new series of immunizations using deglycosylated kidney cell membrane preparations may be effective. Previous studies have shown that specific lectins can be used to enrich hematopoietic stem and progenitor cell subsets in the mouse (Huang and Auerbach, 1993; Lu et al., 1996; Visser et al., 1984). In preliminary studies, we have shown that FITClabeled lectins such as peanut agglutinin (PNA) and potato lectin (PTL) differentially bind to zebrafish kidney subsets. As shown in Fig. 7B, PNA binds to a subset of cells both within the lymphoid and precursor kidney scatter fractions. Staining with PTL also shows that a minor fraction of lymphoid cells binds PTL, whereas the precursor (and other) scatter fractions are largely negative (Fig. 7B). We are currently testing both positive and negative fractions in transplantation assays to determine whether these different binding affinities can be used to enrich HSCs. We have previously demonstrated that long-term HSCs reside in the adult kidney (Traver et al., 2003b). We therefore isolated each of the kidney scatter fractions from gata1a:EGFP transgenic animals and transplanted cells from each into 48 hpf recipients to determine which subset contains HSC activity. The only population that could generate GFP+ cells for over 3 weeks in wild type recipients was the lymphoid fraction. This finding is in accord with mouse and human studies that have shown purified HSCs to have the size and morphological characteristics of inactive lymphocytes (Morrison et al., 1995). Another method that has been extremely useful in isolating stem cells from whole bone marrow is differential dye efflux. Dyes such as rhodamine 123 (Mulder and Visser, 1987; Visser and de Vries, 1988) or Hoechst 33342 (Goodell et al., 1996) allow the visualization and purification of a ‘‘side population’’ (SP) that is highly enriched for HSCs. This technique appears to take advantage of the relatively high activity of multidrug resistance transporter proteins in HSCs that actively pump each dye out of the cell in a verapamil-sensitive manner (Goodell et al., 1996). Other cell types lack this activity and become positively stained, allowing isolation of the negative SP fraction by FACS. Our preliminary studies of SP cells in the zebrafish kidney demonstrated a typical SP profile when stained with 2.5 mg/ml of Hoechst 33342 for 2 h at 28 C (Fig. 7C). This population disappears when verapamil is added to the incubation. Interestingly, the vast majority of SP cells appear within the lymphoid scatter fraction (Fig. 7C). Further examination of whether this population is enriched for HSC activity in transplantation assays is warranted. Finally, there are many other methods to enrich hematopoietic stem and progenitor cells from WKM including sublethal irradiation, cytoreductive drug treatment, and use of transgenic lines expressing fluorescent reporter genes (see Table I). We have shown following 20 Gy doses of g-irradiation that nearly all hematopoietic lineages are depleted within 1 week (Traver et al., 2004). Examination of kidney cytocentrifuge preparations at this time shows that the vast majority of cells are immature


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precursors. That this dose does not lead to death of the animals demonstrates that HSCs are spared and are likely highly enriched 5–8 days following exposure. We have also shown that cytoreductive drugs such as cytoxan and 5-fluorouracil have similar effects on kidney cell depletion, although the effects were more variable than those achieved with sublethal irradiation (A. Winzeler, D. Traver, and L. I. Zon unpublished). Because HSCs are contained within the kidney lymphoid fraction, they can be further enriched by HSC-specific or lymphocyte-specific transgenic markers. Possible examples of transgenic promoters are lmo2, gata2a, or cmyb to mark HSCs and ccr9a, il7r, rag2, lck, or B-cell receptor genes to exclude lymphocytes from this subset (see Table I).

V. In vitro Culture and Differentiation of Hematopoietic Progenitors Hematopoiesis is one of the best-studied models of developmental differentiation because of the multitude of experimental methods developed over the past 60 years to assess the proliferation, differentiation, and maintenance of its cellular constituents. Stem and progenitor cell transplantation into lethally irradiated animal recipients (Ford et al., 1956; McCulloch and Till, 1960) were the first in vivo assays to be developed, followed shortly thereafter by the clonal growth of bone marrow progenitors in vitro (Bradley and Metcalf, 1966). Although these techniques have been substantially refined over the past decades, they still remain the foundation for analyzing the hierarchical organization of vertebrate hematopoietic stem and progenitor cells. In vitro cultures to assess hematopoietic stem and progenitor cell biology generally fall into two categories: growth of progenitor cells on a supportive stromal cell layer, and clonal growth of cells in a semisolid medium with the addition of supplemental cytokines or growth factors. Most stromal culture assays largely derive from the modification and refinement of Dexter cultures (Dexter et al., 1977a, 1977b), whereby stromal cells from hematopoietic organs support the differentiation of HSCs and their downstream progenitors. These early studies were instrumental for the development of cobblestone-area-forming-cell (CAFC) (Ploemacher et al., 1991) and long-term culture initiating cell (LTC-IC) assays, which have been utilized to examine murine (Lemieux et al., 1995) and human (Sutherland et al., 1991) multilineage hematopoietic differentiation. The development of stromal cells from the calvaria of macrophage colony stimulating factor (M-CSF)-deficient mice (Nakano et al., 1994, 1996) were instrumental for examining hematopoietic differentiation of embryonic stem (ES) cells down the hematopoietic pathway, and for differentiation of hematopoietic precursors into multiple mature blood cell types. With refinement, these OP9 cells have proven to be an efficient tool to study T-cell lineage commitment and development (Schmitt et al., 2004; Schmitt and ZunigaPflucker, 2002), once an extremely difficult process to study. To assess the progenitor capacity of normal and mutant zebrafish hematopoietic cells functionally, we created primary zebrafish kidney stromal (ZKS) cells derived

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from the main site of hematopoiesis in the adult fish. Culture of hematopoietic progenitor cells on these stromal cells resulted in their continued maintenance (Stachura et al., 2009) and differentiation (Bertrand et al., 2007, 2010b; Stachura et al., 2009). It also allowed investigation and rescue of a genetic block in erythroid maturation, confirming the utility of these assays (Stachura et al., 2009). Finally, the ZKS culture system has been utilized to investigate the molecular events underlying the progression of T-lymphoblastic lymphoma (T-LBL) to acute T-lymphoblastic leukemia (T-ALL) (Feng et al., 2010). A. Stromal Cell Culture AssaysFS To create a suitable in vitro environment for the culture of zebrafish hematopoietic cells, we isolated the stromal fraction of the zebrafish kidney, the main site of hematopoiesis in the adult fish (Zapata, 1979). The benefit of utilizing hematopoietic stromal layers is two-fold. First, performing culture assays in the zebrafish has been hampered by a paucity of defined and purified hematopoietic cytokines. Most zebrafish cytokines have poor sequence homology to their mammalian counterparts, and as a consequence, have not been well described, characterized, or rigorously tested. Secondly, some hematopoietic cell types, especially T cells, require physical cell-cell interaction for their differentiation.

1. Generation of ZKS Cells To create ZKS cells, kidney was isolated from AB* wild-type fish as described above (also see Stachura et al., 2009). The kidney tissue was sterilized by washing for 5 min in 0.000525% sodium hypochlorite (Fisher Scientific), then rinsed in sterile 0.9X Dulbecco’s PBS. Tissue was then mechanically dissociated by trituration and filtered through a 40 mm filter (BD Biosciences). Flow-through cells (WKM) were discarded, and the remaining kidney tissue was cultured in vacuum plasmatreated vented flasks (Corning Incorporated Life Sciences) at 32 C and 5% CO2.

2. Maintenance and Culture of ZKS Cells ZKS cells are maintained in the following tissue culture medium: 500 ml 350 ml 150 ml 150 mg 15 ml 20 ml 10 ml 100 ml 2 ml

L-15 DMEM (high glucose) Ham’s F-12 Sodium bicarbonate HEPES (1 M stock) Penicillin/streptomycin (5000 U/ml penicillin, 5000 mg/ml streptomycin stock) L-glutamine (200 mM stock) Fetal bovine serum (FBS) Gentamicin sulfate (50 mg/ml stock)


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Medium is made by first adding sodium bicarbonate to the mixture of L-15, DMEM, and Ham’s F-12. Warm the medium to 37 C, and allow the sodium bicarbonate to dissolve. Then, add other medium components and filter-sterilize with 0.22 mm vacuum apparatus. All medium components are available from Mediatech. We utilize FBS from the American Type Culture Collection, but one can use FBS from other sources. It is important to note, however, that different manufacturing lots of FBS investigated in the laboratory differ wildly in their support of hematopoietic progenitor differentiation and proliferation. Once a manufacturing lot is tested and shown to be supportive, we recommend buying a large quantity to minimize experimental variation. ZKS cells are maintained at 32 C and 5% CO2 in a humidified incubator. Cells are grown in vacuum plasma treated 75 cm2 vented flasks (T-75; Corning Incorporated Life Sciences) in 10 ml of medium until 60–80% confluent before passaging. Medium is then removed, and 2 ml trypsin–EDTA (0.25%; Invitrogen) is added to cover the stromal cells. Allow cells to incubate for 5 min at 32 C. Add 8 ml medium to cells to stop trypsinization, pipetting up and down to achieve a single cell suspension. Spin cells for 5 min at 300g, aspirate supernatant and resuspend pellet gently in 10 ml of medium. Take 1 ml of the cell solution, add 9 ml of medium, and move to a new flask. Cells should not be split more than 1:10 to passage, as they are somewhat density-dependent. ZKS cells may be frozen and thawed at a later time. Even though we have never experienced senescence or a decrease in hematopoietic differentiation capacity of ZKS cells in culture, it is useful to perform critical experiments with similar passages of cells to avoid experimental variation. To freeze ZKS cells, first trypsinize a T-75 flask. Spin cells for 5 min at 300g, aspirate supernatant, and resuspend pellet gently in 2 ml of medium. Prepare freezing medium (500 ml of tissue culture-certified DMSO and 1.5 ml of FBS) and aliquot 500 ml into four cryopreservation tubes keeping everything on ice. Add 500 ml of cells to each tube, invert to gently mix, and place tubes into isopropanol-jacketed freezing chamber. Place freezing chamber at –80 C for 24 h. Remove tubes from freezing chamber and place into liquid nitrogen for long-term storage. To thaw ZKS cells at a later date, remove tube from nitrogen, and quickly warm in 37 C water bath. Wear eye protection; if nitrogen seeped into the freezing tubes, the rapid warming may cause the tube to violently rupture. Remove liquid from tube, add slowly to 10 ml of medium in a 15 ml conical tube, and spin at 300g for 5 min. Carefully aspirate all of the medium to remove all traces of DMSO. Resuspend cells in 10 ml of fresh medium and place into T-75 flask. Early the next morning change the medium and determine whether the cells are ready to be passaged or require another day to recover from thawing. As with all tissue culture, strict attention to sterility and cleanliness should be adhered to at all times. All procedures should be performed in a tissue culture laminar flow hood, and it is recommended that vented, filtered flasks be utilized to prevent airborne contamination during culture in incubators.

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3. Protocols for in vitro Proliferation and Differentiation Assays Purify prospective progenitors by FACS as described above. Plate cells on confluent ZKS at a density of 1 104 cells/well in a 12-well tissue culture plate, using 2 ml complete medium per well. Lower density of progenitors is not recommended; if using fewer cells, reduce the size of the tissue culture well and volume of medium. If testing or investigating the effects of growth factors or small molecules, add them to the medium, being sure to have a vehicle only control as well as different concentrations of your experimental factor. 24-well tissue culture plates are extremely useful in this regard, as one can easily plate out a multitude of experimental conditions on one plate. A. Morphological assessment of hematopoietic cells after in vitro culture a. Gently aspirate hematopoietic cells from the ZKS cultures, taking care not to disturb the stromal underlayer. b. Cytocentrifuge up to 200 ml of the hematopoietic cells at 250g for 5 min onto glass slides using a Shandon Cytospin 4 (Thermo Fischer Scientific). It is possible to concentrate the cells before cytocentrifugation at 300g for 10 min. Cytocentrifugation of over 200 ml of cell suspensions is not recommended. c. Perform May-Gr€ unwald/Giemsa staining by allowing slides to air-dry briefly. Then, submerge slide in May-Gr€ unwald staining solution (Sigma Aldrich) for 10 min. Transfer slide to 1:5 dilution of Giemsa stain (Sigma Aldrich) in dH2O for an additional 20 min. Rinse slide in dH2O, and allow to air dry. Coverslip slide with cytoseal XYL mounting medium (Richard-Allan Scientific) and Corning no.1 18 mm square cover glass (Corning). Allow slides to completely dry before visualization on upright microscope, especially if using an oil-immersion lens. B. Proliferation assessment of hematopoietic cells after in vitro culture a. Gently aspirate hematopoietic cells as above. b. Count cells with use of a bright line hemacytometer (Hausser Scientific) using trypan blue dye (Invitrogen) exclusion to assess viability. C. RT-PCR analysis of hematopoietic cells after in vitro culture a. Gently aspirate hematopoietic cells as above. b. Isolate RNA from hematopoietic cells using either Trizol (Invitrogen) or RNAeasy kit (Qiagen). c. Generate cDNA with oligo dT primers and superscript RT-PCR kit (Invitrogen). d. Perform PCR with the desired zebrafish DNA primers. D. Cell labeling and cell division determination of hematopoietic cells after in vitro culture a. Prior to plating cells on ZKS monolayer, wash cells twice with 0.1% bovine serum albumin (BSA) to remove FBS from the medium.


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b. Resuspend cells in 0.1% BSA and 2 ml/ml of 5 mM carboxyfluorescein succinimidyl ester (CFSE; Invitrogen) at room temperature for 10 min, in the dark. c. Wash cells with complete medium supplanted with an additional 10% FBS twice. d. Save 1/10 of the culture and perform FACS (Day 0 time point). Culture remaining cells in complete medium as described above. e. For analysis, remove hematopoietic cells from culture at desired time points as described above and FACS. CFSE is read in the FL-1 channel (on FACS caliber) or with most GFP filters (FACS Aria I and II, LSR-II), and will decrease in fluorescence intensity as cells divide. Compare divisions to Day 0 time point with FloJo software (TreeStar, Ashland, OR, USA). We recommend using the BD LSR-II flow cytometer, as the different scatter profile of mature cells is easily distinguished and directly comparable to profiles shown in Fig. 5.

B. Clonal Methylcellulose-based Assays Although stromal in vitro culture methods have been instrumental for the investigation of hematopoiesis, culturing bulk populations of progenitor cells on stroma cannot distinguish between homogeneous multipotent progenitor populations or heterogeneous lineage-restricted populations without performing limiting dilution assays. The development of clonal in vitro cultures by Metcalf and colleagues allowed not only the growth of murine bone marrow progenitors (Bradley and Metcalf, 1966), but also the study and quantitation of progenitor numbers during hematological disease (Bradley et al., 1967) and exposure to irradiation (Robinson et al., 1967). These assays were utilized to investigate the ontogeny of the developing murine hematopoietic system (Moore and Metcalf, 1970), and refined to study human hematopoietic progenitors dysregulated during leukemogenesis (Moore et al., 1973a, 1973b). Importantly, the utilization of clonal assays was instrumental for the identification and validation of CSFs, secreted proteins that stimulate the differentiation of specific hematopoietic lineages. The ability to isolate, recombinantly produce, and test these factors was a huge advance in hematological research, allowing the sensitive analysis of progenitor differentiation, proliferation, and restriction in the murine and human blood system. This capability to grow progenitors in vitro to test their differentiation capacity in an unbiased manner has greatly advanced the current understanding of hematopoietic lineage restriction. The isolation of putative lineage-restricted daughter cells by FACS coupled with in vitro clonal analysis was pivotal in identifying multipotent (Akashi et al., 2000; Kondo et al., 1997), oligopotent (Akashi et al., 2000), and monopotent progenitor (Mori et al., 2008; Nakorn et al., 2003) intermediates downstream of HSCs in the murine system. While zebrafish likely possess multipotent, oligopotent, and monopotent progenitor cells, their existence has never been proven and remains speculative (see Fig. 4). To investigate whether these cells exist in zebrafish, we developed assays to

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[(Fig._8)TD$IG]

Fig. 8 Recombinantly generated and purified Gcsf and Epo encourage myeloid and erythroid differentiation, respectively, from zebrafish hematopoietic progenitors in clonal methylcellulose assays. (A) Experimental schematic for isolation and culture of mpx:GFP, gata1a:DsRed cells from the precursor (blue) fraction of adult WKM. (B) Brightfield images (top row), mpx: GFP fluorescence (second row), gata1a:DsRed fluorescence (third row), and merged images (bottom row) of colonies grown in various growth factor conditions from the precursor fraction of WKM (conditions listed along top row of images). All images in B taken at 100. Scale bars in top left panels are 50 mm. Arrowheads in top right panels denote mixed colonies. (D) Colonies isolated from precursor fraction methylcellulose cultures cytospun and stained with May Gr€ unwald/Giemsa. Tight colonies were isolated from cultures with only carp serum and Epo (left column), while ruffled and spread colonies were isolated from cultures containing carp serum and Gcsf (right column). All images were taken at 1000, and scale bar in bottom right is 20 mm. (E) RTPCR analysis of colonies isolated from precursor fraction of methylcellulose cultures. Colony morphology is listed on left, and genes assayed are listed along top of gel images. (See Plate no. 8 in the Color Plate Section.)


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investigate progenitors in the zebrafish in a clonal manner by modifying existing methylcellulose culture techniques, exogenously adding the recently identified zebrafish recombinant cytokines erythropoietin (Epo) (Paffett-Lugassy et al., 2007) and granulocyte Gcsf (Liongue et al., 2009) to quantitate the number of myeloid and erythroid progenitors in adult kidney marrow scatter fractions (Fig. 8) (Stachura et al., 2011). This level of precision allows the further testing of prospective hematopoietic progenitors in normal and mutant zebrafish, allowing more careful investigation of lineage determination and its conservation among vertebrate animals. In addition, it allows comparison of hematopoietic progenitor cells and their response to cytokines, furthering our understanding of cytokine signaling. Furthermore, the ability to examine blocks in hematopoietic differentiation, aberrant gene expression, and proliferative regulation is now possible in mutant fish already (and currently being) generated. Finally, it also allows the rapid screening of small molecules, blocking antibodies, and other drug compounds that may affect lineage differentiation, maturation, and proliferation.

1. Methylcellulose To develop a clonal assay to further enumerate and characterize progenitor cells in the zebrafish, we utilized methylcellulose; a semi-solid, viscous cell culture medium used in murine and human progenitor studies. The nature of methylcellulose culture allows individual progenitor cells to develop isolated colonies within the medium, where they can be enumerated after several days in culture. In addition, the use of methylcellulose allows examination of colony morphology and subsequent isolation for further characterization by morphological examination and gene expression.

2. Methylcellulose Stock Preparation Prepare 2.0% methylcellulose by adding 20 g of methylcellulose powder (Sigma Aldrich) to 450 ml of autoclaved H2O and boiling for 3 min. Allow mixture to cool to room temperature before adding 2 L-15 medium powder (Mediatech). Then, adjust the weight of the methylcellulose mixture to 1000 g with sterile water. Methylcellulose should be allowed to thicken at 4 C overnight before being aliquoted and stored at –20 C.

3. Methylcellulose Clonal Assays Complete methylcellulose medium: 10 ml 4.9 ml 2.1 ml 2 ml 300 ml 200 ml 200 ml 40 ml

2.0% methylcellulose stock DMEM (high glucose) Ham’s F-12 FBS HEPES (1 M stock) Penicillin/streptomycin (5000 U/ml and 5000 mg/ml stock, respectively) L-glutamine (200 mM stock) Gentamicin sulfate (50 mg/ml)

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To perform experiments in triplicate, add 3.5 ml of complete methylcellulose to sterile round bottom 14 ml tubes (Becton Dickinson) with 5 ml syringes and 16 gauge needles for each condition. Cells of interest (prospective progenitors) should be isolated, counted, and resuspended in 100 ml of ZKS medium and added to complete methylcellulose, along with cytokines, small molecules, or other agents to be investigated. For myeloid differentiation, add 1% carp serum and 0.3 mg/ml recombinant zebrafish Gcsf to methylcellulose medium. For erythroid differentiation, add 1% carp serum and 0.1 mg/ml recombinant zebrafish Epo to methylcellulose medium. Cytokines and additives should not total more than 10% of the total volume, as the medium will not be viscous enough to discern individual colonies. To observe separable, individual colonies, cells should be resuspended at 1 104–5 104 cells/ml. Tightly cap tubes, and gently vortex solution to mix. In triplicate, aliquot 1 ml of solution into 35 mm Petri dishes (Becton Dickinson). Swirl Petri dishes to distribute the methylcellulose culture evenly, and place plates in a humidified 15 cm dish (made by placing a plate of sterile dH2O inside the 15 cm dish) at 32 C and 5% CO2. Plates should be removed 7 days after plating for microscopic examination, colony isolation, and gene expression analyses. As with all tissue culture, strict attention to sterility and cleanliness should be adhered to at all times. All procedures should be performed in a tissue culture laminar flow hood.

4. Enumeration of Colony Forming Units (CFUs) CFUs are a measurement of how many progenitors are present in a given population of cells; if an individual cell has the capability to proliferate and divide into mature blood cells under certain growth conditions, it will make an individual colony. For example, if 100 putative myeloid progenitor cells are plated under conditions suitable for myeloid differentiation and one myeloid colony arises, 1:100 of the cells plated was a myeloid CFU. To perform enumeration of CFUs, observe and count colonies on an inverted microscope after 7 days in culture. Counting with a 5 or 10 objective (50–100 magnification) is recommended, with the aperture closed down slightly to grant high contrast, which aids in the visualization of colonies. Be careful not to disturb the dish; even though methylcellulose is viscous, excessive movement of the plates will cause colonies to move, complicating further analysis. Depending on the cytokines, growth factors, and other culture additives, colony shape, size, and color will be different; be sure to note and record all of this information. Inclusion of a transgenic marker will aid in identification of colony type and morphology as shown in Figure 8 B and C, whereby erythroid colonies express DsRed driven by the erythroid-specific gata1a promoter, and myeloid colonies express GFP driven by the myeloid-specific mpx promoter. If cultures are to be returned to incubator do not remove Petri plate lids, and take care to wipe down surfaces with 70% ethanol before starting experiment.


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5. Picking and Analyzing Colonies from Methylcellulose Hematopoietic colonies can be carefully plucked from methylcellulose cultures with a pipetteman, preferably a p20 with a fine tip. Pay attention to pick only individual colonies, placing them into 1.5 ml eppendorf tubes with 200 ml of PBS. Pipette up and down gently in the PBS to remove traces of methylcellulose from your tip, and to break up the colony. It is possible to pool colonies of similar morphology for analyses, especially when large cell numbers are required. Colonies may be cytospun and stained with May-Gr€ unwald/Giemsa (Sigma Aldrich) as described above. In addition, colonies may be subjected to RT-PCR analysis for mature lineage gene transcripts as described above.

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David L. Stachura and David Traver Traver, D. (2004). Cellular dissection of zebrafish hematopoiesis. Methods Cell. Biol. 76, 127–149. Traver, D., Herbomel, P., Patton, E. E., Murphy, R. D., Yoder, J. A., Litman, G. W., Catic, A., Amemiya, C. T., Zon, L. I., Trede, N. S. (2003a). The Zebrafish as a Model Organism to Study Development of the Immune System. Academic Press, New York. Traver, D., Paw, B. H., Poss, K. D., Penberthy, W. T., Lin, S., Zon, L. I. (2003b). Transplantation and in vivo imaging of multilineage engraftment in zebrafish bloodless mutants. Nat. Immunol. 4, 1238–1246. Traver, D., Winzeler, E.A., Stern, H.M., Mayhall, E.A., Langenau, D.M., Kutok, J.L., Look, A.T., and Zon, L.I. (2004). Biological effects of lethal irradiation and rescue by hematopoietic cell transplantation in zebrafish. Blood, In press. Trede, N. S., and Zon, L. I. (1998). Development of T-cells during fish embryogenesis. Dev. Comp. Immunol. 22, 253–263. Visser, J. W., Bauman, J. G., Mulder, A. H., Eliason, J. F., and de Leeuw, A. M. (1984). Isolation of murine pluripotent hemopoietic stem cells. J. Exp. Med. 159, 1576–1590. Visser, J. W., and de Vries, P. (1988). Isolation of spleen-colony forming cells (CFU-s) using wheat germ agglutinin and rhodamine 123 labeling. Blood Cells 14, 369–384. Ward, A. C., McPhee, D. O., Condron, M. M., Varma, S., Cody, S. H., Onnebo, S. M., Paw, B. H., Zon, L. I., Lieschke, G. J. (2003). The zebrafish spi1 promoter drives myeloid-specific expression in stable transgenic fish. Blood 102, 3238–3240. Weissman, I., Papaioannou, V., and Gardner, R. (1978). Fetal hematopoietic origins of the adult hematolymphoid system. In ‘‘Differentiation of Normal and Neoplastic Cells,’’ (B. Clarkson, P. A. Marks, and J. E. Till, eds.), pp. 33–47. Cold Spring Harbor Laboratory Press, New York. Westerfield, M. (2000). The Zebrafish Book. A Guide for the Laboratory use of Zebrafish (Danio rerio), 4th ed. University of Oregon Press, Eugene. Willett, C. E., Cortes, A., Zuasti, A., and Zapata, A. G. (1999). Early hematopoiesis and developing lymphoid organs in the zebrafish. Dev. Dyn. 214, 323–336. Willett, C. E., Zapata, A. G., Hopkins, N., and Steiner, L. A. (1997). Expression of zebrafish rag genes during early development identifies the thymus. Dev. Biol. 182, 331–341. Wittamer, V., Bertrand, J.Y., Gutschow, P.W., and Traver, D. (2011). Characterization of the Mononuclear Phagocyte System in Zebrafish. Blood. Yoder, M., Hiatt, K., and Mukherjee, P. (1997a). In vivo repopulating hematopoietic stem cells are present in the murine yolk sac at day 9.0 postcoitus. Proc. Natl. Acad. Sci. U.S.A. 94, 6776. Yoder, M. C., Hiatt, K., Dutt, P., Mukherjee, P., Bodine, D. M., Orlic, D. (1997b). Characterization of definitive lymphohematopoietic stem cells in the day 9 murine yolk sac. Immunity 7, 335–344. Yokota, T., Huang, J., Tavian, M., Nagai, Y., Hirose, J., Zuniga-Pflucker, J. C., Peault, B., Kincade, P. W. (2006). Tracing the first waves of lymphopoiesis in mice. Development 133, 2041–2051. Zapata, A. (1979). Ultrastructural study of the teleost fish kidney. Dev. Comp. Immunol. 3, 55–65. Zapata, A., and Amemiya, C. T. (2000). Phylogeny of lower vertebrates and their immunological structures. Curr. Top Microbiol. Immunol. 248, 67–107. Zhu, H., Traver, D., Davidson, A. J., Dibiase, A., Thisse, C., Thisse, B., Nimer, S., Zon, L. I. (2005). Regulation of the lmo2 promoter during hematopoietic and vascular development in zebrafish. Dev. Biol. 281, 256–269. Zovein, A. C., Hofmann, J. J., Lynch, M., French, W. J., Turlo, K. A., Yang, Y., Becker, M. S., Zanetta, L., Dejana, E., Gasson, J. C., Tallquist, M. D., Iruela-Arispe, M. L. (2008). Fate tracing reveals the endothelial origin of hematopoietic stem cells. Cell. Stem Cell 3, 625–636.

CHAPTER 5

Zebrafish Lipid Metabolism: From Mediating Early Patterning to the Metabolism of Dietary Fat and Cholesterol Jennifer L. Anderson,* Juliana D. Carten* and Steven A. Farber Carnegie Institution for Science, Department of Embryology, Baltimore, Maryland, USA

I.

II. III. IV. V.

VI. VII. VIII.

Abstract Abbreviations Introduction A. The Need for Whole Animal Studies of Lipid Metabolism B. Larval Zebrafish as a Model of Vertebrate Lipid Metabolism Lipid Metabolism in Developing Zebrafish Yolk Metabolism During Early Vertebrate Development Lipid Signaling During Early Zebrafish Development A. Lipid Modifications Influence Primordial Germ Cell Migration Visualizing Lipid Metabolism in Larval and Adult Zebrafish A. Lipophilic Dyes B. BODIPY Fatty Acid Analogs C. BODIPY Cholesterol D. Fluorescent Reporters of Lipid Metabolism Triple Screen: Phospholipase, Protease and Swallowing Function Assays Zebrafish Models of Human Dyslipidemias Summary Acknowledgments References

Abstract Lipids serve essential functions in cells as signaling molecules, membrane components, and sources of energy. Defects in lipid metabolism are implicated in a number of pandemic human diseases, including diabetes, obesity, and hypercholesterolemia. *

These two authors contributed equally to the work.

METHODS IN CELL BIOLOGY, VOL 101 Copyright 2011. Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-387036-0.00005-0

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approaches for disease prevention and treatment. Numerous studies have shown that the zebrafish is an excellent model for vertebrate lipid metabolism. In this chapter, we review studies that employ zebrafish to better understand lipid signaling and metabolism. ABBREVIATIONS LCFA, long chain fatty acid; LD, lipid drop; MCFA, medium chain fatty acid; MTP, microsomal triglyceride transfer protein; SCFA, short chain fatty acid, TAG, triacylglycerol

I. Introduction Lipids play essential roles in cells as signaling molecules, membrane components, and sources of fuel. Given their necessity for proper cellular function, it is not surprising that defects in lipid metabolism underlie a number of human diseases, including obesity, diabetes, and atherosclerosis (Joffe et al., 2001; McNeely et al., 2001; Watanabe et al., 2008). In 2007–08, one-third of US adults and 18% of children were classified as obese (Flegal et al., 2010; Ogden et al., 2010), with obesity and type 2 diabetes on the rise worldwide (Misra and Khurana, 2008). The globalization of the high-fat western diet and the concurrent rise in the incidence of lipid disorders has provided an impetus to better understand lipid metabolism in the context of metabolic dysfunction. This need to investigate the role of lipids in metabolic disease has brought into focus unanswered questions in the field. For instance, although the genes involved in cholesterol and fatty acid (FA) uptake in intestinal cells have been identified, their exact mechanisms of action are highly debated or largely unknown (Klett and Patel, 2004; Nassir et al., 2007; Nickerson et al., 2009; Shim et al., 2009; Stahl et al., 1999). Such gaps in our understanding of these genes and how they function hinder the development of effective therapeutics for lipid disorders and reveal a need to create better approaches to address them. In this chapter, we will review novel approaches undertaken to study lipid signaling and metabolism using the zebrafish model organism, with an emphasis on presenting a diverse array of techniques employed to visualize lipid metabolism during various stages of zebrafish development.

A. The Need for Whole Animal Studies of Lipid Metabolism In vitro studies have laid much of the groundwork for our biochemical understanding of vertebrate lipid metabolism and continue to comprise many of the studies done in the field today; however, a number of caveats arise when attempting to study lipid metabolism in vitro. Such studies are often performed in transformed cultured cells, such as liver HepG2, intestinal Caco2, and adipocyte 3LT3 cells. These cell

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lines, comprised of a single cell type, cannot duplicate the cellular heterogeneity of an entire organ, such as the intestine, which is composed of stem, enteroendocrine, immune, and goblet cells. These multiple cell types are known to influence each other through paracrine signaling that can have global effects on lipid uptake and processing. Furthermore, bile, intestinal mucus, and the gut microbiota are all known to greatly influence dietary lipid processing and absorption in the intestine (Backhed et al., 2004; Field et al., 2003; Kruit et al., 2006; Martin et al., 2008; Moschetta et al., 2005; Pack et al., 1996; Titus and Ahearn, 1992; Turnbaugh et al., 2008), and are absent in cultured cell models. For these reasons, employing whole animal in vivo strategies, in addition to cultured cell work, is vital to better understand how metabolic dysfunction arises and manifests itself in an organism. The importance of utilizing whole animal models to identify drugs to ameliorate metabolic dysfunction is exemplified by how the cholesterol absorption inhibitor ezetimibe (Zetia, Vitorin; Merck-Schering Plough) was developed (Van Heek et al., 1997), and how its mechanism of action was defined (Van Heek et al., 2000). Studies of bile duct-cannulated rats treated with ezetimibe revealed that the bioactive compound responsible for the diminished effect on cholesterol absorption was a glucuronidated form of ezetimibe. Since in vitro studies cannot recreate the complex interplay of neural, chemical, and hormonal cues known to regulate metabolic processes in vivo, there is a need for whole animal approaches to study lipid metabolism as it plays out in a multicellular context.

B. Larval Zebrafish as a Model of Vertebrate Lipid Metabolism The larval zebrafish is well suited for whole animal studies of lipid metabolism. Larval zebrafish possess many of the same gastrointestinal organs present in humans (e.g., the liver, intestine, exocrine and endocrine pancreas, and gallbladder) (Lieschke and Currie, 2007; Pack et al., 1996; Schlegel and Stainier, 2006; Wallace and Pack, 2003; Wallace et al., 2005) as well as the specialized cell types involved in lipid absorption and processing (e.g., intestinal enterocytes, fat-storing adipocytes, hepatocytes in the liver, and acinar cells of the pancreas) (Wallace et al., 2005). These digestive organs and the cell types present in them are formed using similar developmental programs as in mammals with hhex (Wallace et al., 2001), pdx1 (Yee et al., 2005), and shha (Wallace and Pack, 2003) genes playing critical roles in liver and pancreas organogenesis in zebrafish. Notch and its ligands, Delta and Jagged, play a role in the developing pancreas, both by maintaining undifferentiated precursors and regulating acinar, exocrine, and endocrine differentiation (Apelqvist et al., 1999; Esni et al., 2004; Hald et al., 2003; Jensen et al., 2000; Murtaugh et al., 2003; Zecchin et al., 2004). Due to the optical transparency of larvae, these organs and their multiple cell types can all be directly observed through the body wall without the need for invasive surgical manipulations. Work from our lab and others has shown that zebrafish express many of the genes needed to transport and metabolize lipids, such as the lipoprotein gene

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microsomal triglyceride transfer protein (MTP) (Marza et al., 2005) and the FA transport protein (slc27a) and acyl-CoA synthetase (acsl) gene families (Thisse et al., 2005; Thisse et al., 2004; Miyares, unpublished). Zebrafish express the putative cholesterol transporter Niemann-Pick C1-Like 1 (npc1l1) (Farber, unpublished) and treatment of larvae with the human drug ezetimibe, which works through an NPC1L1-dependent pathway and is used to treat hypercholesterolemia, blocks intestinal cholesterol absorption (Clifton et al., 2010). Apolipoprotein C2, a gene needed for lipoprotein assembly in humans, is expressed and required during zebrafish larval development; larvae injected with an apoc2 morpholino exhibit an unabsorbed yolk phenotype (Pickart et al., 2006). Additionally, the enzymes needed to synthesize lipid-signaling molecules, such as prostaglandins and thromboxanes, are highly conserved in zebrafish and these enzymes can be inhibited by commonly used nonsteroidal anti-inflammatory drugs (Grosser et al., 2002). We have presented here only a small sampling of the numerous studies that document the homologies between human and zebrafish lipid metabolism and validate the zebrafish as a model for investigating vertebrate lipid metabolism.

II. Lipid Metabolism in Developing Zebrafish During the first four days of development, a zebrafish embryo relies entirely on its yolk sac for the nutrients needed to sustain its growth and survival. Yolk lipids are the source of essential fat-soluble vitamins and triacylglycerol (TAG), as well as cholesterol, a required component of cell membranes and a precursor for bile acids (Babin et al., 1997; Bownes, 1992; Munoz et al., 1990). Lipids enter the developing embryo at the yolk and embryo interface, an area termed the yolk syncytial layer (YSL). In the YSL, lipoproteins (e.g., apoE, apoA1, apocII, and vitellogenin) and a host of lipid-modifying enzymes (e.g., Mtp) transport lipids from the yolk ball to the embryo (Babin et al., 1997; Marza et al., 2005). Once the circulatory system forms, yolk, hepatic, and intestinal lipids are transported by lipoproteins to specific target tissues throughout the organism via the bloodstream. By 5–6 days postfertilization (dpf), the yolk is depleted and larvae must eat to acquire lipids. Both in the wild and the laboratory, zebrafish consume a lipid-rich diet (10% by weight) high in TAG, phospholipids, and sterols (Enzler et al., 1974; Spence et al., 2008). Prior to absorption by the intestine, these yolk lipids must be processed and solubilized by the digestive enzymes and bile that make up the intraluminal intestinal milieu. As in humans, zebrafish bile is produced by hepatocytes and secreted into an extensive network of intrahepatic ducts, which drains into the gall bladder. In response to hormonal stimulation triggered by food consumption, bile is released into the intestinal lumen to emulsify dietary fat and facilitate its absorption by intestinal enterocytes. The main components of bile are bile acids, phospholipids, and salts, which together promote the formation of micelles by


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inserting themselves between lipid bilayers and reducing the surface tension to allow for membrane curvature (Tso and Fujimoto, 1991; Verkade and Tso, 2001). After dietary fat is emulsified, TAG and phospholipids must be broken down by luminal lipases to release free FA or mono- and di-acylglycerols, which can then enter the specialized absorptive cells (enterocytes) that line the gut (Thomson et al., 1993). The exocrine pancreas is the main source of these fat-splitting enzymes in larval zebrafish (Hama et al., 2009) and known in mammals to secrete lipase- and proteaserich pancreatic juice into the gall bladder (Layer and Keller, 1999). After being emulsified and broken down, TAG can be absorbed by enterocytes, the main absorptive cell type of the zebrafish intestine. These cells are highly reminiscent of mammalian enterocytes (Buhman et al., 2002), with the characteristic microvilli and basal nuclei (Fig. 1). Following food consumption, zebrafish accumulate cytoplasmic lipid drops (LD) in their enterocytes (Walters, unpublished). From there, fats are likely burned via oxidative pathways in the mitochondria or peroxisomes or packaged into chylomicrons, which are secreted from the basolateral surface of enterocytes into lymphatic or blood vessels (Field, 2001; Levy et al., 2007). In chickens, chylomicron production and secretion is highly conserved, with the exception that lipoproteins are secreted from the intestine directly into the portal vein and thus are termed portomicrons (Bensadoun and Rothfeld, 1972; Griffin et al., 1982). This difference has led some to propose that a similar portomicron process occurs in fish (Robinson and Mead, 1973); however, careful ultrastructural studies and isotopic lipid labeling experiments suggest that fish and mammals produce chylomicrons containing largely similar lipoproteins (Sire et al., 1981; Skinner and Rogie, 1978). While it remains to be seen how closely the zebrafish system will model human intestinal lipoprotein metabolism, it is likely that many of the mechanisms of lipoprotein production are conserved.

III. Yolk Metabolism During Early Vertebrate Development To mobilize lipids stockpiled in the yolk sac, rodents express the same genes required for lipoprotein production as those expressed in fully differentiated intestinal enterocytes and liver hepatocytes (e.g., MTP and the apolipoproteins apoE, apoB, and apo-A-IV and apo-A-I (Elshourbagy et al., 1985; Farese et al., 1996; Plonne et al., 1996)). Without these genes, the rapid transport of essential nutrients from the yolk to developing embryonic tissues is impaired and development cannot proceed normally. Not surprisingly, mice null for MTP (Raabe et al., 1998) and apoB (Farese et al., 1996) are embryonic lethal, and DGAT-2 null mice, which lack the enzyme needed to synthesize diacylglycerol, exhibit stunted embryonic growth and die perinatally (Stone et al., 2004). The zebrafish YSL expresses a number of lipoprotein-encoding mRNAs that are also expressed later in the larval intestine and liver, including MTP (Marza et al., 2005), apoC2 (Farber Lab, unpublished), intestinal FA binding protein

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Fig. 1 The intestinal enterocytes of zebrafish and mammals exhibit a high degree of morphological similarity. Electron micrograph of enterocytes from a larval zebrafish (6 dpf). Zebrafish enterocytes exhibit the typical characteristics of mammalian polarized intestinal cells including apical microvilli, which extend into the intestinal lumen (L) and form the brush border, as well as basal nuclei (N). Organelles and subcellular details including mitochondria (M), Golgi bodies (G), and endoplasmic reticulum (ER) are apparent throughout the enterocytes. For comparison, see mouse EMs in Buhman et al. (2002).

(Sharma et al., 2004), and apoE and apoA-I (Babin et al., 1997). Due to the parallel gene expression patterns observed between the embryonic YSL and larval digestive organs, we hypothesize that yolk utilization during early zebrafish embryogenesis is a regulated process highly analogous to intestinal and hepatic lipoprotein-mediated lipid transport.


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To study the role of lipid metabolism genes in the YSL during early zebrafish development, we utilize synthetic antisense morpholinos (MO) to attenuate gene expression and assay subsequent yolk metabolism. MOs are widely used in the zebrafish community to knock down mRNA levels (Heasman, 2002; Nasevicius and Ekker, 2000), and numerous studies have phenocopied mutants by targeting particular mRNA transcripts using this method (Dutton et al., 2001; Karlen and Rebagliati, 2001; Urtishak et al., 2003). By injecting MOs targeting lipid-specific genes into the yolk of 1–4 cell stage embryos, followed by yolk injection of fluorescent lipid analogs, such as BODIPY-labeled FA (Fig. 2A), we can directly assess the necessity of a given gene for yolk metabolism during early larval development.

[(Fig._2)TD$IG]

Fig. 2 BODIPY lipid analogs enable studies of yolk metabolism during early zebrafish development. To assay the function of apoc2 during zebrafish development, embryos were injected with an apoc2 morpholino at the 1–4 cell stage followed by injection of a fluorescent fatty acid (BODIPY-C12) at 24 h postfertilization (hpf). (A) At 48 hpf, embryos injected with BODIPY-C12 retain the fluorescent analog primarily in their yolk. (B, C) apoc2 morphants exhibit an enlarged yolk phenotype (arrowhead) at 48 and 72 hpf, indicating that apoc2 is necessary for yolk utilization during larval development. (D, E) To determine the metabolic consequences of apoc2 deficiency, 1 dpf wild-type and apoc2 morphant larvae were injected with BODIPY-C12 and assayed using fluorescent thin layer chromatography (TLC) 2 days later. (D) TLC analysis shows that BODIPY-C12 is incorporated primarily into triacylglycerol (TAG), diacylglycerol (DG), and phosphatidylcholine (PC) in both wild-type and apoc2 morphant larvae. (E) apoc2 morphants exhibit defects in PC and lysophosphatidylcholine (LPC) metabolism. Total lipid fluorescence was quantified from TLC plates run with total lipids extracted from wild-type and apoc2 morphants. Triacylglycerol (TAG), diacylglycerol (DG), BODIPY C12:0 (C12), phospholipids (PL), PC, and LPC.* p < 0.05.

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Following sequential injections of the morpholino and BODIPY-labeled FA, embryos are allowed to develop and total larval lipids are extracted at 2–3 dpf. We assay the incorporation of the fluorescent FA into metabolites using thin layer chromatography (TLC). This approach allows one to identify metabolic abnormalities in morphants that may not exhibit obvious morphological phenotypes. Furthermore, we can assay the effects of essential genes at earlier stages of larval development prior to lethality caused by insufficient transcript amounts. We are currently using this technique to better understand how the apolipoprotein apoc2 functions in yolk utilization during early zebrafish development. This gene was first identified in a MO screen targeting secreted proteins of the unknown function. In a screen of approximately 100 MOs, we identified one with an unabsorbed yolk phenotype (Fig. 2B and 2C). TLC analysis revealed that these larvae exhibit metabolic defects in phospholipid production, as morphants incorporated less BODIPY-C12 into phosphatidylcholine (PC) and lysophosphatidylcholine (LPC) (Fig. 2D and 2E). Unabsorbed yolk and phospholipid metabolic deficiency in apoc2 morphants suggest that this gene has a function in the yolk or YSL during zebrafish development that is unrelated to its known role in the activation of lipoprotein lipase in peripheral tissues (Jong et al., 1999).

IV. Lipid Signaling During Early Zebrafish Development It is not surprising that many lipid-signaling molecules are derived from membrane phospholipids. Such a system allows for rapid transmission of extracellular signals via membrane, and constituents to the intracellular environment to activate appropriate signaling cascades and cellular responses. The synthesis of membranederived signaling lipids is often dependent on phospholipases, enzymes that cleave phospholipids in response to specific cellular signals. Phospholipase A2 (PLA2) is one such enzyme that catalyzes the hydrolysis of the second fatty acyl bond of glycerophospholipids to liberate lysophospholipid and free FA, both lipid-signaling precursors. In the last decade, PLA2 activity and its products have been implicated in a wide range of cellular phenomena including inflammation, membrane remodeling, and cancer. Lipid signaling events during early development are not well elucidated and have only been recently explored in zebrafish. Studies examining PLA2 enzymatic activity throughout larval development using whole embryo lysates revealed varying activity levels during different stages of development, with larvae exhibiting a significant PLA2 activity during somitogenesis (Fig. 3A). Pharmacological inhibition of PLA2 activity causes developmental arrest at epiboly (Farber et al., 1999). Further experiments identified the primary source of phospholipase activity to be the Ca2+-dependent cytosolic PLA2 (cPLA2) type, a crucial mediator of stimulusinduced eicosanoid release. cPLA2 activity releases the polyunsaturated FA arachidonic acid from membranes, allowing it to participate in the synthesis of eicosanoids, a potent class of lipid-signaling molecules that exhibit both paracrine and


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Fig. 3

Phospholipase activity is required for proper zebrafish development. (A) Cytosolic phospholipase A2 (cPLA2) mRNA levels and PLA2 enzymatic activity were quantified from whole embryo lysates during early stages of zebrafish development. cPLA2 expression peaks during somitogenesis (10 h) while enzymatic activity steadily increases as development proceeds. PLA2 activity is required to generate prostaglandins. (B) Blocking prostaglandin production during early zebrafish development through inhibition of prostaglandin–endoperoxide synthase (Ptgs1) via morpholino knockdown results in developmental arrest at epiboly. Developmental arrest can be rescued by adding back the exogenous enzyme product (PGE2). Reproduced with permission by Development (Speirs et al., 2010). (C) Embryos exposed to the PGE2 analog (16,16-dimethyl-PGE2; dmPGE2) exhibit increased expression of runx11 and cmyb1 as evidenced by in situ hybridization. These genes are expressed in the ventral wall of the dorsal aorta in a region analogous to the mammalian aorta–gonad–mesonephros and are required for mammalian hematopoietic stem cell development. Reprinted by permission from Macmillan Publishers Ltd.: Nature (North et al., 2007). Prostaglandin E2 regulates vertebrate haematopoietic stem cell homeostasis. Nature 447, 1007–11, copyright 2007.

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autocrine influences on cells and tissues (Burke and Dennis, 2009; Clark et al., 1991) and are commonly associated with provoking inflammatory and immune responses. The developmental arrest observed in cPLA2 morphants suggests that cPLA2 activity and eicosanoid signaling are essential for early embryonic patterning, pointing to a novel role for lipid signaling during embryogenesis. Arachidonic acid can act as a substrate for a number of other lipid modifying enzymes (e.g., cyclooxygenase) that are critical for cell movements required to pattern the early zebrafish embryo. Cha et al. (2006) found that PGE2, an eicosanoid downstream of cyclooxygenase, is essential for the morphogenic movements of convergence and extension (Cha et al., 2006; Cha et al., 2005). Moreover, the arrest of epiboly that results from the inhibition of cyclooxygenase 1 (also known as Prostaglandin–endoperoxide synthases, Ptgs1) using an antisense morpholino oligonucleotide can be completely rescued by the addition of PGE2 (Speirs et al., 2010) (Fig. 3B). The importance of PGE2 during zebrafish development was further demonstrated in a small molecule screen performed in zebrafish larvae (36 hpf) by North et al. (2007). Using this approach PGE2 was found to impact hematopoietic stem cell proliferation as evidenced by a dramatic increase in the expression of hematopoietic markers in larvae treated with a PGE2 analog (North et al., 2007) (Fig. 3C). Taken together, these data provide evidence for the importance of PLA2-derived lipid mediators during zebrafish development.

A. Lipid Modifications Influence Primordial Germ Cell Migration In addition to uncovering a role for lipid signaling during the early movements of gastrulation, work on the zebrafish has shown that lipid modifications influence another critical cellular movement during early embryonic development: primordial germ cell (PGC) migration. PGC migration in zebrafish embryos can be visualized in embryos as early as 80% epiboly by injecting in vitro transcribed, capped GFPnanos mRNA (which consists of the coding sequence of GFP fused to the 3’UTR of the nanos gene) into zebrafish embryos at the one-cell stage. GFP-nanos message and protein are stabilized preferentially in PGCs such that they maintain their fluorescence throughout early development, facilitating the detailed study of PGC migratory behavior (Doitsidou et al., 2002). Because lipid metabolism is highly conserved across vertebrates, human drugs can be used to block particular steps in metabolic pathways to determine what pathway or metabolites are necessary for a developmental process, such as PGC migration. In Drosophila, loss of 3-hydroxyl-3-methylglutaryl-CoA reductase (HMGCoAR) results in PGC migration defects (Van Doren et al., 1998). Similarly, studies in zebrafish have shown that pharmacologic inhibition of HMGCoAR by atorvastatin (Lipitor) results in abnormal development and PGC migration (Thorpe et al., 2004). HMGCoAR is a rate-limiting step in the synthesis of cholesterol and is the target of statins (Fig. 4). Zebrafish embryos soaked in


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Fig. 4

Cholesterol biosynthesis and protein lipidation are highly conserved in vertebrates. Due to the high genetic conservation of these pathways, human drugs can be used to block specific biochemical steps to determine the roles that cholesterol and protein lipidation play in zebrafish development. Reprinted from Development Cell with permission from Elsevier (Thorpe et al., 2004).

statins, either mevinolin (Lovastatin) or simvastatin (Zocor), exhibit developmental arrest, blunted axis elongation, misshapen somites, and head and axial necrosis (Fig. 5). Additionally, PGC migration is profoundly perturbed. Embryos treated with a more hydrophobic statin (Lipitor) exhibit PGC migration defects and only mild morphologic abnormalities. To determine which downstream products of HMGCoAR mediate these developmental defects, a ‘‘block and rescue’’ approach was taken through injection of putative biochemical pathway intermediates downstream of HMGCoAR following statin-mediated inhibition. Injection of mevalonate, the product of HMGCoAR’s reduction of ß-hydroxy-ß-methylglutaryl-CoA, completely rescued all the phenotypes associated with the statin treatment (Fig. 6). Similar experiments using intermediates downstream of mevalonate (Fig. 7) indicated that the prenylation pathway, responsible for adding polyunsaturated lipids to proteins, was likely mediating the effect of statins on PGC migration. High doses of the selective farnesyl transferase inhibitors L-744 or FTI-2153 (Crespo et al., 2001; Sun et al., 1999) had no effect on PGC migration. However, embryos treated with geranylgeranyl transferase I (GGTI) inhibitor (GGTI-2166) exhibited a strong PGC migration phenotype with only a

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Fig. 5

Embryos treated with statins exhibit developmental defects. In comparison to untreated embryos (A), embryos soaked in a low dose of mevinolin (0.06 mM). (B) exhibit mild developmental defects, as evidenced by tail kinks. (C) Exposure to higher doses of mevinolin (1.2 mM) results in blunted axis elongation, necrosis and developmental arrest. (D) Simvastatin treated embryos (2.0 mM) exhibit similar developmental defects. (E) Dose response of mevinolin on developmental arrest (mean + SEM, n = 3). (F) Dose response of simvastatin on developmental arrest (mean + SEM, n = 3). Reprinted from Development Cell with permission from Elsevier (Thorpe et al., 2004).

slight disruption of the notochord. These data suggest that protein prenylation, specifically by GGTI, is required for correct PGC migration (Thorpe et al., 2004).

V. Visualizing Lipid Metabolism in Larval and Adult Zebrafish A. Lipophilic Dyes Lipophilic dyes, known as lysochromes, are one of the first tools used to visualize lipids in cells. These dyes are capable of labeling a variety of lipids and lipidcontaining structures including TAG, FA, and lipoproteins. Dyes, such as oil red O (ORO), sudan black B, and nile red, were initially used to label LDs in tissue sections and cultured cells and continue to be used today. Marza and colleagues (Marza et al., 2005) utilized sudan black B to identify LDs in histological sections of fed adult zebrafish. The authors found that feeding a high-fat meal increased the expression of MTP in intestinal epithelial cells. This protein is required for proper


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Fig. 6 Isoprenoid intermediates rescue the developmental defects caused by statin treatment. Embryos at early cell stages were injected with mevalonate and then soaked overnight in mevinolin, simvastatin, or atorvastatin. (A) Embryos treated with mevinolin show severe developmental defects at 24 hpf. (B) Embryos injected with the isoprenoid intermediate mevalonate (1–16 cell stages) and then treated with mevinolin exhibit normal morphology. (C) Mevalonate injection rescues the somatic defects observed in embryos treated with different statins. Embryo morphology was scored at 24 hpf. Data represent the MEAN SEM from 3–4 experiments. Reprinted from Development Cell with permission from Elsevier (Thorpe et al., 2004).

assembly and secretion of hepatic and intestinal ApoB-containing lipoproteins, chylomicrons, and very low-density lipoproteins (VLDL) (Gordon et al., 1995). Their observation that LDs are coincident with an upregulation of MTP expression is consistent with MTP’s known function in humans (Marza et al., 2005). Lysochromes can also be used to visualize endogenous lipid stores in whole fixed zebrafish to generate an overall picture of neutral lipid localization during development. Schlegel and Stainier used ORO to assess the consequence of MTP knockdown (via MO) on lipid absorption in whole zebrafish larvae (Schlegel and Stainier, 2006). MTP morphants exhibited decreased yolk consumption and an inability to absorb dietary neutral lipids, resulting in death by 6 dpf. Although lysochromes such as sudan black B and ORO consistently label neutral lipids in tissue sections and fixed larvae, fixation techniques are laborious and staining procedures have been shown to cause artificial fusion of adjacent LDs and mislocalization of the LD marker, adipose differentiation-related protein (Adrp/Perilipin2) (Fukumoto and Fujimoto, 2002). More recent techniques to visualize LDs have focused on staining these drops in vivo.

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Fig. 7 Statin treatment causes abnormal primordial germ cell (PGC) migration in zebrafish embryos. Compared to wild-type (A), embryos treated with statins (B) display ectopic PGCs. The arrows indicate ectopic PGCs that have failed to migrate to the developing gonad. (C) The PGC migratory defect observed following statin treatment is prevented by injections of isoprenoid intermediates. Embryos at early stages (1–16 cell) were injected with gfp-nos mRNA and mevalonate and then soaked overnight in mevinolin, simvastatin or atorvastatin. At 24 hpf, embryos were scored for ectopic PGCs, with a score of 1 indicating a wild-type single gonadal cluster and score of 4 indicating no discernable PGC cluster. Data represent the mean SEM from 3–4 experiments. (D) The PGC migratory defect observed following statin treatment is prevented by increasing the levels of isoprenoid synthesis intermediates. Embryos injected at the 1–16 cell stage with farnesol, geranylgeraniol or mevalonate and then soaked overnight with atorvastatin (10 mM) show normal PGC migration. Data represents MEAN SEM, *p < 0.01 difference from atorvastatin alone. Reprinted from Development Cell with permission from Elsevier (Thorpe et al., 2004).


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Greenspan et al. (1985) first showed the utility of nile red (9-diethylamino-5Hbenzo[a]phenoxazine-5-one) to label intracellular LD in live cultured peritoneal macrophages and smooth muscle cells. Nile red is an uncharged heterocyclic molecule that only fluoresces in a hydrophobic environment. Labeled neutral lipids fluoresce a yellow-gold to red color, depending on their relative hydrophobicity, with no detectable damage or deformation of dye-infused tissues (Fowler and Greenspan, 1985). More recently, Jones et al. (2008) used nile red to visualize neutral lipid deposits in live zebrafish larvae. They initially demonstrated that daily exposure of larvae to nile red-containing embryo media for 4 days (from 3 to 7 dpf) consistently labeled lipid-rich tissues. The authors then sought to test the effects of known pharmacological inhibitors of triglyceride metabolism on total larval lipid content. Treatment with nicotinic acid, a potent pharmacological inhibitor of adipocyte lipolysis (Carlson, 1963), resulted in an increase in total triglyceride content and decreased cholesterol levels. Treatment with resveratrol, a compound known to inhibit FA synthase (Tian, 2006), resulted in a decrease in total triglyceride content as detected by nile red staining. Total triglyceride content was further decreased when resveratrol was supplemented with norepinephrine (Jones et al., 2008). While fluorescent dyes and stains are useful for identifying lipid deposits in cells and tissues, issues arise regarding the distribution and affinity properties of these compounds. Nonspecific labeling of tissues devoid of lipid deposits may be observed and staining and washing procedures must then be carefully optimized to minimize this effect. Additionally, nile red staining does not distinguish between FA and cholesterol in vivo, although some discrimination based on staining intensity of tissue sections is possible (Fowler and Greenspan, 1985). B. BODIPY Fatty Acid Analogs The wide variety of fluorescent lipid analogs commercially available allows one to fully exploit the optical clarity of zebrafish larvae to study lipid metabolism. One type of analog widely used in cultured cells to visualize lipid dynamics is BODIPYlabeled FA. These analogs consist of an acyl chain of variable length attached to the BODIPY (4,4-difluoro-4-bora-3a, 4a-diaza-S-indacene) fluorescent moiety. First synthesized by Treibs and Kreuzer in 1968 (Treibs and Kreuzer, 1969), the BODIPY fluorophore possesses a number of advantageous qualities including high photostability, strong and narrow wavelength emission in the visible spectrum, and an overall uncharged state (Monsma et al., 1989; Pagano et al., 1991). To administer BODIPY FA analogs to live zebrafish larvae, we developed a feeding assay that utilizes liposomes to create a suspension of relatively hydrophobic FA analogs in embryo media (Carten, unpublished). Following a short liposome feed, digestive organ structure and metabolic function can be assessed, as the fluorescent FAs accumulate readily throughout numerous larval organs and tissues. With this assay, we have observed that different chain length FAs (short, medium, and long) accumulate in distinct patterns throughout digestive organs and tissues, with each chain length suited to visualize particular larval organs and cellular structures. Long

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chain fatty acids (LCFA) appear in cytoplasmic LD in enterocytes and hepatocytes. Short chain fatty acids (SCFA) accumulate primarily in the hepatic and pancreatic ducts and are particularly suited to illuminate ductal networks. Medium chain fatty acids (MCFA) accumulate in LDs throughout a wide range of cell types, as well as ductal and arterial networks, and reveal the subcellular structures of multiple cell types. Because this feeding assay enables the rapid assessment of digestive function and FA metabolism in live zebrafish larvae, it has the potential for use in genetic and pharmacologic screens to identify genes involved in FA metabolism and potential drug targets.

C. BODIPY Cholesterol In addition to FA analogs, a number of sterol analogs are available for use with cultured cells and zebrafish larvae (e.g., NBD-cholesterol, BODIPY-cholesterol). Initially created to visualize cholesterol partitioning into membranes, these analogs were found to preferentially enter into liquid-disordered domains, making them less useful for studies of sterol trafficking in cells (Li et al., 2006). To address these limitations a new BODIPY-tagged cholesterol analog was synthesized with a modified fluorophore linker (Li and Bittman, 2007). Recent studies utilizing the improved BODIPY-cholesterol analog found it to partition into the cholesterol-rich liquid-ordered membrane domain (Ariola et al., 2009) and interact with membranes in ways similar to native sterols, making it a powerful new tool for imaging sterol trafficking in live cells (Marks et al., 2008). Studies done in the zebrafish have found that BODIPY-cholesterol localizes to the yolk of developing zebrafish larvae (Holtta-Vuori et al., 2008). Ongoing work in the Farber lab is examining the localization of BODIPY-cholesterol in zebrafish larval intestinal enterocytes after a high-fat meal and comparing its localization to that of LD (revealed by BODIPY-labeled FA). Recent data suggest that the initial uptake of sterol and of FA segregate into nonoverlapping compartments (Walters, unpublished). These types of experiments will ultimately enable the development of a clearer model for the uptake and trafficking of dietary lipid in intestinal enterocytes.

D. Fluorescent Reporters of Lipid Metabolism The ability to perform forward genetic studies in zebrafish by mutagenizing the entire genome and screening for particular phenotypes has made this vertebrate model popular (Driever et al., 1996; Haffter et al., 1996). Mutagenesis methods commonly utilized by the zebrafish community include gamma-ray irradiation to generate large deletions and translocations (Fisher et al., 1997), soaking founder fish in mutagenic chemicals such as ethylnitrosourea (ENU) to generate point mutations, (Driever et al., 1996; Haffter et al., 1996), and retrovirus- or transposon-mediated gene insertions (Chen and Farese, 2002; Ivics et al., 2004). We perform an ongoing


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ENU-mutagenesis screen to search for mutations that perturb lipid processing. To screen for mutants defective in lipid processing, we soak zebrafish larvae in fluorescent lipid reporters that are swallowed and allow lipid processing to be visualized in vivo (Farber et al., 2001). One fluorescent reporter we routinely use in our screen is the phosphoethanolamine analog PED6, [N-((6-(2,4-dinitro-phenyl)amino)hexanoyl)-1-palmitoyl-2BODIPY-FL-pentanoyl-sn-glycerol-3-phosphoethanolamine] (Fig. 8A). Following ingestion, PED6 is cleaved by phospholipase A2 (PLA2), resulting in the release of the fluorescent labeled acyl chain (Farber et al., 1999) (Fig. 8C). When zebrafish larvae (5 dpf) are immersed in media containing PED6, bright green fluorescence is observed in the intestine, gall bladder, and liver (Fig. 8E). We also utilize the sterol analog 22-NBD-cholesterol (22-[N-(7-nitronbenz-2-oxa-1,3-diazol-4-yl) amino]23,24-bisnor-5-cholen-3-ol) (Fig. 8B) to visualize cholesterol absorption in live larvae and screen for mutants (Fig. 8F). This reagent is different from PED6 in that it continuously fluoresces and due to its hydrophobicity it is slightly more difficult to administer via feeding. Because PED6 and NBD-cholesterol provide rapid readouts for digestive organ morphology and lipid processing, we used them to perform the first forward genetic physiologic screen in zebrafish to identify new genes that regulate lipid metabolism. One mutation identified with these lipid reporters, fat-free (ffr), was a recessive lethal mutation. Although ffr mutants appeared morphologically normal, they exhibited significantly diminished fluorescence in their intestine and the gall bladder following PED6 treatment (Fig. 9) (Farber et al., 2001). ffr larvae were further characterized using NBD-cholesterol to determine if the uptake and trafficking of sterol-like molecules was also impaired. In contrast to wild-type larvae, which accumulated NBD-cholesterol in their gall bladders within a few hours of feeding, ffr mutants were unable to concentrate NBD-cholesterol in their gall bladders. This observation suggests that ffr larvae have a significant defect in bile secretion and/or transport. When ffr mutants were incubated with BODIPY FL-C5, a medium chain FA analog, they had nearly normal digestive organ fluorescence. Because this medium chain length FA is less hydrophobic and is not dependent on emulsifiers, such as bile, for absorption, this further suggests that the ffr mutation attenuates biliary synthesis or secretion (Ho et al., 2003). To better characterize the metabolic defects of ffr larvae, we immersed mutants in media containing radioactive oleic acid (250 mCi/mmol, 3 h), followed by whole embryo lipid extraction and TLC analysis. We found that ffr larvae have significantly reduced radioactivity incorporated into phosphatidylcholine (PC) fraction (p95%, which is similar to murine resections, and the high conservation with mammalian models allows the rapid assessment of chemical and genetic modulation in organ injury and repair. Protocol – Liver Resection 1. Anesthetize adult zebrafish with tricaine. 2. Transfer anesthetized fish to lid of 10 cm culture dish, located under dissection stereoscope. 3. Using McPHERSON-VANNAS microdissecting spring scissors (Biomedical Research Instruments 11–1050) and #55 forceps, make an 3 mm incision on the ventral abdomen, just caudal to opercula. 4. Remove the inferior liver lobe, using gentle tugging motion. The zebrafish liver has no capsule and can easily tear. Use caution not to injure the intestine as this will result in higher mortality. 5. No wound closure is required after the completion of the liver resection because the scales will cover over the resection site. 6. Return fish to fresh zebrafish water. (Note: Care should be taken to complete the procedure in a timely manner; if the fish does not exhibit gill movement after prolonged anesthesia, then manual perfusion of the gills with a transfer pipette may accelerate recovery.) 7. Sacrifice zebrafish at defined time interval after resection with approved methods, such as tricaine overdose. Either process entire fish for histological evaluation or dissect out endodermal organs for length measurements of inferior lobe and remnant stump (see schematic in Fig. 1). C. Genetic Ablation Another method for targeted and timed hepatocyte injury was recently introduced by Curado et al. (2007). This approach has also been demonstrated in other cell types, including pancreatic beta cells, cardiomyocytes, retina, and testis. Here, transgenic fish are created that express the Escherichia coli enzyme nitroreductase under the control of organ-specific promoters, in this case fabp10a (Tg(fabp10: CFP-NTR)s891). Although nitroreductase expression has no effect on the target organ by itself, its enzymatic activity reduces metronidazole (1-(2-hydroxyethyl)2-methyl-5-nitroimidazole), a commonly used and typically nontoxic antibiotic, to form a potent DNA interstrand cross-linking agent, which causes cell death. This technology can be applied to fish of all ages: embryos, larvae, juveniles, and adults.

10. Endoderm Specification, Liver Development, and Regeneration

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Fig. 1 Schematic depiction of zebrafish liver resection. The inferior liver lobe is resected. Regenerative capacity can be analyzed after en bloc dissection of the endodermal organs and measurement of the total inferior lobe length (indicated in black) and the length of the remnant liver stump (indicated in grey, often identified by blood clots). The ratio of total:remnant lengths indicates the regenerative index.

In this model, toxicity and cell death are exclusively limited to the nitroreductaseexpressing cells, potentially allowing highly targeted cellular injury. However, this also makes this system extremely dependent on a suitable promoter, efficient transgenic expression, and availability of metronidazole to the cells to achieve significant damage of the entire organ (Curado et al., 2008) and may not allow complete ablation of hepatic tissue (Curado et al., 2010).

D. Transient Genetically Induced Apoptosis Another model of hepatic recovery was also introduced by Curado and colleagues (2010). This approach is based on the observation of a novel genetic mutant, oliver, with a mutation of the mitochondrial protein translocase of the outer mitochondrial membrane 22 homolog (tomm22) gene. These mutants exhibit a smaller liver consisting of mainly biliary tissue caused by hepatocyte apoptosis. Transient knockdown of tomm22 by morpholino injection results initially in a corresponding phenotype; however, with waning efficacy of the morpholino, the liver recovers. The morphant provides a model for organ recovery after apoptotic cell death in larval stages where the role and impact of other signaling pathways can be tested, which has been done for WNT and FGF. This model may be amenable to chemical modifier screens on a larger scale.

IV. Assessment of Liver Function In recent years, several approaches have been developed to assess clinical and functional parameters of hepatic physiology and damage in zebrafish. These

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methods are identical to or reflect the assessment of liver damage and function obtained in patients suspected of having liver disease. These measures enable the correlation of zebrafish phenotypes with clinical liver disease.

A. Liver Enzymes Liver enzyme tests are the cornerstone of the clinical assessment of hepatobiliary damage. The assessment of serum parameters in the adult zebrafish has been limited due to the small volume of blood that can be obtained (Murtha et al., 2003). By direct cardiac puncture using micropipettes, however, blood samples can be pooled to obtain minimum quantities (50 ml) of serum for analysis (100 ml whole blood). This can be processed in a clinical laboratory to obtain ‘‘individual’’ cohort test values. We were able to demonstrate APAP-induced liver damage in adult fish by an increase in circulating alanine aminotransferase levels (ALT) (North et al., 2010). Other relevant parameters are the assessment of aspartate aminotransferase for hepatocyte damage, alkaline phosphatase for biliary injury, and total bilirubin for the metabolic capacity of the liver. In humans, albumin is routinely measured to assess protein synthesis in the liver; however, recent genomic (Noel et al., 2010) and proteomic analyses (North et al., 2010) demonstrate the absence of albumin in the zebrafish. Overall, this method is resource-intensive, because 10–25 fish are required to obtain one pooled sample sufficient for analysis. Despite this, it can serve as a resource when demonstrating the clinical importance of zebrafish studies, using therapeutically relevant parameters.

B. Lipid Metabolism In addition to the evaluation of early liver morphology and its regenerative capacity, recent studies have demonstrated the feasibility of evaluating functional aspects of the intestine, pancreas, and liver in vivo. One technique is intravital staining with Nile red which, like oil-red-O in histological sections, stains neutral lipids, which could be used in small molecule screens to discover novel antiobesity drugs (Jones et al., 2008). Although Nile red stains all lipid-rich tissues, it cannot distinguish between cholesterol and fatty acids. The development of BODIPY fluorophores has enabled the dynamic assessment of lipid metabolism in vivo. One compound, N-([6-(2,4-dinitro-phenyl)amino]hexanoyl)-1-palmitoyl-2BODIPY-FL-pentanoyl-sn-glycerol-3-phosphoethanolamine (PED6), has recently been used to demonstrate lipase activity in vivo (Farber et al., 2001). Zebrafish larvae exposed to PED6 exhibit green fluorescence in the intestine, gall bladder, and liver. This approach has enabled genetic screens to identify modulators of lipid metabolism, such as in the mutant fat-free (c11orf2, also called ffr) (Farber et al., 2001) and others in medaka (Watanabe et al., 2004). More recently, PED6 has been combined with a quenched protease reporter, EnzChek (Invitrogen), which can assess exocrine pancreas function, and with microspheres, which determine


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swallowing and intestinal capacity, giving a comprehensive functional in vivo assessment of gastrointestinal physiology (Hama et al., 2009).

V. Summary Over the past decade, many new aspects of early endoderm formation, liver specification, differentiation, and growth have been discovered in the zebrafish. In zebrafish the liver does not function as a hematopoietic organ; therefore, genetic and chemical effects that lead to impaired liver formation can be studied without the interference of death from anemia. This has revealed the interaction of central signaling pathways in liver formation, and a direct assessment of their function in vivo. In addition, the study of larval and adult zebrafish has enabled the development of models of liver injury and regeneration as well as in vivo assessment of organ function, with a high conservation between fish and humans of toxic and therapeutic drug effects. These findings indicate the potential of the zebrafish for discovery of disease-relevant aspects of endoderm and liver structure and function. References Alexa, K., Choe, S. K., Hirsch, N., Etheridge, L., Laver, E., Sagerstrom, C. G. (2009). Maternal and zygotic aldh1a2 activity is required for pancreas development in zebrafish. PLoS One 4, e8261. Alexander, J., Rothenberg, M., Henry, G. L., and Stainier, D. Y. (1999). casanova plays an early and essential role in endoderm formation in zebrafish. Dev. Biol. 215, 343–357. Amali, A. A., Rekha, R. D., Lin, C. J., Wang, W. L., Gong, H. Y., Her, G. M., Wu, J. L. (2006). Thioacetamide induced liver damage in zebrafish embryo as a disease model for steatohepatitis. J. Biomed. Sci. 13, 225–232. Anderson, R. M., Bosch, J. A., Goll, M. G., Hesselson, D., Dong, P. D., Shin, D., Chi, N. C., Shin, C. H., Schlegel, A., Halpern, M., et al. (2009). Loss of Dnmt1 catalytic activity reveals multiple roles for DNA methylation during pancreas development and regeneration. Dev. Biol. 334, 213–223. Aoki, T. O., David, N. B., Minchiotti, G., Saint-Etienne, L., Dickmeis, T., Persico, G. M., Strahle, U., Mourrain, P., Rosa, F. M. (2002). Molecular integration of casanova in the Nodal signalling pathway controlling endoderm formation. Development 129, 275–286. Bally-Cuif, L., Goutel, C., Wassef, M., Wurst, W., and Rosa, F. (2000). Coregulation of anterior and posterior mesendodermal development by a hairy-related transcriptional repressor. Genes Dev. 14, 1664–1677. Brown, J. L., Snir, M., Noushmehr, H., Kirby, M., Hong, S. K., Elkahloun, A. G., Feldman, B. (2008). Transcriptional profiling of endogenous germ layer precursor cells identifies dusp4 as an essential gene in zebrafish endoderm specification. Proc. Natl. Acad. Sci. U.S.A. 105, 12337–12342. Burdine, R. D., and Schier, A. F. (2000). Conserved and divergent mechanisms in left-right axis formation. Genes Dev. 14, 763–776. Chen, W., Burgess, S., and Hopkins, N. (2001). Analysis of the zebrafish smoothened mutant reveals conserved and divergent functions of hedgehog activity. Development 128, 2385–2396. Chung, W. S., Shin, C. H., and Stainier, D. Y. (2008). Bmp2 signaling regulates the hepatic versus pancreatic fate decision. Dev. Cell 15, 738–748. Curado, S., Anderson, R. M., Jungblut, B., Mumm, J., Schroeter, E., Stainier, D. Y. (2007). Conditional targeted cell ablation in zebrafish: a new tool for regeneration studies. Dev. Dyn. 236, 1025–1035.

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CHAPTER 11

Morphogenesis of the Zebrafish Jaw: Development Beyond the Embryo Kevin J. Parsons,* Viktoria Andreeva,y W. James Cooper,* Pamela C. Yelicky and R. Craig Albertson* *

Department of Biology, Syracuse University, Syracuse NY, Tufts University, Boston, Massachusetts

y

Department of Oral and Maxillofacial Pathology, Tufts University, Boston, Massachusetts

Abstract I. Postembryonic Development – Framing the Questions and Understanding the Challenges II. Obtaining Phenotypes A. Postembryonic Mutagenesis Screens in Zebrafish B. Heterozygous Craniofacial Phenotypes in Previously Identified Homozygous Recessive Early Lethal Zebrafish Mutants C. Rescue of Homozygous Recessive Lethal Mutations to Examine Postembryonic Defects III. Quantitative Methods for Studying Adult Phenotypes A. Geometric Morphometrics B. Data Collection C. General Data Analysis D. Analysis of Shape E. Analysis of Shape Variation F. Rates of Shape Change G. Trajectories of Shape in Zebrafish Lines H. Morphological Integration I. Synthesis of Quantitative Data IV. A Complementary Approach: The Use of Natural Variation to Complement that Generated in the Lab for Understanding Jaw Morphogenesis V. Implications and Conclusions References

Abstract The zebrafish has emerged as an important model for vertebrate development as it relates to human health and disease. Work in this system has provided significant METHODS IN CELL BIOLOGY, VOL 101 Copyright 2011. Elsevier Inc. All rights reserved.

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insights into the variety of genetic signals that direct the cellular activities and tissue interactions necessary for proper assembly of the pharyngeal skeleton. Unfortunately our understanding of craniofacial development beyond embryonic stages is far less complete. Stated another way, we know a great deal about the early patterning of the skull, but we know comparatively little about how mature craniofacial shape is determined and maintained over time. Here we propose ways to expand the current molecular genetic paradigm beyond the embryo to gain an understanding of the processes and mechanisms that guide growth and remodeling of mineralized craniofacial, skeletal, and dental tissues. First, we discuss sources of adult mutant phenotypes that can be used to study of postembryonic development. Next, we review salient quantitative methods that are necessary to define complex adult phenotypes. We also discuss how other organismal systems can be used to inform and complement studies in zebrafish. We conclude by discussing the implications for such studies within the context of furthering an understanding of the etiology and pathophysiology of human craniofacial malformations, as well as informing an understanding of adaptive craniofacial variation among natural populations.

I. Postembryonic Development – Framing the Questions and Understanding the Challenges In stark contrast to the extensive body of literature focused on understanding the basis of early craniofacial patterning, relatively little is known about the mechanisms that underlie skeletal development beyond embryonic stages. Craniofacial development can be broadly divided into chronological stages based on the appearance of tissues, the formation of structures, and the initiation of various morphogenic processes. The cranial neural crest (CNC) cells are the first to arise during embryonic development that directly contribute to the craniofacial skeleton. A PubMed literature search (May 12, 2010) for CNC identified nearly 1400 peer-reviewed research articles. By early larval stages of development the CNC cells condense and differentiate to form craniofacial cartilages, which soon thereafter develop into pharyngeal bones and other components of the skull (Hall, 1999). A search for craniofacial ‘‘cartilage development’’ returned over 600 articles, whereas craniofacial ‘‘bone development’’ identified 200 references. Bone growth and remodeling are initiated during larval development, and continue throughout juvenile and adult stages. A PubMed search for craniofacial ‘‘bone growth’’ and ‘‘bone remodeling’’ each identified fewer than 100 articles. This brief survey of the literature reveals a dearth of peer-reviewed articles that explore craniofacial development beyond embryonic stages. It also reveals an inverse relationship between developmental chronology and our understanding of the mechanisms that regulate this process. The main purpose of this chapter is to describe ways in which the experimentally tractable zebrafish can be used to promote a more robust understanding of craniofacial development beyond the embryo.

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The zebrafish is an excellent vertebrate model with which we study skeletogenesis at all stages of development. They are easily bred and maintained, have short generation times, and large numbers of progeny can be obtained from a single mating (200–500/week). The zebrafish genome has been sequenced and zebrafish developmental genetics are easily manipulated through functional analyses, transplantation, and transgenics. Zebrafish are also touted for having transparent embryos, allowing various developmental processes to be observed in the living embryo with minimal processing or manipulation. This attribute also holds true for postembryonic zebrafish, particularly for the skeletal system. Most craniofacial elements lie just underneath a relatively thin layer of tissue and are easily observed in live or fixed animals with minimal processing (Fig. 1). Another relevant attribute of the zebrafish is that, like most teleosts, they exhibit dynamic patterns of allometric growth, whereby anatomical changes accompany their increase in size (Fuiman, 1983; Hernandez, 2000; Osse, 1990; Weatherly, 1990). The life cycles of bony fishes are often characterized by dramatic shifts in ecology and behavior, accompanied by concomitant changes in the body shape (Loy et al., 2001; Zelditch and Fink, 1995). Allometric growth is even considered to be a defining characteristic of fish early life histories (Hernandez, 2000). The use of a model that exhibits pronounced size-related shape change will likely facilitate our efforts to understand the mechanisms that underlie skeletal growth and remodeling. Zebrafish craniofacial mutants have traditionally been characterized using qualitative methods (i.e., the gross presence or absence of structures) up to, and rarely past, 5 days postfertilization (dpf). We have argued previously (Albertson and Yelick, 2004, 2007; Cooper and Albertson, 2008) that a shift from qualitative to quantitative descriptions of phenotype will facilitate a more comprehensive understanding of craniofacial development, particularly as it pertains to development beyond larval stages. The impetus for this assertion stems, in part, from the fact that the adult skeleton is significantly more complex than that of the larvae. Fig. 2A depicts a 5-day larval skeleton, while Fig. 2B shows the adult complex. The most obvious

[(Fig._1)TD$IG]

Fig. 1 Skeletal preparations in two adult zebrafish. (A) A living fish stained with the fluorescent compound, Calcein (Sigma), which binds to free calcium (note the cranial sutures). (B) An adult AB zebrafish that was fixed, enzymatically cleared, and stained with Alizarin red that also binds to calcium (Sigma, St. Louis, MO). In both specimens detailed skeletal anatomy is apparent.

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[(Fig._2)TD$IG]

Fig. 2 Ontogenetic changes in craniofacial bone in AB zebrafish. (A) depicts a 5-day larval skeleton, while (B) shows the adult complex. The most obvious difference between the two structures is that the larval skeleton is only barely ossified as indicated by the predominance of alcian blue staining.

difference between the two structures is that the larval skeleton is minimally ossified. For the most part, only cartilages are present, and thus only a fraction of the elements that comprise the adult complex is evident. In fact, entire functional units are absent from the larval form, including the upper jaws and portions of the cranium (Harris et al., 2008). So while significant progress has been made toward an understanding of the molecular/genetic factors that regulate development of larval pharyngeal cartilage morphology, we know virtually nothing about major functional complexes of the skeleton that appear later in development (i.e., mineralized jaws). We contend that a major challenge facing developmental biologists is to extend the current developmental genetic paradigm beyond the embryo. This challenge can be framed by asking two basic questions: (1) What are the molecular determinants of craniofacial development beyond embryonic stages? and (2) What is the molecular basis for quantitative shifts in craniofacial shape? The answer to these and related questions will inform a broad range of scientific disciplines, as we seek a better understanding of both normal and abnormal variation in craniofacial form.

II. Obtaining Phenotypes A. Postembryonic Mutagenesis Screens in Zebrafish Over the years, the zebrafish has proven to be an excellent subject for forwardgenetic mutagenesis screens. By screening for early-lethal phenotypes, a large number of mutants relevant to human development and disease have been discovered, including a variety of craniofacial mutants (Neuhauss et al., 1996; Piotrowski et al., 1996; Schilling et al., 1996). These studies have shed significant light on our understanding of molecular events regulating early craniofacial and tooth development. Our current focus is to move beyond the embryonic stages in order to identify and characterize genes and signaling pathways that regulate the growth and


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remodeling of adult mineralized tissues. Studies of viable, adult mineralized craniofacial tissue phenotypes are particularly relevant to determining the genetic basis of human craniofacial pathophysiology and disease. Several recent publications have demonstrated the strengths of the zebrafish model for adult craniofacial and skeletal morphogenesis. For example, Fisher et al. (2003) utilized a radiographic analysis of adult ENU-mutagenized F1 fish to identify the dominant skeletal dysplasia chihuahua (chi) mutant. The chi mutant zebrafish harbors a dominant mutation in the type I collagen gene (col1a1) which leads to uneven mineralization, fragility and defects in bone growth (Fisher et al., 2003), features that are associated with human skeletal dysplasia osteogenesis imperfecta (OI). OI is a dominant heritable disorder of connective tissues in humans, caused in approximately 90% of individuals by mutations in type I collagen genes (col1a1 and col1a2), for which there currently is no effective clinical treatment (reviewed in Basel and Steiner, 2009). Thus, the zebrafish chi mutant is a valuable tool and model with which we can extend our knowledge of OI, and hopefully identify effective clinical treatments. Another large-scale mutagenesis screen for defects in adult structures led to the discovery of two similar mutants finless (fls) and Nackt (Nkt), both of which exhibited defects in the skeletal elements of the skull, fins, scales, and teeth (Harris et al., 2008). It has been shown that Nkt harbors a mutation in the ectodysplasin (eda) gene,and that mutations of the ectodysplasin receptor (edar) gene were found in fls mutants (Harris et al., 2008). Remarkably, mutations of eda and edar genes are responsible for the majority of hypohydrotic ectodermal dysplasia (HED) cases in humans (Mikkola, 2009). HED is a hereditary disorder characterized by hair and teeth defects, as well as defects in a number of other ectodermal organs (Mikkola, 2009). In the Yelick laboratory, we have identified several mutants that exhibit craniofacial and bone defects in adult fish, by applying high-throughput Alizarin red (AR)

[(Fig._3)TD$IG]

Fig. 3 Analysis of bka and knz mutants. (A) AR staining of bka mutant and wild-type control. (B) GM analysis of bka mutant and wild-type control. The arrows in (A) and (B) point to reduced upper jaw in bka mutant. (C) AR staining of knz mutant and wild-type control. (D) GM analysis of knz mutant and wildtype control. The arrows in (C) and (D) point to changes in upper jaw morphology of knz mutant.

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staining of ENU-mutagenized F3 generations at 6–8 weeks postfertilization (Stewart-Swift et al., 2010; 9th International Zebrafish Development and Genetics Conference, Madison, WI, June16–20, 2010). One of them, homozygous recessive mutant belkal (38N), exhibits a midface hypoplasia phenotype, and reduced and fused maxilla (Fig. 3A, B). In addition, approximately 50% of sqr mutants develop scoliosis. A second mutant, which seems to be heterozygous semidominant, knjaz’ (78N), displays craniofacial shape defects including the shape of preorbital region possibly due to a caudal displacement of the upper jaw in addition to scoliosis in the tail region, as detected by AR staining, and as confirmed by geometric morphometric (GM) analyses, the methodology for this will be explained in the following (Fig. 3C, D). Efforts to map each of these mutations, and to identify the genetic mutations responsible for these phenotypes, are currently underway. B. Heterozygous Craniofacial Phenotypes in Previously Identified Homozygous Recessive Early Lethal Zebrafish Mutants Another approach to identify adult craniofacial and bone mutants is to use AR staining and GM analyses to look for heterozygous phenotypes previously described early-lethal homozygous recessive zebrafish mutants – particularly those harboring gene mutations in craniofacial and tooth-expressed genes. One example is the acerebellar (ace) mutant, caused by a mutation in the fibroblast growth factor (fgf8) gene. The ace/fgf8 homozyougous mutants exhibit an asymmetric craniofacial pharyngeal skeleton, among other defects (Albertson and Yelick, 2005), and die at approximately 7 dpf (Brand et al., 1996). Our study of heterozygous ace/fgf8 zebrafish allowed investigations of craniofacial development to be extended beyond the early embryonic stages, and identified roles for fgf8 in adult craniofacial form and function (Albertson and Yelick, 2007). Our geometric shape GM analyses revealed that fgf8 haploinsufficency leads to craniofacial asymmetries and defects in cranial sutures, and staining for alkaline phosphatase and tartrate resistant acid phosphatase activities showed aberrant osteoblast and osteoclast activities, respectively, which contributed to abnormal bone formation and remodeling in the lower jaw (Albertson and Yelick, 2007). Thus, these studies demonstrated that subtle but highly informative phenotypes can be revealed by investigations of viable adult heterozygous mutant zebrafish. Analyses of the viable adult heterozygous zebrafish mutants have high clinical relevance, since the majority of human hereditary disorders are induced by mutations in a single copy of the gene. For instance, single allele mutations in fibroblast growth factor receptors (fgfr)types 1–3, have been found in several human syndromes that are characterized by craniofacial defects including craniosynostosis, or premature fusion of cranial sutures, as well as defects in bone formation and growth (Morriss-Kay and Wilkie, 2005). C. Rescue of Homozygous Recessive Lethal Mutations to Examine Postembryonic Defects An additional way to extend our knowledge of craniofacial morphogenesis beyond embryonic stages is by rescuing homozygous recessive mutants from early lethality


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using wild-type mRNA injections. This approach was successfully used by LeClair et al. (2009) to discover the effects of glypican 4 (gpc4) on craniofacial organization in adult zebrafish (LeClair et al. (2009). gpc4 belongs to the family of extracellular glycoproteins that can modulate Wnt, bmp, and fgf signaling (Filmus and Selleck, 2001; Fransson, 2003). The gpc4/knypek homozygous mutant embryos normally die around 5–7 dpf, and exhibit defects in convergence and extension movements in the ectoderm and mesoderm (Topczewski et al., 2001). However, when investigators rescued gpc4/knypek homozygous mutant embryos by single cell injections of gpc4 mRNA, and then allowed the rescued mutants to develop for up to 1 year, they discovered the loss or rearrangement of several adult craniofacial bones, defects in craniofacial cartilages, and changes in the shape of neurocranium (LeClair et al., 2009). These elegant studies have shown that adult craniofacial phenotypes can be identified and examined in previously identified homozygous recessive lethal mutants, if they are allowed to survive beyond critical early developmental stages.

III. Quantitative Methods for Studying Adult Phenotypes A. Geometric Morphometrics Shape is a fundamentally important feature of organisms but has historically been very challenging for biologists to quantify. Until recently morphometric analyses have largely involved measuring sets of linear distances on a form, applying a chosen method of size correction to these distances, and finally applying a multivariate statistical test to these data to discern patterns. While these methods are still used, they are fraught with disagreements over what size corrections to apply, and difficulties in determining the biological meaning of statistical results (Parsons et al., 2003). The field of morphometrics is undergoing a radical transformation through the introduction and expansion of GM (Adams et al., 2004; Rohlf and Marcus, 1993). These powerful techniques have now become the standard method to quantify the shape and offer researchers a greater ability to discern and interpret trends in shape variation. Simply put, we contend that morphometrics is a quantitative approach to compare shape differences that zebrafish biologists have been describing qualitatively for years. Moreover, these newer methods are powerful enough that they can extract meaningful information from situations where only subtle or continuous variation occurs, even when these patterns are not apparent to the naked eye. Although the use of GM in zebrafish research is still in its infancy, there is enormous potential for biologists to adopt these methods to study development beyond the embryo (Albertson et al., 2005; Albertson and Yelick, 2007; Cooper and Albertson, 2008; LeClair et al., 2009). We recognize that GM can seem mathematically complex, abstract, or even obscure, and that this perception could easily lead to discouragement. Accordingly, we have written the following sections as a primer for zebrafish biologists, with largely heuristic explanations of the mathematics

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and statistics being applied. We will also illustrate how data from GM can be collected and analyzed in a developmental context using samples from the commonly available AB and TL lines of zebrafish. Specifically, we will describe techniques that are useful for assessing developmental robustness (i.e., canalization), the trajectory of morphological development (i.e., allometric repatterning), the rate of shape change over development (i.e., heterochrony), and morphological integration.

B. Data Collection The beginning of any GM analysis involves the selection and photographic imaging of samples, and the collection of x,y Cartesian landmark coordinates. Samples can be chosen based on their membership to a particular group of interest (i.e., wildtype vs. mutant zebrafish), and at multiple stages of ontogeny. Imaging is a relatively straightforward but extremely important process whereby samples are photographed in a reproducibly standard position in the presence of a scale bar. These images are then used as a source from which x, y landmark data are collected. Currently the most popular software for x, y data collection is TPSdig2 (Rohlf, 2010), which is used in coordination with TPSutil (Rohlf, 2010). TPSutil creates a file that allows images to be read sequentially into TPSdig2, using the ‘‘build tps files from images’’ function so that the coordinate locations of homologous landmarks can be recorded on images (available at http://life.bio.sunysb.edu/morph/). It is extremely important that x, y coordinates are collected in the same sequence at homologous points on each individual so that they are recognized correctly. Ultimately, these data will be used to statistically test for shape differences between groups or ontogenetic stages. The quality of any statistical test depends upon the quality and amount of data collected. We therefore recommend the use of multiple individuals per group, and

[(Fig._4)TD$IG]

Fig. 4 Three stages of ontogeny in AB and TL lines of zebrafish analyzed for morphometric variation. Note that the seven landmarks used in this study are in homologous positions throughout ontogeny.


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the collection of as many landmarks as are reasonable for any GM study. Here we use a total of 146 specimens and seven landmarks across three stages of ontogeny (10 dpf n = 24 AB, 20 TL; 30 dpf n = 30 AB, 26 TL; and adult n = 25 AB, 21TL) (Fig. 4).

C. General Data Analysis A first step toward analyzing x, y coordinate data is to eliminate variance that is due to differences in the size of specimens, their orientation, and position. Several superimposition algorithms have been developed but the most common approach is a generalized procrustes analysis (GPA), which is available seperately or automatically implemented in a number of morphometric programs (Zelditch et al., 2004). This method superimposes landmark configurations through a process of translation, rotation, and scaling for size (see Fig. 5), in order to minimize the sum of squared distances between corresponding landmarks across specimens (Adams et al., 2004).

[(Fig._5)TD$IG]

Fig. 5

Processes carried out during a generalized Procrustes analysis.

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Once performed, this analysis opens the gateway for a variety of further statistical methods. However, it is important that researchers comparing multiple groups use a single GPA across all specimens, as the comparison of data from separate GPAs can create misleading results. D. Analysis of Shape A landmark-based approach to shape analysis is far more powerful than traditional methods used to discern and describe patterns of shape variation (Parsons et al., 2003). In the simplest terms a geometric approach can double the number of traits examined as compared to traditional linear methods. For example, consider three distances on the head of a zebrafish: the depth of the eye, the length of the lower jaw, and the overall length of the skull. These measurements represent three traits, but if one were simply to place landmarks at the ends of each linear distances, the number of traits would double to six. Moreover, in a landmark-based analysis variation at each landmark is evaluated relative to every other landmark, leading to a more comprehensive picture of how the geometry of a given shape varies between experimental manipulations or over time. After GPA, GM shape analysis typically involves performing a thin-plate spline (TPS) analysis. The analogy that is typically used in the field is that TPS analysis models a starting form (e.g., that of a wild-type zebrafish) as an infinitely thin metal plate that is constrained at some combination of points (i.e., landmarks), but is otherwise free to adopt a target form (e.g., that of a mutant zebrafish) in a way that minimizes bending energy (Bookstein, 1991). Bending energy can be thought of as the degree of deformation that is required to bend the Cartesian coordinate (x, y) system such that two landmark configurations can be superimposed on top of one another (Bookstein, 1991). The total deformation of the TPS can be decomposed into geometrically orthogonal components based on scale/bending energy such that less energy is required to bend a metal plate while holding it at distantly positioned points, whereas much more energy is required to bend the plate to the same vertical extend while holding two points in close proximity (Rohlf and Marcus, 1993). These components (partial warps) can be localized to describe precisely what aspects of shape are different. Because the number of partial warps can be large, as they scale with the number of landmarks, a data reduction analysis is typically performed, principal component analysis (PCA) being the most common. Performing a PCA on partial warp scores is formally referred to as relative warp analysis (Zelditch et al., 2004), and can be an effective method to identify the major axes of shape variation. By applying these methods, we are able to show that adult AB and TL strains of zebrafish significantly differ in craniofacial geometry along PC1 (one-way ANOVA: F-ratio = 35.32, p < 0.001). Differences are largely restricted to the anterior head, and show that TL zebrafish on average have longer upper (yellow region) and lower (purple) jaws than AB wild-type fish, whereas the retroarticular process is longer in ABs than it is in TLs (Fig. 6). TL fish harbor a mutation that results in the expression of their characteristic long tails. It is interesting that, while a craniofacial phenotype


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[(Fig._6)TD$IG]

Fig. 6

Results of a relative warps analysis on craniofacial shape in adult AB and TL lines. Morphological variation between lines along RW1 (the horizontal axis) and RW 2 (the vertical axis). The scatter plot suggests that RW1, the major axis of variation, distinguishes AB and TL lines. Deformation grids along this axis show that AB fish have a relatively smaller maxillae (yellow) and shorter mandible (purple). However, the length of the retroarticular process is larger in ABs. Note that this analysis utilizes 15 landmarks in contrast to the seven used in our ontogenetic analysis because more homologous regions were apparent at adult stages. However, despite this reduction in the amount of shape variation collected AB and TL lines are still distinguishable with seven landmarks. (See Plate no. 12 in the Color Plate Section.)

has never been noted for this stain, TL fish (at least in our lab) also possess elongated dermal bones in the jaw. These results demonstrate the power of GM to discriminate subtle but discrete differences in craniofacial morphology.

E. Analysis of Shape Variation Zebrafish biologists may also be interested in the effects elicited by an experimental treatment or mutation with respect to shape variation. Magnitudes of shape

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variation may also change over the course of ontogeny (Zelditch et al., 2004). With GM it is possible to quantify the degree to which a sample varies and statistically test for differences in the magnitude of variation among groups. While many biologists are familiar with how variance is calculated for univariate traits, shape is a more complex multivariate trait that is not compatible with these more traditional methods. Fortunately, a metric for calculating variance in multivariate traits exists; instead of ‘‘variance’’ the term ‘‘disparity’’ is used. Disparity is calculated using the approach of Foote (1993) as follows: X 2 D¼ ðdi Þ=ðN1Þ Here di represents the Procrustes distance of the centroid of individual i from the centroid of all N individuals. Procrustes distance is normally calculated by comparing an individual specimen to the average shape of a group (which is called the consensus) to provide a standard metric of shape difference. Here we test for differences in measures of disparity across ontogenetic stages between our AB and TL lines. To test for differences in disparity, 1000 permutations of the difference in the disparities between stages within AB and TL lines, as well as between these lines at the same stage were performed. For the permutation tests, residuals of the Procrustes fit were randomly assigned within each treatment group, and then recombined with the mean form of that group to calculate the disparity of the permuted group. The observed results were then compared with the range of values obtained via permutation to determine if the observed values were significant. These tests were performed using the program DisparityBox6 (available at www3.canisius.edu/sheets/ morphsoft.html). Our analyses show that overall disparity decreases over time in both lines of zebrafish. However, in ABs this decrease was statistically significant between 10 and 30 dpf, whereas in TLs there was a significant reduction in disparity that occurred between the 30 dpf and adult stages (Table I). This suggests that although reductions in shape disparity take place in both lines, these changes are initiated at different times between lines. Comparisons between lines over ontogeny show that initially the AB line is approximately twice as disparate than the TL line after 10 dpf (p < 0.001, Table I), but no significant difference in disparity was observed at 30 dpf or in adults (all p > 0.16). In fact, levels of disparity are nearly equal at adult stages

Table I Changes in craniofacial shape disparity over ontogeny in AB and TL lines Ontogenetic comparisons p-values from 1000 permutations

Disparity Line AB TL

10 dpf 0.011 0.005

30 dpf 0.005 0.008

Adult 0.004 0.004

10 to 30 dpf < 0.0001 0.388

30 dpf to adult 0.108 0.047

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Table II The rate of morphological development as measured by Partial Procrustes distancesa Partial Procrustes distances, F, and p-value from comparisons between ontogenetic stages Line AB TL a

10 to 30 dpf 0.125, F = 28.24, p = 0.001 0.138, F = 31.08, p = 0.001

30 dpf to adult 0.129, F = 53.21, p = 0.001 0.140, F = 37.10, p = 0.001

10 dpf to adult 0.218, F = 82.44, p = 0.001 0.239, F = 132.11, p = 0.001

The F and p-values from a test performed between developmental stages within lines is presented. Note that partial Procrustes distances are similar between AB and TL lines and did not differ significantly.

suggesting that an equally canalized phenotype has been achieved in both lines by adulthood (Table I, Fig. 7).

F. Rates of Shape Change Heterochrony has most often been studied in the context of evolutionary biology and is usually defined as a change in the developmental rate or timing of the appearance of features that create parallels between ontogeny and phylogeny (Gould, 1977). More general definitions also exist that do not consider this parallelism, and only require that a developmental event occurs at a different time or rate in a descendant relative to an ancestor (McKinney and McNamara, 1991;

[(Fig._7)TD$IG]

Fig. 7

Plotted GPA superimposed landmarks for each measured stage (10 dpf, 30 dpf, and adult stages) of craniofacial development in AB and TL lines of zebrafish. Note that the scatter of landmarks tends to decrease over ontogeny, happening earlier in the AB line during the period between 10 and 30 dpf, while for TLs a reduction in scatter does not occur until the period between 30 dpf and adult stages.

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McNamara, 1995, 1997). In other words, heterochrony causes a decoupling of shape from developmental time and may also be a theory of interest to developmental biologists comparing wild-type (ancestor) to mutant (descendant) phenotypes. Since any developmental process is temporal in nature, thereby making it susceptible to heterochronic modifications, it is likely that heterochrony is a widespread phenomenon for zebrafish mutants relative to their respective wild-type ancestors. The rate of morphological development over ontogeny can be determined empirically in the absence of trajectories, but we recommend that researchers use the approaches for quantifying trajectories discussed below in a complementary fashion to gain a comprehensive idea of morphological development. Determination of the ontogenetic rate of morphological development is based on the morphometric distance between shapes at a given ontogenetic stage in relation to the average specimen (of a treatment group at one stage of the experiment), and this is calculated as the partial Procrustes distance (Bookstein, 1996; Dryden and Mardia, 1998; Zelditch et al., 2004). We tested whether rates of morphological development differed over ontogeny among AB and TL zebrafish lines (Table II). Ontogenetic tests for differences in the amount of shape change in morphometric distance between 10 and 30 dpf, 30 dpf and adult, and 10 dpf and adult specimens were performed between lines using 900 bootstrap replicates while stage specific tests between lines were conducted by performing Goodall’s F-test in the program TwoGroup6h (available at http://www3.canisius.edu/sheets/morphsoft.html). Goodall’s F-test compares the difference in the mean shape between two samples relative to the shape variation found within the samples. Our analysis showed that rates of morphological development did not differ between our zebrafish lines at any of the ontogenetic intervals tested. However, stage specific tests did reveal that significant differences in the position of AB and TL lines in shape space (distance) were present at all three stages of the experiment, indicating that lines did differ in shape at each stage. Taken together these data suggest that while rates of shape change do not differ between lines at any stage, both lines are characterized by marked shifts in rates of morphological development over time, which may be a common feature of zebrafish craniofacial development. G. Trajectories of Shape in Zebrafish Lines Mutations may also modify the direction of ontogenetic shape change in zebrafish. This aspect of morphological development is distinct from changes in shape due to alterations in the rate or timing of development (i.e., heterochrony). Whereas heterochrony refers to changes in the timing of developmental events, allometric repatterning refers to changes in the trajectory of development. Allometric repatterning can gradually develop over long intervals of ontogeny whereas heterochrony may be sudden. Since both heterochrony and allometric repatterning can affect shape, it is useful to consider both phenomena in relation to each other. We can quantitatively test whether allometric repatterning has occurred between AB and TL zebrafish lines by measuring their shape trajectories in relation to a


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Table III The angles between ontogenetic trajectories of craniofacial shape in AB and TL linesa Angle between ontogenetic stages Line AB TL a

10 to 30 dpf 84.1 * 140.5 *

30 dpf to adult 127.0 * 76.5

10 dpf to adult 101.5 * 101.5

Trajectories are measured between all stages including 10 dpf, 30 dpf, and adults.

continuous variable (ln centroid size in this case). We accomplish this here by measuring ontogenetic growth angles, which are the arc cosines of vector correlations (Zelditch et al., 2000), between AB and TL zebrafish at 10 dpf, 30 dpf, and adult stages. Similar to the analysis of disparity described above the calculation of angle is simply a metric and there is no true statistical model. We therefore compared the observed angle between ontogenetic vectors to bootstrapped estimates of these angles. The null hypothesis was an angle of zero, which is biologically equivalent to the hypothesis of a conserved ontogenetic trajectory of shape between AB and TL lines. To test this null hypothesis, 95% confidence intervals for the angles were created through a procedure involving 900 bootstrapped replicates. The angle between the vectors would be considered statistically significant if they exceeded those drawn from a bootstrapped distribution within a group. In other words the differences between groups of interest would have to exceed those generated from bootstrapping within both groups. This explains how smaller overall differences in trajectories may sometimes be significant, whereas larger differences are not. Growth angles and bootstrap estimates of 95% confidence intervals were calculated in VecCompar6c (available at http://www3.canisius.edu/sheets/morphsoft.html). Our analysis shows that ontogenetic trajectory differs between lines at the 10 and 30 dpf stages (110.1 and 94.7 respectively). Trajectories also differ between stages within lines. This is observed for each measured stage within our AB line, and for all but one comparison in out TL line (Table III). However, the trajectory for TLs at 10 dpf is only marginally different from the one present in adult TLs. This suggests that ontogenetic trajectories are relatively dynamic in ABs, while in TLs the trajectory is set early in development and persists with little change to adulthood. Considered in the context of data reported above, these results are consistent with the hypothesis that the unique TL phenotype is due, in part, to an altered growth trajectory established early in development.

H. Morphological Integration Morphological integration refers to the degree to which functionally related traits are correlated (Cheverud et al., 1996; Olson and Miller, 1958). Implicit to this theory is the concept of modularity. A module refers to a complex of traits that are inherited

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together, and which are independent of other character complexes (Wagner, 1996). Developmental architecture figures prominently in questions of integration and modularity, because genetic and functional modules are mediated by developmental processes (Atchley and Hall, 1991; Cheverud, 1996; Klingenberg, 2004; Klingenberg, 2003). For example, the integration of component parts may result from any combination of the following phenomena: common function, shared developmental pathways, pleiotropy, or heritable epigenetic effects. One way to quantitatively assess general levels of morphological integration is through the combined use of GM and PCA. High levels of positional covariation among landmarks will skew the distribution of shape variation among the PC axes such that most of the variation will be explained by only a few axes (Wagner, 1984, 1990; Young, 2006). This would be reflected by the existence of one or a few axes that explains a high amount of variation followed by a ‘‘distinct’’drop in explanatory power in subsequent PCs. Several methods exist for determining whether differences in the degree of integration exist between groups of interest. For example, a Chisquared test can be used to determine if and when there was a strong drop in the levels of variation explained by subsequent PC axes. This method determines whether the total shape variation in a dataset is spread out among many PC axes (a low level of integration) or concentrated within a small number of the initial PC axes (a higher level of integration). This method is incorporated into the program PCAgen (available at http://www3.canisius.edu/sheets/morphsoft.html). For a detailed explanation of this method, see p. 211–254 in Morrison (2004). Alternatively, it is possible to simply compare the variance of eigenvalues (the explanatory power of each PC) produced from a PCA on each group by using an F-test. This should be available in most statistical packages, and it is even manageable to perform this test by hand. An increased level of variation in eigenvalues is indicative of increased integration. Our analysis of integration revealed differences both between AB and TL lines, and between ontogenetic stages within lines. At 30 dpf the TL line showed a significantly higher degree of integration than ABs (variance = 1.32006 vs. 2.45007, respectively, F-ratio9,9 = 3.21, p = 0.02). The comparisons between 10 and 30 dpf in ABs (variance = 2.45006 to 2.41007 respectively, F-ratio9,9 = 0.0982, p-value = 0.002) showed a reduction in integration had occurred. This period of ontogeny in ABs likely accounts for the majority of change in levels of integration as differences were not detected between 30 dpf and adult stages, but a similar degree of difference was present between 10 dpf and adult stages (variance = 2.45006 to 1.94007, respectively, F-ratio9,9 = 0.079, p < 0.001). For TLs a difference in integration did not appear until 30 dpf and adult stages were compared (variance = 1.32006 vs. 1.64007, respectively, F-ratio9,9, p = 0.005), and this period of ontogeny is also likely to be responsible for the differences observed between 10 dpf and adult stages (7.62007 to 1.64007, respectively, F-ratio9,9 = 0.215, p = 0.032). Thus, within both lines we observed a decrease in integration over ontogeny, but this drop occurred earlier and more precipitously in ABs, leading to the observed difference in integration between lines at 30 dpf.


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[(Fig._8)TD$IG]

Fig. 8 An outline of our approach that incorporates the use of GM and quantitative methods to study development. GM offers an ability to quantify and assess variation at an intermediate stage of discovery. This means that GM can provide a way of testing hypotheses about potential mutant phenotypes and their development, as well as providing data for further explorations into the genetic basis of mutant phenotypes.

Taken together these analyses suggest that integration is an ontogenetically dynamic property of phenotypes. It is also noteworthy that the timing of changes in integration seems to coincide with the changes in growth trajectory that were observed earlier. In this case it may be that patterns of integration are a surrogate for trajectories in that

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interactions among anatomical features also determine the spatial direction of development.

I. Synthesis of Quantitative Data We have demonstrated here that GM can be used to identify differences in craniofacial shape. More importantly we have shown that with the proper experimental design GM can be used to quantify the dynamic processes involved in development and provide an explicit quantitative link between development within and beyond the embryo. We feel this provides a major opportunity to understand when and how key changes occur to produce clinically and evolutionary relevant variation. For clinicians the approaches demonstrated here may be especially useful for the identification of progressive syndromes at very early stages when therapeutic approaches may have their greatest impact. They also enable the identification of life stages when mutational effects are most pronounced. For evolutionary biologists, these techniques may lead to the identification of adaptively important ontogenetic processes. Opposite to developmental biology, the traditional paradigm in evolutionary biology has been to study adult phenotypes. We suggest that the use of GM can be a means to link embryonic and adult life history stages, and is thus equally valuable to both fields of study (see Fig. 8 for an outline of our approach). Indeed, this approach has recently gained traction in evolutionary biology, and such studies have fallen under the title of ‘‘evo-devo’’ (Carroll et al., 2005; Gilbert and Epel, 2009). Critical to both of these biological disciplines is a consideration of developmental processes that occur after embryonic but before adult stages, and GM can provide an empirical avenue for this view.

IV. A Complementary Approach: The Use of Natural Variation to Complement that Generated in the Lab for Understanding Jaw Morphogenesis While the use of mutagenic analyses in a select group of model organisms, including the zebrafish, has provided a powerful approach for understanding the genetic basis of traits that emerge very early in development, they have been less effective for understanding complex traits that manifest themselves later in development. This may be due to an ascertainment bias introduced by researchers focusing on embryonic stages, but there are likely other complications. The most confounding effects of induced mutations concern pleiotropy. Most mutations isolated in mutant screens occur in the coding regions of genes and lead to the severe attenuation or complete knockdown of gene function. As a result genetic screens typically recover mutant animals with defects at early developmental stages when the gene first provides an essential function, generally in early embryonic development. Such mutations often result in early lethality, which in turn masks the


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ascertainment of their effects on adult phenotypes. Thus, phenotype-driven mutagenesis screens often preclude analyses at later developmental stages, and while we have discussed different methods for mitigating this limitation in zebrafish, in the following section we will review another, complementary approach that can be used to identify the genetic mechanisms that contribute to craniofacial development – the analysis of variation among natural populations. The details of this approach have been elaborated on elsewhere (Albertson et al., 2009), but a brief overview is worth repeating here. Closely related cichlid fishes from lakes in the East African Rift Valley have undergone extensive evolutionary modifications of their oral jaws and faces, providing an array of ‘‘evolutionary mutant’’ models for medically important human craniofacial variation that may not always be present in model organisms like the zebrafish (e.g., micrognathia, midface hypoplasia, facial asymmetries). The utility of integrating models organisms with emerging models was illustrated by Albertson et al. (2005), where a role for bmp4 in directing the development of mandibular shape was assessed through a combination of quantitative trait loci analysis and experimental embryology. Both the expression of and allelic variation in bmp4 was shown to be associated with quantitative differences in the shape of the lower jaws of cichlid fishes. However, because cichlids were less amenable to genetic manipulation zebrafish were used as a surrogate to investigate how altering levels of bmp4 would affect the development of the craniofacial shape. Manipulations of bmp4 levels in zebrafish lead to variable effects on the jaw, but this variation mirrored the natural, quantitative variation in jaw shape seen among cichlid species. These data suggested a novel role for levels of bmp4 in modulating the jaw shape, and underscored the utility of integrating work in both model and nonmodel organisms to paint a more comprehensive picture of how morphology develops and evolves. The results of recent research in Darwin’s finches bears striking similarity to that in cichlids. Here, investigators identified roles for bmp4 and calm1 in regulating differences in beak width/height and beak length, respectively, which were confirmed independently using genetic manipulations in chicken and duck models (Abzhanov et al., 2004, 2006; Wu et al., 2004, 2006). These studies underscore the utility of integrating work in the laboratory with studies of evolutionary model systems, as these approaches have highly complementary attributes; laboratory models bring experimental tractability to the table, whereas evolutionary models provide extensive amounts of novel phenotypic variation to bear on questions related to the morphogenesis of the craniofacial shape. Fortunately, whereas postembryonic screens are hard to come by, for researchers interested in understanding craniofacial shape there is no shortage of examples where adaptive divergence has created a plethora of variation in adult head and jaw characteristics. Space limitations preclude a thorough review of other natural systems that embody both the experimental and phenotypic potential to advance our understanding of craniofacial development, but these would most certainly include the blind cavefish (Astyanax mexicanus), which has been used to study roles for SHH signaling in jaw development (Yamamoto et al., 2009), the Antarctic notothenioid species

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flock used to study craniofacial patterning and skeletogenesis (Albertson et al., 2009, 2010), as well as the threespine stickleback (Gasterosteus aculeatus) to analyze an array of mineralized tissue phenotypes (Cresko et al., 2004; Colosimo et al., 2005; Kimmel et al., 2005; Shapiro et al., 2004; 2006). With technological advances in molecular biology and genomics, the once distinct line that separated model from nonmodel organisms is becoming increasingly blurred (Albertson et al., 2009). While the tools and resources that are available to zebrafish researchers far exceed those that can be applied in nontraditional fish systems, we contend that a niche also exists for these nonmodel systems in investigations of the morphogenesis of craniofacial shape.

V. Implications and Conclusions The zebrafish has become an invaluable model system that has provided detailed knowledge of the molecules that regulate early embryonic development. However, due to the early lethal phenotypes exhibited by most zebrafish mutants, significantly less is known about craniofacial development beyond embryonic stages. There is a significant gap in our understanding of later developmental events including tooth replacement, as well as bone development, growth, and remodeling. We argue that an expansion of the current paradigm is needed, and that this will include (1) postembryonic mutagenesis screens, (2) the application of quantitative shape analyses to evaluate and statistically compare patterns of phenotypic variation among wild-type and mutant lines over extended periods of development, and (3) comparative analyses in evolutionary models that exhibit relevant patterns of phenotypic variation (Albertson and Yelick, 2009). From a biomedical perspective, the study of the zebrafish beyond the embryological stages may provide insights into the developmental and genetic basis of progressive craniofacial syndromes. Over 70% of all birth defects are associated with craniofacial malformations (Hall, 1999), and while surgery can correct many of these (e.g., orofacial clefting), syndromes associated with oral, dental, and craniofacial defects continue to pose major therapeutic challenges for clinicians (NIDCR, 2007). The NIDRC initiative, FaceBase, states that ‘‘much research is needed to achieve a molecular and cellular understanding of the mechanisms by which genes and gene products interact to generate complex phenotypes,’’ and argues for broadening support for projects ‘‘that merge genetics, developmental biology, and modeling expertise and are aimed at achieving a systems biology comprehensive understanding of the mechanisms that underlie complex craniofacial phenotypes.’’ Many of the statistical frameworks presented here could be folded into genetic and developmental studies to contribute to this initiative. Indeed, by modeling the craniofacial skeleton as a dynamic multivariate trait and analyzing accordingly, we will gain a more accurate understanding of phenotype. Since mutational analysis are, by definition, based on phenotype, these insights will enable a far more comprehensive understanding of the number, type,


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mode of action and interaction of genes that underlie craniofacial form at all stages of development. From an evolutionary perspective, the alteration of developmental processes in craniofacial traits over ontogeny often plays a key role in producing adaptive diversity (Carroll et al., 2005). Adaptive phenotypes in the wild are most often studied in adult animals, and while there is mounting evidence to suggest that adaptive variation studied in adults has an early embryonic origin (e.g., Abzhanov et al., 2004, 2006; Albertson et al., 2005), there are undoubtedly myriad adaptive phenotypes that also arise postembryonically (West-Eberhard, 2003). In the sections above, we have presented ways in which natural variation in the wild can compliment that produced by induced mutations, but this may well be a two-way street. Which prompts us to ask, can craniofacial variation induced via a chemical mutagen inform an understanding of the molecular nature of evolutionary change? Since many of the most spectacular adaptive radiations among vertebrates are associated with extensive modifications of the craniofacial skeleton, and given that a proximate genetic basis for natural variation in craniofacial form remains unknown, this question is well worth thinking about. References Abzhanov, A., Protas, M., Grant, B. R., Grant, P. R., and Tabin, C. J. (2004). Bmp4 and morphological variation of beaks in Darwin’s finches. Science 305, 1462–1465. Abzhanov, A., Kuo, W. P., Hartmann, C., Grant, B. R., Grant, P. R., Tabin, C. J. (2006). The calmodulin pathway and evolution of elongated beak morphology in Darwin’s finches. Nature 442, 563–567. Adams, D. C., Rohlf, F. J., and Slice, D. E. (2004). Geometric morphometrics: ten years of progress following the ‘revolution’. Ital. J. Zool. 71, 5–16. Albertson, R. C., and Yelick, P. C. (2004). Morphogenesis of the jaw: development beyond the embryo. Methods Cell Biol. 76, 437–454. Albertson, R. C., and Yelick, P. C. (2007). Fgf8 haploinsufficiency results in distinct craniofacial defects in adult zebrafish. Dev. Biol. 306, 505–515. Albertson, R. C., and Yelick, P. C. (2005). Roles for fgf8 signaling in left-right patterning of the visceral organs and craniofacial skeleton. Dev. Biol. 283, 310–321. Albertson, R. C., Streelman, J. T., Kocher, T. D., and Yelick, P. C. (2005). Integration and evolution of the cichlid mandible: Molecular basis of alternate feeding strategies. Proc. Natl. Acad. Sci. U.S.A. 102, 16287–16292. Albertson, R. C., Cresko, W., Detrich, H. W., and Postlethwait, J. H. (2009). Evolutionary mutant models for human disease. Trends Gen. 25, 74–81. Albertson, R. C., and Yelick, P. C. (2009). Morphogenesis of the jaw: development beyond the embryo. Methods Cell Biol. 91, 453–473. Albertson, R. C., Yan, Y. -L., Titus, T. A., Pisano, E., Vacchi, M., Yelick, P. C., Detrich, W. H., Postlethwait, J. H. (2010). Molecular pedomorphism underlies craniofacial skeletal evolution in Antarctic notothenioid fishes. BMC Evol. Biol. 10, 4. Atchley, W. R., and Hall, B. K. (1991). A model for the development and evolution of complex morphological structures. Biol. Rev. 66, 101–157. Basel, D., and Steiner, R. D. (2009). Osteogenesis imperfecta: recent findings shed new light on this once well-understood condition. Genet. Med. 11, 375–385. Bookstein, F. L. (1991). Morphometric Tools for Landmark Data: Geometry and Biology. Cambridge University Press, Cambridge.

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Kevin J. Parsons et al. Bookstein, F. L. (1996). Biometrics, biomathematics and the morphometric synthesis. Bull. Math. Biol. 58, 313–365. Brand, M., Heisenberg, C. -P., Jiang, Y. -J., Beuchle, D., Lun, K., van Eeden, F. J. M., Furutani-Seiki, M., Granato, M., Haffter, P., Hammerschmidt, M., Kane, D. A., Kelsh, R. N., Mullins, M. C., Odenthal, J., N€ usslein-Volhard, C. (1996). Mutations in zebrafish genes affecting the formation of the boundary between midbrain and hindbrain. Development 123, 179–190. Carroll, S. B., Grenier, J. K., and Weatherbee, S. D. (2005). From DNA to diversity: molecular genetics and the evolution of animal design, 2nd edn. Blackwell, Oxford. Cheverud, J. M. (1996). Developmental integration and the evolution of pleiotropy. Am. Zool. 36, 44–50. Cooper, W. J., and Albertson, R. C. (2008). Quantification and variation in experimental studies of morphogenesis. Dev. Biol. 321, 295–302. Cresko, W. A., Amores, A., Wilson, C., Murphy, J., and Currey, M., et al. (2004). Parallel genetic basis for repeated evolution of armor loss in Alaskan threespine stickleback populations. Proc. Natl. Acad. Sci. U.S.A. 101, 6050–6055. Dryden, I. L., and Mardia, K. V. (1998). Statistical shape analysis. Wiley, Chichester. Filmus, J., and Selleck, S. B. (2001). Glypicans: proteoglycans with a surprise. J. Clin. Invest. 108, 497–501. Fisher, S., Jagadeeswaran, P., and Halpern, M. E. (2003). Radiographic analysis of zebrafish skeletal defects. Dev. Biol. 264, 64–76. Foote, M. (1993). Contributions of individual taxa to overall morphological disparity. Paleobiology 19, 40n3–419. Fransson, L. A. (2003). Glypicans. Int. J. Biochem. Cell. Biol. 35, 125–129. Fuiman, L. A. (1983). Growth gradients in fish larvae. J. Fish Biol. 23, 117–123. Gilbert, S. F., and Epel, D. (2009). Ecological Developmental Biology: Integrating Epigenetics, Medicine, and Evolution. Sinauer Associates, Sunderland, MA. Gould, S. J. (1977). Ontogeny and Phylogeny. Belknap Press, Cambridge. Hall, B. K. (1999). The Neural Crest in Development and Evolution. Springer-Verlag, New York. Harris, M. P., Rohner, N., Schwarz, H., Perathoner, S., Konstantinidis, P., N€ usslein-Volhard, C. (2008). Zebrafish eda and edar mutants reveal conserved and ancestral roles of ectodysplasin signaling in vertebrates. PLoS Genet. 4, e1000206. Kimmel, C. B., Ullmann, B., Walker, C., Wilson, C., and Currey, M., et al. (2005). Evolution and development of facial bone morphology in threespine sticklebacks. Proc. Natl. Acad. Sci. U.S.A. 102, 5791–5796. Klingenberg, C. P. (2004). Integration, modules and development: molecules to morphology to evolution. In ‘‘Phenotypic Integration: Studying the Ecology and Evolution of Complex Phenotypes,’’ (M. Pigliucci, and K. Preston, eds.), pp. 213–230. Oxford University Press, New York. Klingenberg, C. P. (2003). Developmental instability as a research tool: using patterns of fluctuating asymmetry to infer the developmental origins of morphological integration. In, ‘‘Developmental instability: causes and consequences.’’ Oxford University Press, New York. LeClair, E. E., Mui, S. R., Huang, A., Topczewska, J. M., and Topczewski, J. (2009). Craniofacial skeletal defects of adult zebrafish Glypican 4 (knypek) mutants. Dev. Dyn. 238, 2550–2563. Loy, A., Bertelletti, M., Costa, C., Ferlin, L., and Cataudella, S. (2001). Shape changes and growth trajectories in the early stages of three species of the genus Diplodus (Perciformes Sparidae). J. Morph. 250, 24–33. McKinney, M. L., and McNamara, K. J. (1991). Heterochrony: the evolution of ontogeny. Plenum, New York. McNamara, K. J. (1995). Evolutionary change and heterochrony. Wiley, Chichester, UK. McNamara, K. J. (1997). Shapes of time: the evolution of growth and development. Johns Hopkins University Press, Baltimore. Mikkola, M. L. (2009). Molecular aspects of hypohidrotic ectodermal dysplasia. Am. J. Med. Genet. A 149, 2031–2036.


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Morrison, D. F. (2004). Multivariate Statistical Methods. Duxbury Press, Pacific Grove, CA, p. 480. Morriss-Kay, G. M., and Wilkie, A. (2005). Growth of the normal skull vault and its alteration in craniosynostosis: insights from human genetics and experimental studies. J. Anat. 207, 637–653. Neuhauss, S. C., Solnica-Krezel, L., Schier, A. F., Zwartkruis, F., Stemple, D. L., Malicki, J., Abdelilah, S., Stainier, D. Y., Driever, W. (1996). Mutations affecting craniofacial development in zebrafish. Development 123, 357–367. Olson, E. C., and Miller, R. L. (1958). Morphological Integration. University of Chicago Press, Chicago. Osse, J. W. M. (1990). Form changes in fish larvae in relation to changing demands of function. Neth. J. Zool. 40, 362–385. Parsons, K. J., Robinson, B. W., and Hrbek, T. (2003). Getting into shape: an empirical comparison of traditional truss-based morphometric methods with a newer geometric method applied to New World cichlids. Environ. Biol. Fishes 67, 417–431. Piotrowski, T., Schilling, T. F., Brand, M., Jiang, Y. J., Heisenberg, C. P., Beuchle, D., Grandel, H., van Eeden, F. J., Furutani-Seiki, M., Granato, M., Haffter, P., Hammerschmidt, M., Kane, D. A., Kelsh, R. N., Mullins, M. C., Odenthal, J., Warga, R. M., N€ usslein-Volhard, C. (1996). Jaw and branchial arch mutants in zebrafish II: anterior arches and cartilage differentiation. Development 123, 345–356. Rohlf, F. J., and Marcus, L. F. (1993). A revolution in morphometrics. Trends Ecol. Evol. 8, 129–132. Schilling, T. F., Piotrowski, T., Grandel, H., Brand, M., Heisenberg, C. P., Jiang, Y. J., Beuchle, D., Hammerschmidt, M., Kane, D. A., Mullins, M. C., van Eeden, F. J., Kelsh, R. N., Furutani-Seiki, M., Granato, M., Haffter, P., Odenthal, J., Warga, R. M., Trowe, T., N€ usslein-Volhard, C. (1996). Jaw and branchial arch mutants in zebrafish I: branchial arches. Development 123, 329–344. Shapiro, M. D., Marks, M. E., Peichel, C. L., Blackman, B. K., and Nereng, K. S., et al. (2004). Genetic and developmental basis of evolutionary pelvic reduction in threespine sticklebacks. Nature 428, 717–723. Shapiro, M. D., Bell, M. A., and Kingsley DM. (2006). Parallel genetic origins of pelvic reduction in vertebrates. Proc. Natl. Acad. Sci. USA 103, 13753–13758. Stewart-Swift, C., Andreeva, V., Gibert, Y., Connolly, M., Lee, Y., Fraher, D., Burt, J., Cardarelli, J., Lattanzi, V., Sullivan, D., Albertson, R. C., and Yelick P. C. (2010). Forward Genetic ENU Chemical Mutagenesis Screen for Mineralized Craniofacial, Skeletal and Tooth Mutants in Zebrafish. 9th International Zebrafish Development and Genetics Conference, Madison, WI, June16–20, 2010. Topczewski, J., Sepich, D. S., Myers, D. C., Walker, C., Amores, A., Lele, Z., Hammerschmidt, M., Postlethwait, J., Solnica-Krezel, L. (2001). The zebrafish glypican knypek controls cell polarity during gastrulation movements of convergent extension. Dev. Cell. 1, 251–264. Wagner, G. P. (1984). Coevolution of functionally constrained characters: prerequisites for adaptive versatility. Biosystems 17, 51–55. € Wagner, G. P. (1990). A comparative study of morphological integration in Apis mellifera. Zeitschrift fYr zoologische. Systematik und Evolutionsforschung 28, 48–61. Wagner, G. P. (1996). Homologues, natural kinds and the evolution of modularity. Am. Zool. 36, 36–43. Weatherly, A. H. (1990). Approaches to understanding fish growth. Trans. Am. Fish. Soc. 119, 662–672. West-Eberhard, M. J. (2003). Developmental Plasticity and Evolution. Oxford University Press, New York. Wu, P., Jiang, T. X., Shen, J. Y., Widelitz, R. B., and Chuong, C. M. (2006). Morphoregulation of avian beaks: comparative mapping of growth zone activities and morphological evolution. Dev. Dyn. 235, 1400–1412. Wu, P., Jiang, T. X., Suksaweang, S., Widelitz, R. B., and Chuong, C. M. (2004). Molecular shaping of the beak. Science 305, 1465–1556. Yamamoto, Y., Byerly, M. S., Jackman, W. R., and Jeffery, W. R. (2009). Pleitropic functions of embryonic sonic hedgehog expression link jaw and taste bud amplification with eye loss during cavefish evolution. Dev. Biol. 330, 200–211. Young, N. M. (2006). Function, ontogeny and canalization of shape variance in the primate scapula. J. Anat. 209, 623–636.

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CHAPTER 12

Associative Learning in Zebrafish (Danio rerio) Robert Gerlai Department of Psychology, University of Toronto, Mississauga, Ontario, Canada

Abstract I. Introduction A. Why Study Associative Learning? II. Rationale A. Why Zebrafish? B. Behavior: The Bottleneck III. Methods and Discussion A. Ethology: A Fruitful Guiding Principle in Designing Learning Tasks B. What Motivates Zebrafish? C. The Perceptual Demands of the Learning Task: Focus on Visual Stimuli D. What can Zebrafish Learn: Simple Two Cue Association versus Complex Relational Learning? E. How Can We Make Associative Learning Tasks High Throughput? F. The Screening Strategy G. Memory: A Blank Slate H. The Devil is in the Details I. Things We Did Not Cover IV. Summary Acknowledgments References

Abstract The zebrafish has been one of the primary study species utilized in developmental biology. However, it is also gaining increasing amount of interest in other disciplines of biology including behavioral neuroscience; the numerous genetic tools developed and the large amount of genetic information accumulated for this species by now make it an excellent tool for the analysis of the mechanisms of complex central nervous system characteristics. Although several studies have investigated the METHODS IN CELL BIOLOGY, VOL 101 Copyright 2011. Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-387036-0.00012-8

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biological and genetic underpinnings of associative learning (and memory), given the complexity of these phenomena, much remains to be discovered. In the past, the zebrafish has been employed particularly successfully in screening applications where a large number of mutations or drug effects had to be analyzed. Briefly, the practical simplicity and system complexity of the zebrafish may make this species an excellent tool also for the analysis of the mechanisms of associative learning. Screening, however, requires appropriate phenotypical (in this case behavioral) paradigms. A step in this direction is the characterization of learning abilities of zebrafish. The number of studies focused on cognitive and/or mnemonic characteristics of zebrafish is orders of magnitude smaller than those with rats or mice, but recently zebrafish has also started to be utilized in this research. The current chapter reviews these most recent developments. It also discusses certain unique features of zebrafish that must be taken into account when designing an associative learning task and how these tasks may be made high throughput.

I. Introduction A. Why Study Associative Learning? Learning and memory is a ubiquitous feature of all animals. From single cell organisms through the nematode and Drosophila to higher order vertebrates, learning and memory allow the organism to respond plastically to the changing environment. Instead of having to wait for hundreds, perhaps thousands, of generations to make individuals genetically adjust (evolve), that is, to adapt properly to local environmental conditions, learning and memory can achieve the ‘‘adaptation’’ within a fraction of the lifetime of the organism. Learning in a broad sense here is defined as the acquisition of experience-dependent change in behavior, and memory is thought of as the consolidation, storage, and recall of the acquired information. Here, learning and memory are used interchangeably in the sense that these processes represent the two sides of the same coin: a temporal series of neurobiological mechanisms that lead to the manifestation of experience-dependent behavioral change. Associative learning, the focus of the current chapter, is one of the most widely studied and also complex forms of learning and memory (e.g., a simple Medline (PubMed) search with the keyword ‘‘associative learning’’ returns more than 20,000 entries). Associative learning requires the acquisition of temporal and/or causal relationship between at least two stimuli. The different forms of associative learning are not reviewed here because this question has been extensively discussed by others in the literature (e.g., Eichenbaum, 2006; Squire, 2004; Tulving, 1987). Suffice it to say that the simplest form of associative learning is when the animal (human or nonhuman) learns the association between two stimuli, US (unconditioned stimulus) and CS (the conditioned stimulus). One of the most complex forms of associative learning is relational learning, in which the animal learns loose relationships of a potentially large number of cues or stimuli (Eichenbaum, 2004). An example of the latter is episodic memory and spatial learning (Squire, 2004).

12. Associative Learning in Zebrafish (Danio rerio)

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Apart from the scientifically fascinating aspect of the question of how associative learning works, there is also an important human clinical relevance of these studies. Numerous CNS disorders, for example neurodegenerative disorders including Alzheimer’s disease, are associated with impaired associative learning, particularly of the relational type (Dickerson and Eichenbaum, 2010). Concerted efforts have been and are being made to understand the mechanisms of these learning processes and to develop treatment applications. Indeed, by now we know a lot about intricate molecular, synaptic electrophysiological, and neuroanatomical (microstructural as well as macrostructural) aspects of these processes. For example, the second edition of the book, Mechanisms of Memory (one of the most comprehensive treatments of the biochemistry of learning and memory) (Sweatt, 2010), enlists hundreds of already known molecular components involved in one way or another in processes associated with memory formation. Is there anything else left to discover? The answer to this question is a resounding yes. According to conservative estimates, vertebrate genomes harbor approximately 30,000 to 40,000 genes and at least 50% of these genes are expressed in the brain (Dıaz, 2009; Fritzsch, 1998). A large proportion of these genes is believed to subserve neural plasticity, one of the fundamental and most crucial functions of the brain (Sweatt, 2010). Briefly, one may expect as many as 10–15 thousand genes involved in learning and memory processes, a staggering number compared to a few hundred genes so far known to underlie these phenomena.

II. Rationale A. Why Zebrafish? Most of the studies that led to the discoveries of the molecular mechanisms of learning and memory used mammals, mainly the house mouse, as a model organism, and by now most research methods from transgenic techniques (Babinet et al., 1989) to the Morris Water Maze spatial learning task (Gerlai, 2001) have been thoroughly tried and tested for the mouse. Why would the zebrafish, a newcomer in this research, be useful? At this point it is too early to know whether it indeed will be; however, there are some compelling reasons for optimism. Perhaps the most important of these is the fact that zebrafish is highly amenable to high-throughput screening. Identification of the thousands of molecular components involved in learning and memory may require such screening applications. Forward genetics screens have the potential to cover the entire genome and identify numerous mutations leading to the localization and identification of the genes and molecular pathways involved (e.g., Haffter and N€ ussleinVolhard, 1996). Screening for learning mutants has, of course, been attempted in the house mouse (Reijmers et al., 2006), but so far the results have not been stellar. This may partly be due to the high costs and the practical limitations associated with having to work with thousands of mice. Due to its small size (4 cm when

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adult), ease of maintenance, and prolific nature (200–300 eggs per spawning per female every other day), zebrafish has been particularly appropriate for forward genetics screens (Haffer and N€ usslein-Volhard, 1996; Patton and Zon, 2001). One can house tens of thousands of zebrafish in a small room, whereas housing this number of mice would require a substantially larger amount of space. In addition, but very importantly, by now numerous genetic markers have been developed for the zebrafish (Patton and Zon, 2001), and along with other methods (for a recent review see e.g., Gerlai, 2010) identification of the genes harboring the mutations is within our reach. Also important is the translational relevance of zebrafish, that is, the high nucleotide sequence homology and functional similarities between mammalian and zebrafish genes (e.g., Reimers et al., 2004; Renier et al., 2007). Taken together, these features of zebrafish may make forward genetic screens (and also drug screens) more efficient with this species. Thus, given the complexity of, that is, the potentially large number of genes involved in learning and memory processes, zebrafish may offer a unique solution.

B. Behavior: The Bottleneck The cornerstone of forward genetics studies is the phenotypical screening and characterization tools (Gerlai, 2002; Gerlai and Clayton, 1999, Sison et al., 2006). Although the zebrafish has a great potential in behavioral brain research, there is a serious bottleneck that may limit its utility: the phenotypical testing tools that would be required for the screening are often rudimentary or not available at all (Sison et al., 2006). A simple Medline (PubMed) literature search with key words ‘‘zebrafish’’ and ‘‘learning’’ returns 102 publications, a fraction of which is actually about learning and/or memory. A similar search with the keywords ‘‘mouse’’ and ‘‘learning’’ gives 14,616 publications. This is not because zebrafish cannot learn or cannot be used in learning studies. Indeed, most recently there is an upsurge of publications in which zebrafish learning and memory are analyzed. For example, zebrafish have been found to perform well in learning tasks including a one trial avoidance learning paradigm (Blank et al., 2009), olfactory conditioning (Braubach et al., 2009), place conditioning (Eddins et al., 2009), appetitive choice discrimination (Bilotta et al., 2005), active avoidance conditioning (Xu et al., 2007), and visual discrimination learning in the T-maze (Colwill et al., 2005), and even an automated learning paradigm has been published (Hicks et al., 2006). Our laboratory is one of those that focus on this problem. We, as others before us (for review see e.g., Gerlai, 2001; also see Gerlai and Clayton, 1999), believe that the level of sophistication the past decades have seen in recombinant DNA technologies must be met with similarly powerful behavioral methods, an argument especially true for forward genetic screens where the detection of mutants is solely dependent upon the behavioral tools. Therefore, characterization of the cognitive and mnemonic characteristics of zebrafish, and the development of learning and memory paradigms are important.


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In this chapter, I review the growing literature on associative learning in zebrafish, but admittedly with a bias. I focus on those questions we, that is, our laboratory, find most interesting. Although others may have a different bias, I hope that the current chapter will give a sufficient snapshot of this fast developing and very new research area: the analysis of associative learning in zebrafish. In this chapter, I discuss such fundamental questions as what motivates zebrafish, how one can measure simple associative learning, and whether zebrafish are capable of learning more complex, that is, relational-type associative tasks. Last, I discuss some points as to how one could automate a learning task and make it high enough throughput for mutagenesis or drug screening, and what screening strategies may make it possible to find mutations that specifically affect learning and/or memory.

III. Methods and Discussion A. Ethology: A Fruitful Guiding Principle in Designing Learning Tasks Unfortunately, zebrafish researchers do not have the luxury that characterizes the work of others studying more traditional laboratory model organisms including the house mouse, the rat, or even the fruit fly. We do not have the ability to rely on several decades of research comprising tens of thousands of publications when we face the question: How shall we test behavior, in this case associative learning, in our subjects? We know little about what motivates zebrafish, what motor and perceptual constraints and peculiarities these animals possess, and how they would respond to the artificial confines of a test environment. When faced with such questions, some, including us (Gerlai and Clayton, 1999) have argued that perhaps focusing on natural behavior and factoring in species-specific characteristics, the ecological demands, and evolutionary past of the study species may be helpful. For example, knowing what is rewarding or negatively reinforcing for zebrafish under natural conditions will help us find how to motivate our subjects in a learning task. Similarly, knowing what stimuli zebrafish can perceive, and how zebrafish respond to these stimuli will help us provide the right type of stimulus in the learning task. Last, knowing the behavioral repertoire of zebrafish, that is, its motor responses will also be crucial in the proper design of experiments. Although these statements may appear trivial, the following pages will demonstrate that the task to meet these goals is far from simple. Nevertheless, I hope that these pages will also show that the development of high throughput, simple, yet efficient associative learning tasks for zebrafish is possible.

B. What Motivates Zebrafish? Motivation is a crucial question in learning tasks. Without appropriate reinforcement, learning, although it may occur, cannot be demonstrated under experimental conditions. There are primarily two main classes of reinforcers: negative and positive. Negative reinforcement, or punishment, is often a powerful

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motivator. Positive reinforcement has also been successfully employed in animal research in, so-called, appetitive (reward based) tasks. Food appears to be a trivial choice as a positive motivator (Berridge, 1996). All animals need food to sustain their homeostasis and thus should be compelled to obtain it. The question, however, of how well food may be employed as a reinforcer for zebrafish appears somewhat controversial. Given the small number of studies conducted on this subject it is difficult to decide how well food may be employed as a motivator in learning tasks. The issue is that the small and cold-blooded zebrafish can eat and satiate so quickly that, unlike rodents that require a steady supply of energy, the fish remain well fed for prolonged time periods even after a single and short episode of eating. We have experienced significant difficulties with food rewardbased learning tasks (Gerlai, unpublished results); our experimental zebrafish started to acquire the association between the food reward and a CS (a red cue card in this case) but often stopped performing after a couple of days, presumably due to being satiated and no longer motivated to seek out food. The food we utilized was a gelatin-based commercial fish food (gellybelly) and once we meticulously limited the amount of food delivered at each trial, we were able to make the task work (Sison and Gerlai, 2010). The nauplii of Artemia (brine shrimp), a favorite of many aquarium fish including zebrafish, may lead to better success. Given the small particle size of this food item, it is likely that satiation will be less of a problem with it. Once appropriate delivery methods (precise localization of delivery and nauplii count) have been worked out (nauplii are delivered live and can swim or float away from the delivery location), this food item may have good utility in appetitive conditioning tasks for zebrafish. Briefly, systematic analysis of the type and amount of food and the mode of food delivery will significantly contribute to the development of successful appetitive learning tasks in zebrafish. Although this goal should be attainable, we decided to take a short cut and employed a somewhat unorthodox positive reinforcer, the sight of zebrafish itself. The zebrafish is a highly social species (e.g., Miller and Gerlai, 2007; Saverino and Gerlai, 2008); individuals tend to aggregate, a behavior termed shoaling. Briefly, zebrafish prefer to stay in close proximity to each other. A single fish, thus, is expected to seek out and join a group of zebrafish. Indeed, this is the natural behavior we have utilized in our learning task (Al-Imari and Gerlai, 2008). We have shown that the sight of conspecifics is enough to motivate zebrafish in a simple associative learning task (Al-Imari and Gerlai, 2008). The advantage of this motivator is that zebrafish do not appear to ‘‘satiate’’ with it, that is, the motivation to seek out and attempt to join a group of conspecifics remains stable over the extended periods of time and across multiple repeated trials. Negative reinforcers have also been successfully utilized in studies of animal learning (Gerlai, 1998). However, with zebrafish, our knowledge of the utility of such reinforcers is extremely limited. Future studies will tell whether and what punishment may work best in learning tasks designed for zebrafish. Nevertheless, we (Bass and Gerlai, 2008; Gerlai et al., 2009a; Parra et al., 2009; Speedie and


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Gerlai, 2008), as well as others (Blaser et al., 2010; Jesuthasan and Mathuru, 2008), have started the characterization of the behavioral responses to stimuli that are assumed to be fear inducing. These negative reinforcers may turn out to be appropriate for punishment-based associative learning tasks. For example, a piscivore (fish predator) believed to be sympatric with zebrafish, the Indian leaf fish (Nandus nandus), has been shown to induce significant fear responses in zebrafish (Bass and Gerlai, 2008), and the computer animated (moving) image of this predator has also been found equally effective (Gerlai et al., 2009a) (Fig. 1). The latter finding is noteworthy as it suggests that precise and consistent delivery of a negative stimulus, both in terms of its location and timing, should be possible. Other forms of punishment may also be available. The natural alarm substance of zebrafish has been shown to induce fear responses efficiently (Hall and Suboski, 1995; Speedie and Gerlai, 2008), and most recently a synthetic variety of this alarm substance has been found to be effective in zebrafish (Parra et al., 2009) (Fig. 2). Last, electric shocks have also been employed in learning tasks for fish, including zebrafish (Blank et al., 2009). Although the simplicity of delivery of this punishment makes this reinforcer appear quite practical, I would advise against it for two reasons. It is a rather artificial stimulus and as such it may evoke unexpected reactions in zebrafish, and more importantly, the electric current injected into the tank of the experimental subject may affect brain function directly. Simply put, injecting electricity into the brain is not a good idea when one wants to analyze learning-induced electrochemical (and underlying molecular) changes.

[(Fig._1)TD$IG]

Fig. 1 The sympatric predator of zebrafish (Indian leaf fish, Nandus nandus) elicits a robust antipredatory behavior, elevated number of jumps. Panel A shows a comparison of the effect of live stimulus fish. Note that sympatric harmless fish (the Giant danio) or allopatric harmless or predatory fish (the Swordtail fish and the Compressed cichlid) do not induce the behavior. Note that the stimulus fish were presented continuously during the 10 min period of behavioral recording. Panel B shows the effect of a computer animated (moving) image of the sympatric predator (Indian leaf fish). Here the image is shown for three 1 min periods (black filled circles) separated by 3 min no stimulus intervals (white unfilled circles). For further details, methods, and results, see Bass and Gerlai (2008) and Gerlai et al (2009a).

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[(Fig._2)TD$IG]

Fig. 2

Both the natural (Panel A) and the synthetic (Panel B) alarm substance increases the number of erratic movement episodes in a dose-dependent manner in zebrafish. Note that for the natural alarm substance only the relative dose (dilution sequence) but not the absolute dose could be established. Also note the chemical structure of the synthetic alarm substance in Panel B. For further details, methods, and results, see Speedie and Gerlai (2008) and Parra et al. (2009).

C. The Perceptual Demands of the Learning Task: Focus on Visual Stimuli One of the reasons why the zebrafish has an advantage over classical rodent laboratory model organisms in the analysis of learning and memory is that rodents are nocturnal whereas the zebrafish is a diurnal species. Being active during the light phase of the photoperiod not only makes zebrafish easier for us to observe, but it also means that this species possesses a highly developed visual system, one which can detect visual cues well (for a recent review, see Fadool and Dowling, 2008). Why does this represent advantage over rodent species? Our own species is also diurnal and as such many of our devices are vision based, including TV monitors and cameras that can be purchased cheaply, and used readily off the shelf for zebrafish. Furthermore, visual stimuli are easy to manipulate; images can be delivered precisely in terms of their location and timing. As mentioned above, computer-generated images of the sympatric predator of zebrafish have been shown to be just as effective as the live predator in eliciting fear reactions in these fish (Bass and Gerlai, 2008; Gerlai et al., 2009a). Similarly, computer-animated images of zebrafish, an artificial shoal, have been shown to induce robust shoaling responses in zebrafish, and thus can be utilized as positive reinforcement (Fernandes and Gerlai, 2009a, 2009b; Gerlai et al., 2009b) (Fig. 3). Although other stimuli, for example chemical (smell of food, or the alarm substance), or tactile (vibration detected by the lateral line of zebrafish) as well as auditory cues may also be useful, I predict that the presentation of visual stimuli will enjoy the brightest future, at least in the analysis of associative learning of zebrafish.


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Fig. 3 Zebrafish exhibits strong preference for the sight of live (Panel A) or animated (moving) computer images (Panel B) of its own species. Panel A shows the amount of time (in percent) (mean SEM) zebrafish spent in the preference area during a 10 min recording session. Experimental subjects were tested singly. The preference area is the gray zone indicated in the plus maze (above the bar graph) that is close to the stimulus tank. The stimulus tank contained five live zebrafish (dotted rectangle). Results are shown for the first and the tenth trial (each trial lasting for 5 min). Note that identical stimulus tanks are placed at each arm of the plus maze (white unfilled rectangles) but these tanks did not have the stimulus fish in them. Also note that during both trials the experimental subject spent significantly more time than random chance (18%, indicated by the horizontal line) near the stimulus tank. Panel B shows the response (mean SEM) to the animated images of zebrafish presented on one of the computer screens placed on the side of the tank (black-sided gray rectangles). Experimental fish were tested singly and each of them received the stimulus presentation on only one side (one of the screens), but which side it was varied randomly from subject to subject. The zebrafish images mimicking a natural shoal were shown from the tenth to twentieth minute (black symbols) of a 30 min behavioral session. The images were not shown during the first and the last 10 min of the session (gray symbols). Note the robust decrease of the distance from the stimulus screen during image presentation. For further details, methods, and results, see Al-Imari and Gerlai (2008) and Fernandes and Gerlai (2009).

D. What can Zebrafish Learn: Simple Two Cue Association versus Complex Relational Learning? In the simplest forms of associative learning tasks, the temporal contiguity between the US and the CS is learned. This form of learning has been demonstrated in fish (Woodard and Bitterman, 1973) including zebrafish (Sison and Gerlai, 2010). For example, in a plus-shaped maze (which had four arms and a center starting location, Fig. 4) zebrafish were able to learn that a red cue card predicted the presence of food (Sison and Gerlai, 2010). The task was as follows. Zebrafish were extensively habituated to the maze environment, a point I will return to later, and subsequently were allowed to explore the maze singly, that is, one fish at a time. Fifty percent of the subjects (the ‘‘paired group’’) had access to food (gellybelly, delivered

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Fig. 4 Simple associative (Panel A) and complex spatial learning (Panel B) performance in zebrafish during a probe trial that followed a 20 trial training. Fish were trained singly, and during regular training trials the fish had access to food from one of the food delivery stations (gray rectangles at the end of the arms of the plus maze). However, during the probe trial, no food was given, and the responses shown here are elicited by the conditioned stimulus (CS) alone. Amount of time (in percent) in the target arm is significantly higher (** p < 0.01) in fish (paired group) that were presented a food reward (unconditioned stimulus or US) and a visual cue (CS) together as compared to fish (unpaired group) for which these stimuli were given randomly (Panel A). Amount of time in the target arm is also significantly higher (* p < 0.05) for fish that were presented the food (US) at a fixed (unmarked) location (CS) (paired group) as compared to fish that received the food at random locations (Panel B). Chance level performance is indicated by the horizontal line. Mean SEM are shown. The test apparatus is shown above the graph. The center starting box is shown by the black rectangle. At the end of each arm there is a food delivery station (gray rectangles) behind a perforated acrylic screen (broken lines). Only one of these delivery stations provides access to food but all give the same olfactory cue. For further details, methods, and results (see Sison and Gerlai, 2010).


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through a 1 mm diameter Teflon tube) at the end of one of the arms of the maze where the delivery station was marked by a red cue card. The other arms had identical delivery stations and also provided the smell of food but no access to it, and these locations were not marked by the red cue card. Where exactly the food was delivered, that is, which arm of the plus-maze was the correct choice, varied randomly from trial to trial (extra- and intramaze cues were made irrelevant), but the baited arm was always marked by the red cue card. The other 50% of the fish received the same training procedure as the paired group, except that for them the cue card and the accessible food were not presented together (the ‘‘unpaired group’’). That is the location of the food and the location of the cue did not covary but was random with respect to each other. A day after 20 training trials, zebrafish entered a probe trial. During the probe trial no food was delivered and the cue card was placed in one of the arms. Zebrafish in the paired group showed significant preference for the cued arm, that is, spent significantly more time near the cue card than anywhere else in the maze, whereas fish in the unpaired group showed no such preference and were at random chance, that is, distributed their location evenly in all parts of the maze (Fig. 4). These results demonstrated that zebrafish are able to learn and remember the association between the food reward and the cue card. It is not surprising that zebrafish can acquire the association between the US and CS. But in nature, these fish may also be faced with more complex tasks. For example, remembering where appropriate hiding places are located, where predators are more frequent, or where insects are most abundant may all have a high fitness value. Briefly, the ability to learn and remember spatial information (the location of things) may be important for the survival of the individual. Spatial learning is a complex form of associative learning whereby the animal (or human) needs to acquire the association among loosely related bits and pieces of information. This type of learning is called relational because performance in these tasks is dependent upon establishing the relationships among multiple stimuli, in this case landmark cues. Briefly, spatial learning entails the acquisition of a map, the dynamic relationship of visuospatial cues. Good spatial learning performance has been already demonstrated in zebrafish (Sison and Gerlai, 2010). The paradigm was very similar to the simple associative task conducted in the plus maze described above. The main difference here, however, was that instead of marking the location of the food reward with a salient visible cue, the food delivery station was not marked. But instead, food was delivered in a fixed location relative to external visual cues. That is, the maze was rotated from trial to trial and the arm in which the food was delivered was randomized, but the spatial location, relative to the external visual cues found in the experimental room where the maze was, remained constant. In tasks such as this, the assumption is that successful navigation through the maze requires acquisition of the dynamic relationships of external visual cues, because the view of these cues changes at every point in the maze as the subject navigates in the maze. Zebrafish performed in this spatial task just as well as in the simple associative (two cue association) learning task (Sison and Gerlai, 2010), suggesting that this species is capable of relational learning (Fig. 4).

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E. How Can We Make Associative Learning Tasks High Throughput? The learning tasks discussed above have proven to be successful and attest to the learning capabilities of zebrafish. However, maze navigation-based learning tasks, especially spatial tasks, are notoriously slow and labor intensive to use (Gerlai, 2001). In the case of zebrafish, the experimenter needs to place the subject in the maze individually, and the training trial takes several minutes (5–15 min). But more importantly, the trials must be repeated multiple times (in the examples above, 20 times) to achieve a robust enough learning performance (acquisition of the association), and each time the fish need to be handled individually (removed from the home tank and placed into the test maze and then returned to the home tank). Briefly, these learning tasks are not appropriate for high-throughput screening. Recently, however, a novel maze-based learning task for zebrafish has been published that may be appropriate for this purpose (Go´mez-Laplaza and Gerlai, 2010). The task is termed ‘‘latent learning paradigm’’. Latent learning has been somewhat of a conundrum for classical psychology, because in these tasks animals learned seemingly without any external reinforcement. In latent learning, the experimental subjects were almost always exposed to some form of novelty, novel cues, new places, or new objects. Later it turned out that these tasks are not principally different from those employed in the classical animal psychology laboratory; they also provided an external reinforcer, novelty itself. Exploratory behavior has been shown to be highly adaptive in multiple species from fish (Gerlai et al., 1990) to mammals (Crusio and van Abeelen, 1986). Exploring novel places and objects presumably allows the subject to gather vital information about escape routes, sources of food, location of competitors, and potential mates, etc. Exploration, just like learning, appears to be a ubiquitous feature of most animal species. The recently published latent learning paradigm utilizes this exploratory drive, the motivation to seek out novelty. The paradigm was as follows. Groups of zebrafish (10 subjects at a time) were allowed to explore a complex maze (Fig. 5). The maze, made from transparent acrylic, consisted of a start chamber, and left, center, and right tunnels that connected the start chamber to a goal chamber. No reward or punishment was delivered in the maze, and the fish could freely swim in it, except that one group could swim to a goal chamber (see below) only via the left tunnel (while the right and center tunnels were closed by a guillotine door), another group could swim to the goal chamber only via the right tunnel (while the left and center tunnels were closed by a guillotine door), and fish in the third group were not exposed to the maze. Specifically, fish in groups 1 and 2 were exposed to their respective training environment for 1 h once a day for 16 consecutive days. After training, each zebrafish was tested singly in a probe trial, during which both the left and the right tunnels were open and the goal chamber contained a group of stimulus fish, conspecifics equal in size and identical in color to the test fish. Importantly, during probe trial, the experimental subject could view the stimulus fish from any part of the maze (the maze was transparent) and could get to the goal chamber via either tunnel. Which tunnel did the fish choose? According to the expectations, fish that were allowed to explore the maze with the left tunnel open


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Fig. 5 Latent learning in zebrafish. Fish were exposed to a complex maze shown on the left, equipped with guillotine doors (broken lines). The animals (group of ten each trial) were started from the start box and were allowed to explore the maze via particular routes as described in the text and in Go´mez-Laplaza and Gerlai (2010). During training no external motivation was provided. During the probe trial (bar graph on the left) each experimental fish was tested singly and the reward chamber contained stimulus fish (conspecifics) that were clearly visible from all directions in the maze. The number of fish choosing the left versus the right tunnel was counted. Binomial test indicated a significant left bias in left tunnel open trained and a significant right bias in the right tunnel open trained fish, whereas naı¨ve fish were at random chance (broken horizontal line). For further details, methods, and results (see Go´mez-Laplaza and Gerlai, 2010).

chose this route significantly more frequently, and fish that experienced the right tunnel open chose this rout preferentially, whereas fish that had both arms open chose at random (Fig. 5). In summary, prior exploration of the maze, led to a significant spatial bias, acquisition of the routes available, that is, spatial memory. Why is this paradigm high throughput? Although training took more than 2 weeks, multiple fish were trained at a time, and in principle multiple apparatuses could be used in parallel. In fact one could envision a rack system in which tens of such mazes are used and hundreds of fish are trained at the same time. The only part of the task that requires careful monitoring and analysis is the probe trial in which the spatial navigation of the single subject is video recorded. But this trial lasts only for 10 min, and with the use of video tracking systems (Go´mez-Laplaza and Gerlai, 2010), multiple mazes can be monitored at the same time. Within less than 3 weeks, the learning performance of hundreds of fish may be analyzed. Another potentially useful high-throughput application for zebrafish is a shuttle box paradigm that has also been recently published (Pather and Gerlai, 2009). In this

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task zebrafish learn the location of a reward, the sight of animated computer images of zebrafish, a ‘‘synthetic shoal’’. The task is very simple. A single subject is placed in a rectangular fish tank that is equipped with computer monitors placed next to the left and the right side of the tank. After a habituation period, the subject is presented with the synthetic shoal on one side for 20 s. This stimulus period is followed by a 90 s no-stimulus period during which both computer screens are blank (black). The no-stimulus period is followed by another 20 s stimulus presentation period. The stimulus and no-stimulus periods continue to alternate 30 times. The task is run in three different modes. One, the stimulus is presented on alternating sides (left, right, left, right, etc.); two, it is presented on the same side only (e.g., left, left, left, etc.); and three, it is presented in a random manner (left, left, right, right, right, left, right, left, left, left, etc.). A single subject in an empty tank is highly motivated to seek out and stay close to conspecifics. The stimulus presentation thus achieves two goals. One, it allows the fish to habituate to the test environment faster, making it swim actively as opposed to eliciting freezing or other fear responses. Two, it provides sufficient motivation (reinforcement) for associative learning to occur. The most interesting presentation mode is the systematically alternating side presentation. Under this schedule, zebrafish are expected to learn that once the synthetic shoal is gone, it should reappear on the opposite side of the tank. Thus, as zebrafish are acquiring the task, they are expected to move increasingly sooner away from the side where the stimulus has just been shown during the no-stimulus period. Indeed, this is what we found. Within a few training trials, zebrafish significantly decreased the amount of time they spent near the side where the stimulus had just been shown (Fig. 6). Because during the no-stimulus interval no cues could guide the fish as to where the synthetic shoal would reappear, this response demonstrates that zebrafish learned the task and remembered that the shoal would reappear always on the opposite side relative to the previous location. This behavioral response was not found in the group exposed to the random side sequence, and those fish that received the synthetic shoal on one side only tended to stay on that side. Why are these results remarkable? First, one could argue that the good learning performance under the alternating side stimulus presentation schedule is an indication of temporal learning, that is, the experimental subjects learned when and where the synthetic shoal would appear. This, however, may not necessarily be true because the good learning performance of the fish could be due to a simpler process. The disappearance of the synthetic shoal on one side may be regarded as the CS with which the US (the rewarding stimulus, that is, the reappearance of the shoal) is associated. That is, the task can be solved without precise temporal encoding, but instead by establishing the association, with a slight delay, between CS and US (a training technique called ‘‘trace conditioning’’). Irrespective of how zebrafish solve this task, another important point is that this task is fully automated. Stimulus delivery is computerized, as is recording of the animal responses (video tracking or photocells). The subject does not need to be handled (fished out and placed back) between trials, and the entire paradigm (with 30 nostimulus intervals) takes less than an hour. Due to computerization, the compact size of the test (the test apparatus is a 40 L aquarium with two flat screen monitors), and


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Fig. 6 Shuttle box learning performance suggests good associative learning in zebrafish. Mean SEM are shown. Zebrafish were tested singly in this task. Panel A shows the performance of fish that received an alternating sequence of stimulus presentation. For 20 s a group of animated (moving) images of zebrafish were shown on one of the computer screens (black-sided gray rectangles). Following the stimulus presentation, the images were turned off for 90 s and reappeared for another 20 s on the opposite side, and this alternating sequence continued 30 times. The data show the performance during the 90 s no stimulus periods (interstimulus intervals). The significant decrease of time on the side of the previous stimulus presentation shows that zebrafish have learned to expect the appearance of the stimulus on the opposite side. Panel B shows the results when the experimental fish were presented with the image on randomly changing sides. These fish show no change in their behavior. For further details, methods, and results, see Pather and Gerlai (2009).

the fact that all components are commercially available in the public domain (videoequipment is available from any electronics store and behavioral monitoring software is available from several commercial vendors), the task is easy and inexpensive to scale up. The only step missing before its use in high-throughput screening is the optimization of its parameters, that is, the systematic analysis of what works best for the temporal parameters of stimulus delivery, the characteristics of the shoal stimulus, and the physical parameters of the test apparatus, to mention but a few features.

F. The Screening Strategy The examples above should by now persuade the reader that high-throughput screens are feasible for associative learning related characteristics. However, up

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until now we conveniently ignored the fact that learning cannot be measured per se. What we measure in learning tasks is not learning, but learning performance. The distinction may appear semantic at first glance, but it runs much deeper. Whether learning has occurred and whether the subject remembers something can be inferred only from its behavior, that is, its movement in space and time. Any change that alters the ability of the subject to perceive or to respond to stimuli may alter its learning performance. That is, altered learning performance is not necessarily an indication of altered learning or memory (Gerlai, 2001). It may be due to abnormal perception, abnormal motor function, or altered motivation. How can one be certain that in a large scale associative learning screen the mutants that perform differently from normal wild-type animals indeed have a learning alteration? This is a complex question, one that has been debated for decades in the behavioral brain research and neurobehavioral genetics literature (Gerlai, 2002). There are no simple solutions, but how one tries to address this problem will determine the screening strategy one needs to employ. There are two principally different screening strategies we need to consider: the bottom up versus the top down approach (Gerlai, 2002). In the bottom up approach, the investigator first screens for trivial performance characteristics. If a mutationinduced change is found, it is not considered useful for further investigation at least in the sense that this mutant is not considered a potential candidate for identification of genes involved in learning processes. Those mutants that pass the performance evaluation go to a secondary, more complex screen in which basic forms of learning are analyzed. Mutants that are found at this level may be interesting learning mutants. A third-level screen of those mutants that showed no abnormalities in the first two levels may also be conducted. At this level, particular complex forms of learning (e.g., relational vs. simple associative learning) or particular phases of memory (e.g., consolidation vs. retention or recall) may be analyzed specifically to identify mutants that suffer only from alterations in some of these specific processes. The systematic aspect of the bottom up approach is its advantage, but it requires a lot of work. An alternative strategy is the top down approach, which works the opposite way. First, a complex learning paradigm, say a relational learning task, is employed. If a mutation-induced change is detected, follow up analyses are conducted using offspring of the mutant animal to ascertain what may be behind the alteration, a simple performance abnormality, or altered cognitive or mnemonic processes. I suggest, the top down approach better suits zebrafish forward genetics. It appears more efficient in the sense that one does not waste time screening and finding mutants that are unrelated to the main goal of identification of genes involved in associative learning. It also fits the breeding strategy of forward genetics because a phenotypically deviant subject would need to be bred to confirm heritability of its alteration anyway. Another important point to consider is that there are no specific tests for all specific behavioral phenomena. For example, although some tasks may be more or less sensitive to particular forms of learning, they may not be sensitive only to alterations in that form of learning. The argument is similar to what we have discussed above regarding performance factors. Thus, it is recommended


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that one employs a battery of tasks, a series of behavioral tests that tap into the same or similar form of learning, say relational learning, but utilizes different performance characteristics (e.g., one is based on active motor responses, while the other requires passivity, or one is associated with reward, while the other with punishment). If a mutant shows a similar level of alteration across the test battery consisting of learning tasks with different performance demands, one can be more certain that it is the common feature, say the relational aspect of these tasks that is affected by the mutation. Notably, the discussion above is only for the future. Currently, zebrafish researchers do not have enough behavioral test paradigms to populate their screens. Nevertheless, it is important to bare the importance of multiple tests in mind as these tests are being developed. Another important point to consider here is the breeding strategy for mutagenesis screening. This question has been thoroughly covered by others in the literature (e.g., see Guo, 2004; Patton and Zon, 2001) and also in this book. Nevertheless, I would like to briefly mention it here as it specifically pertains to the identification of mutants with altered associative learning phenotypes. The first question is whether we can expect dominant negative mutations. We define dominant negative here as mutations that are not completely recessive and that lead to phenotypical changes even in heterozygotes. The mammalian literature suggests that indeed such mutations do exist for learning and memory related phenotypes (Gerlai et al., 2001 and references therein), and thus we may expect them in zebrafish, too. It is also likely, however, that several mutations of genes associated with learning and memory may be fully recessive and do not manifest in a heterozygous phenotype. Given that retroviral and ethyl nitroso-urea-induced mutations are expected to affect only one allele, that is, will be in a heterozygous form in the population one screens, they may not be detectable at the phenotypical level. In this case, an additional round of breeding is necessary to generate a segregating generation in which homozygous recessives (as well as phenotypically wild-type homozygotes and heterozygotes) will be found. Another issue concerns the fact that behavior is variable. Although error variation in learning tasks is somewhat reduced by the repeated trials, one may still need to worry about whether a single mutant will be detectable against the backdrop of environmental noise. A solution to the problem is again to breed further, that is, to create a population (e.g., by crossing every mutant to a wild-type animal and then backcrossing the offspring to the original mutant) in which the same mutation is represented in multiple individuals. I advocate this approach because I suspect it will be crucial in the identification of mutants exhibiting alterations in such complex behavioral phenomena as associative learning. G. Memory: A Blank Slate By now it has become clear that memory is not a unitary process; it has multiple types and multiple phases (Squire, 2004; Sweatt, 2010). Unfortunately, however, memory has not been systematically analyzed using zebrafish. We do not know how

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long zebrafish may remember and whether the strength of memory is dependent upon the type of learning task employed or other factors. We also do not know whether memory in zebrafish has mechanistically distinguishable phases (short, medium, and long term) and whether the mechanisms of these phases are similar to those discovered in mammals. Some pioneering results, however, suggest that zebrafish is not so different from mammals. For example, long-term potentiation, a synaptic phenomenon believed to subserve memory, has been demonstrated in zebrafish (Nam et al., 2004), and inhibition of memory consolidation by antibodies against cell adhesion molecules has been achieved suggesting homologous mechanisms between mammalian and zebrafish memory processes (Pradel et al., 1999). H. The Devil is in the Details This short chapter is not meant to be a detailed collection of recipes on how to conduct associative learning tasks with zebrafish. Nevertheless, some practical issues may be mentioned. These are often ignored or considered not important to describe in peer-reviewed publications, yet can make a difference between a successful behavioral study and complete failure. I suspect many such issues will come up as zebrafish behavioral studies progress. Here I only mention a couple of them, which I have found particularly important and/or problematic. The first one is the habituation of the experimental subjects. Many learning tasks require active behavioral responses. Under aversive conditions, however, zebrafish often exhibit passivity, freezing, or even worse, erratic movement, a classical escape reaction that is believed to be adaptive because it leads to the stirring up of debris on the bottom of the small lake or stream in which zebrafish live in nature (e.g., Speedie and Gerlai, 2008). Fish that perform such reactions almost certainly will not attend to the associative cues and will not be motivated to learn appetitive (reward based) tasks. Thus, habituating the experimental subjects to the handling and experimental procedures as well as the physical environment of the test situation is an absolute must. The problem is that the method of proper habituation is difficult to describe. At minimum, fish should never be chased with a net, and removal from the water must be minimized or completely avoided (fish can be gently cornered by a net and guided into a beaker, or a beaker could be placed under the net before the fish is removed from its home tank). Making sure that fear reactions do not develop requires patience and a lot of practice. In our laboratory, we also follow a meticulous habituation protocol whereby we place multiple fish at a time in the test tank. Most learning paradigms require testing single subjects, but zebrafish are highly social and find isolation, especially in a novel environment, particularly aversive. Thus, the initial employment of multiple subjects during the habituation trial eases the stressful nature of the situation. Following the first habituation trial, in the subsequent habituation trials, we gradually decrease the number of subjects until a single subject is exposed to the task alone (Sison and Gerlai, 2010). Whether habituation, that is, minimizing experimental handling and novelty-induced fear, is required for aversive (punishment-based) conditioning paradigms remains to be established. Nevertheless, it is likely that aversive


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conditioning paradigms will also require the habituation described above if one needs to contrast and compare pre-fear stimulus and post-fear stimulus behavioral responses. Among the many ‘‘small’’ practical issues, another one concerns how to detect the location and movement patterns of fish. Manual behavioral quantification techniques are less prone to technical errors, but may be subjective and are definitely very labor intensive (Blaser and Gerlai, 2006). Computerized (automated) methods such as video tracking are superior in terms of sophistication and their ability to quantify motor responses on a continuous scale (speed, turn angle, etc.), and because the experimenter does not have to sit in front of every fish tank or monitor, such techniques allow scaling up the tests, that is, facilitate high throughput. The problem with video tracking, however, is that it may be sensitive to numerous technical errors associated with image quality. The image quality depends tremendously on lighting conditions. For example, shaded areas and reflections on the glass can all confuse the tracking system. Given that light itself is an important factor in the behavior of fish, it may not be easy to optimize its delivery according to the needs of the tracking system (e.g., in light dark choice paradigms, the subject may not be visible on one side of the tank, or in the case of visual cue delivery experiments, the cue itself can confuse the tracking). It is recommended, therefore, to use a light source that emits in the infrared range not visible to zebrafish with a camera capable of detecting this wavelength of light. Another simple, but often cumbersome issue in video tracking is floating debris or development of bubbles, small errors that show up as altered pixels from frame to frame that may confuse the tracking system. Debris can occur as a result of the use of unfiltered water or simply due to defecation by the fish. Bubbles can develop in the test tank if the water placed in the tank comes from a pressurized source. Tracking systems can compensate for this to a certain degree because some may be set to ignore frame to frame changes under a particular preset pixel count, but it is best to avoid the use of pressurized water (e.g., directly from the faucet), and it is a good idea to filter the system water thoroughly and provide no food to the test subject at least for 2 h immediately before the test is administered. I. Things We Did Not Cover I did not consider in this chapter several important questions. For example, this chapter did not discuss nonassociative learning, for example habituation or sensitization, instrumental or operant conditioning, or learning in the larvae (Best et al., 2008). Nonassociative learning may be subserved by biological mechanisms partially shared with those underlying more complex forms of learning, and thus nonassociative tasks may also be a useful tool for forward genetics or drug screens aimed at discovering mechanisms of learning and memory. Instrumental (or operant) conditioning may also be an excellent tool for zebrafish. Operant conditioning entails reinforcing particular responses performed. As far as I know, this form of learning has not been demonstrated with zebrafish, but with other fish species it has been successfully employed (e.g., Gerlai and Hogan, 1992). Importantly, I also did not review here the learning studies conducted with larval zebrafish (e.g., Best et al.,

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2008). These also may be quite useful for obvious practical reasons. Primarily, one does not need to wait 3 months for the subjects to grow up and instead can use the fish within 5 days from fertilization. However, studying learning that requires repeated trials in fast developing larvae may be problematic; it may be confounded by developmental changes during the period required to acquire the task. The last point I did not cover but want to mention here is an entire field to itself, psychopharmacology. Most past studies and future plans with zebrafish include genetics. However, pharmacological tools may also be successfully utilized, and the behavioral tests developed may also be appropriate not only for mutational analysis, but also for drug screening (Chakraborty et al., 2009).

IV. Summary Zebrafish is gaining increasing popularity in numerous fields of science because this species represents an excellent compromise between system complexity and practical simplicity. Although a relatively complex vertebrate, it is highly prolific, easy, and inexpensive to maintain in a laboratory. An increasing number of behavioral paradigms is becoming available, and, thus, mutation and drugs screens for complex phenotypes including associative learning and memory will soon be a reality. The complexity of these behavioral phenomena and the large number of potential molecular players involved in them will necessitate thorough and systematic screening, and this is where zebrafish may swim into fame. However, as the orders of magnitude larger rodent neurobehavioral genetics field has taught us, the foundation of these screens, the phenotypical (in this case behavioral) testing tools, must be well laid down. The rapidly evolving zebrafish behavioral neuroscience research suggests that, indeed, this foundation is being well built. Acknowledgments Supported by NIH/NIAAA (USA) and NSERC (Canada). I wish to thank all my students and collaborators for their excellent work.

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INDEX

A AB and TL lines of zebrafish craniofacial development in, 237 craniofacial shape analysis in, 234–237 ontogenetic stages within, 240 trajectories of shape in, 238–239 Acetaminophen exposure in adult fish, 214–215 toxicity of, 214 Acid-Schiff staining, 212 Acridine orange staining, apoptosis detection by, 30–31 Adult hematopoiesis blood production, 82 derived from HSCs, 82 site of, 91 Adult hematopoietic sites, histological analyses of, 83 Adult zebrafish regenerative angiogenesis, 186 Alexa-647-labeled tubulin, 7 preparation of, 8 Amphibian embryos vs. zebrafish embryo, 2–3 Anaphase-promoting complex/cyclosome, 22 Angioblast specification genes, 186 Angiogenesis chemical approach for investigating blastemal and fin tip VEGF ligand expression, 190–191 chemical sensitization, 189–190 genetic sensitization, 190 mural cell recruitment, 191 vascular function analysis. See Vascular function analysis, in transgenic zebrafish line zebrafish as animal model for, 182, 184, 189 Anterior lateral plate mesoderm (ALPM), cardiac progenitor migration into, 164 Anthrax lethal toxin entry harmful effects of, 191 zebrafish vascular model for, 192 Anti-acetylated tubulin antibodies affinity for cilia, 53 hair cell staining with, 64 Anti-VEGF antibody drug, 182 Antivin, 206 Aorta, gonads, and mesonephros (AGM), 81

APC/C. See Anaphase-promoting complex/ cyclosome Apoptosis control of, 29–30 definition, 29 detection by acridine orange staining, 30–31 TUNEL staining, 30 Apoptotic cells, 24 Arachidonic acid, 120 arl13b/sco-eGFP-GFP mRNA over-expressants Kupffer’s vesicle imaging using, 49–51 pronephric duct imaging using, 51–52 spinal central canal imaging using, 51–52 arl13b/scorpion-eGFP transgenic fish, cilia imaging using, 52 Arrhythmia, 176 AR staining bka mutant, 229 bone mutants, 230 craniofacial mutants, 230 Arterial occlusion, 199 Associative learning, 250 impaired, CNS disorders associated with, 251 screening strategies, 263–264 Associative learning in zebrafish advantages of, 251–252 disadvantages of, 252 high-throughput learning tasks, 260–263 latent learning, 261 learning tasks design principles, 253 and motivation natural and synthetic alarm substance, 256 negative, 253–255 positive, 254 practical issues, 266–267 shuttle box learning performance, 263 US and CS, 257–259 visual stimuli, 256–257 Atrioventricular canal development errors, 172 markers, 172–173 specification and mutations, 172–173 Atrioventricular valve

271

272

Index maturation and function, time-lapse imaging of, 173 retrograde blood flow and, 174 AVC. See Atrioventricular canal AVV. See Atrioventricular valve B Bardet–Biedl syndrome, 154–155 BBS. See Bardet–Biedl syndrome Biliary differentiation, 209–210 Blastula recipients, transplanting cells into, 90 Blood cell precursors, 77 Blood mutants, 86 Blood production, 82 Blood vessel formation, 182, 184 in mammals and zebrafish, similarities between, 183 VEGFR inhibitor blocking, 183 BMP signaling. See Bone morphogenetic protein signaling BODIPY fatty acid analogs administration to live zebrafish larvae, 125–126 composition, 125 BODIPY fluorophores, 218 BODIPY lipid analogs, 117–118 BODIPY-tagged cholesterol analog, 126 Bone morphogenetic protein signaling role in hepatogenesis, 208–209 role in liver development, 208–209 Bone mutants, 230 Bony fishes, life cycles of, 227 BrdU incorporation, zebrafish embryos, 24, 26–27 C caf1b in zebrafish, 21 Cancer treatment, anti-VEGF antibody drug for, 182 Cardia bifida mutations, 168 Cardiac conduction, optical mapping of, 175 Cardiac conduction system, 174 Cardiac cone, 169 Cardiac defects clinical manifestations of, 162 identification using zebrafish, 162 prevalence of, 161–162 Cardiac function, defects in arrhythmia, 176 cardiac conduction, 176 contractile apparatus abnormalities, 174–176 monitoring in zebrafish embryo, 174

types of, 174 Cardiac phenotypes in zebrafish embryos defects in cardiac function. See Cardiac function, defects in heart shape defects assessment using markers, 170–172 cardia bifida, 168–169 dysmorphic hearts, 172 errors in AVC formation, 172–173 origins, 168 valve function defects, 173 heart size defects characterization techniques, 164 as function of cell number defects, 163–164 and Hh signaling, 164–165 late-differentiating cells, 166 myocardial differentiation, 166–168 Cardiac progenitor cells differentiation and heart size defects, 162, 163 Hh signaling influence on, 164–165 locations of, 163–164 specification of, 166 Cardiomyocytes counting of, 163 differentiation, assay for timing of, 167 differentiation of progenitors into, 164 fluorescence of, 166 movements during cardiac fusion, 168–169 during heart tube elongation, 169–170 wild-type patterns, 169 CCK-RA. See CCK receptor A CCK receptor A, 133 CCK signaling. See Cholecystokinin signaling CD41 (integrin molecule), 78 Cell-counting experiments, 164 Cell cycle role in regeneration, 21 in zebrafish, 20 Cell-cycle inhibitors, 21 Cell division, 31 in zebrafish, 20 Cell proliferation, control of, 19 Centrosome detection protocol, 25–26 CFUs. See Colony Forming Units Chemical biology using zebrafish model, 185 Chemical library screening, 182 zebrafish embryos, 187 Chemically assisted suppressor/enhancer genetic screen, 189–190 Chemical screens impact of, 210

273

Index for liver formation, 211–212 Chemical sensitization of vascular phenotypes, 189–190 Chemical suppressors of zebrafish cell-cycle mutants, 31 mechanism of action of, 32 protocol for identifying, 32–34 Cholecystokinin signaling, 133 Cholesterol biosynthesis and protein lipidation, 121 Cilia apical, 41 definition, 40 functions of, 40–41 microtuble-based core cytoskeleton of, 40 morphology and motility, analytical tools for, 47 motility, 49 Cilia in zebrafish, 41 fluorescently labeled, live imaging of arl13b/scorpion-eGFP transgenic embryos, 52 protein expression, 52–54 scorpion/arl13b-eGFP mRNA overexpression, 49–52 localization of proteins in, 48 markers of, 60–61 SCCs and MCCs, 44 Cilia-related mutant phenotype analysis degeneration of sensory neurons, 58–59 hair cells basal bodies in, 66–67 neuromast, 67 whole-mount staining and imaging, 64–66 kidney cysts, 57–58 left–right asymmetry defects heart positioning, 54–56 using in situ hybridization, 56–57 olfactory neurons, 68 photoreceptor cell layer morphology, 59, 61–63 Ciliary defects in renal epithelial cells, 43–44 Ciliated cells in zebrafish, markers of, 60–61 Cleavage-stage zebrafish embryos, mounting of, 5–6 Clonal methylcellulose-based assays CFU enumeration, 104 hematopoietic colonies, 105 methylcellulose, 103–104 myeloid and erythroid differentiation in, 102 vs. stromal in vitro culture methods, 101 utilization of, 101 CNC cells. See Cranial neural crest cells CNS progenitor cells, lineage analysis of, 21 Colony Forming Units enumeration of, 104

picking and analyzing, 105 Comet Assay, 27–28 Contractile apparatus abnormalities, 174–176 Cranial neural crest cells, 226 Craniofacial bone in AB zebrafish, ontogenetic changes in, 228 Craniofacial morphogenesis classification of, 226 morphological development rates, 238 zebrafish model for, 229 Craniofacial mutants, approach to identify, 230 Craniofacial phenotype, quantitative methods for studying generalized procrustes analysis, 233–234 geometric morphometrics, 231–232 GM shape analysis, 234–235 morphological development over ontogeny, 238 morphological integration, 239–242 quantitative data, 242 shape change rates, 237–238 shape variation analysis, 235–237 statistical test, 232–233 trajectories of shape in zebrafish lines, 238–239 Cyclins, 31 Cytokinesis, 11 cytoreductive drug treatment, stem cell isolation by, 96, 97 Cytosolic phospholipase A2 (cPLA2) mRNA levels and PLA2 enzymatic activity, 118–120 D DAPI-stained double transgenic embryos, confocal imaging of, 166 Definitive hematopoiesis, precursor subsets initiated by, 78 Differential dye efflux, stem cell isolation using, 96 Digestive function screening, multiple reagents for, 133–134. See also Triple screening method DNA content analysis, 22–23 DsRed expression, 163, 166 Dysmorphic hearts, 170–172 E Early mitotic inhibitor 1 (emi1), 21 Embryonic development, cilia role in, 40 Embryonic hematopoiesis, 77 Embryonic vascular response, 186 EMPs. See Erythromyeloid progenitors EMTB-3GFP-expressing embryos, microtubule imaging in

274

Index Alexa-647-labeled tubulin injection, 7 probe introduction, 8 EMTB-3GFP probes affinity of, 10 disadvantage of, 14 Endoderm progenitor differentiation and specification, 206–207 ENS. See Enteric nervous system Enteric nervous system coexpression of neurochemical markers in, 151 composition, 144 development future prospects, 155–156 genetic approaches to studying, 148–149 and Hh signaling, 155 molecular mechanisms of, 149–150 transcription factors implicated in, 144 differentiation, 150–151 early development of, 146–148 expressing neuronal markers, 152 model for human diseases, 153 Bardet–Biedl syndrome, 154–155 Goldberg–Shprintzen Syndrome, 154 HSCR, 154 role in modulating intestinal activity, 152 EnzChek protease reporter, 131 EnzChk protease assay, 134 Eosin staining, 58 Epifluorescence microscopy, 13 Epigenetic regulation in liver development, 209 Erythromyeloid progenitors in definitive hematopoietic program, 76, 78 gata1+lmo2+ cells, 80 isolation, 78 lmo2 gene expression and, 81 specification and differentiation, 78–79 Ezetimibe, 134 F FACS. See Fluorescence-activated cell sorting FACS profiling, 86 Fate-mapping studies, 79, 81, 164 cardiac progenitor cells, 166 heart size defects, 164 fat-free (ffr) larvae metabolic defects in, 127, 130 positional cloning of, 127 FGF signaling. See Fibroblast growth factor signaling Fibroblast growth factor signaling endoderm formation inhibition by, 206

role in liver development, 208–209 Fish assay, 187 FITC-labeled lectins, 96 Flow cytometric assay, 84–85 Fluorescence-activated cell sorting, 22, 212–213 Fluorescence-imaging modalities, 12–14 Fluorescent bead assay, 188 Fluorescent lipid reporters, 126 NBD-cholesterol, 127–128 phosphatidylcholine, 127 phosphoethanolamine analog PED6, 127–128 for zebrafish ffr mutation, 129–131 Forkhead box transcription factors, 206–207 Forward chemical genetic approach, 185 Forward genetic screens, 20 PED6 and NBD-cholesterol, 127 pH3, 21–22 Forward genetic studies in zebrafish, 126 Forward scatter (FSC), 82 Functional in vitro differentiation studies, 80 G Generalized procrustes analysis, 233–234 Genetic ablation, 216–217 Genetic screens, 162 for lipid metabolism, 218 mutant animals with defects, 242 in zebrafish, screening criteria used in, 82 Genetic sensitization of vascular phenotypes, 189–190 Geometric morphometrics, 231–232 craniofacial shape analysis, 242 geranylgeranyl transferase I (GGTI) inhibitor (GGTI-2166), 121–122 glypican 4 (gpc4) and craniofacial organization, 231 GM shape analysis bone mutants, 230 craniofacial mutants, 230 data collection in, 232–233 and TPS analysis, 234 Goldberg–Shprintzen Syndrome, 154 GOSHS. See Goldberg–Shprintzen Syndrome GPA. See Generalized procrustes analysis gridlock phenotype collateral blood flow in, 185 VEGF-A mRNA expression in, 186 grl and VEGF signaling pathway, connection between, 185–186 Gut-looping, bHLH transcription factor in, 207 Gut microbiota, 146

275

Index Gut motility contractions, 151–152 regulation of, 152–153 gutwrencher mutants, ENS neurons and gut motility in, 152–153 H Hair cell analysis basal bodies, 66–67 by histology, 64 by immunohistochemistry, 64 neuromast, 67 whole-mount staining and imaging, 64–66 Hair cell kinocilia, 45 HCT. See Hematopoietic cell transplantation Heart function assessment, 174 Heart shape defects assessment using markers, 170–172 cardia bifida, 168–169 dysmorphic hearts, 170–172 errors in AVC formation, 172–173 origins, 168 valve function defects, 173 Heart size defects characterization techniques, 164 as function of cell number defects, 163–164 and Hh signaling, 164–165 late-differentiating cells, 166 myocardial differentiation, 166–168 Heart tube morphology, 170 Heart, zebrafish cardiac function defects in, 174–176 composition of, 162 shape defects in, 168–174 size defects in, 163–168 Hedgehog signaling and ENS development, 155 myocardial progenitor cell specification, 164–165 Hematopoiesis, 97 adult, 82–86 definitive, 78–81 primitive, 76–78 timing of, 77 transgenic zebrafish lines for studies of, 79 Hematopoietic cell transplantation adult donor cells, 91–94 embryonic donor cells, 87 hematopoietic cell isolation, 88–89 transplantation into blastula recipients, 90 transplantation into embryonic recipients, 89–90

transplantation into 48 hpf embryos, 90–91 strategies for, 87–88 Hematopoietic progenitors clonal methylcellulose-based assays. See Clonal methylcellulose-based assays differentiation, 98 in vitro culture of categories of, 97 stromal, 98–101 Hematopoietic stem cells enrichment strategies Hoechst 33342 dye, 95, 96 irradiation and cytoreductive drug treatment, 96, 97 lectin binding to WKM scatter fractions, 95, 96 mouse monoclonal antibodies, 94, 95 formation of, 81 migration to CHT, 77 multidrug resistance transporter proteins in, 96 transdifferentiation, 91 Hematoxylin and eosin staining, 58, 212 Hemostasis definition, 197 main initiating event in, 199 mechanism of, 198 zebrafish as model to study, 198 Hepatic precursors, markers of, 207 Hepatobiliary differentiation, 210 Hepatogenesis genetic markers of, 207–208 signals involved in FGF and BMP signaling, 208–209 RA signaling, 209 WNT signaling, 208 H&E staining. See Hematoxylin and eosin staining Heterochronic transplantation strategies, 87 Heterochrony, 237–238 Heterozygous craniofacial phenotypes, 230 High-cholesterol diet (HCD)-fed fish, lipoprotein profiles of, 134 High-speed videomicroscopy techniques, 49 High-throughput screens, 263 Hirschsprung’s disease and BBS, 155 clinical manifestations of, 154 pathology, 154 phenotype of GOSHS, 154 phenotype of Mowat-Wilson syndrome, 154 Histological assessment, 212 Histone H3 phosphorylation, 23–24 HMGCoAR. See 3-Hydroxyl-3-methylglutarylCoA reductase

276

Index Hoechst 33342 dye, stem cell isolation using, 95, 96 Homozygous recessive early lethal zebrafish mutants heterozygous craniofacial phenotypes in, 230 rescue of, 230–231 HSCR. See Hirschsprung’s disease HSCs. See Hematopoietic stem cells 3-Hydroxyl-3-methylglutaryl-CoA reductase downstream products of, 121 PGC migration defects and, 120 Hypercholesterolemia, 134 I ICCs. See Interstitial cells of Cajal ICM. See Intermediate cell mass Immunohistochemistry cilia-related defects in photoreceptor cells, 62–63 hair cell analysis, 64 of hair cells, 64 Intermediate cell mass, 78 Interstitial cells of Cajal, 151 Intestinal enterocytes of zebrafish and mammals, 116 Intestinal tract composition, 144, 145 mammalian and zebrafish, comparison of, 145–146 Intravital staining with Nile red, 218 Irradiation, stem cell isolation by, 96, 97 J Jaw morphogenesis, 242–244 K Kaede photoconversion, 167–168 Kidney cell types, 84 Kidney cysts detection in zebrafish, 57–58 Kif1-binding protein and Stmn2 (stathmin-like protein), interaction between, 154 Kif1-binding protein, null mutations in, 154 Kinocilium, 45 Kupffer’s vesicle cilia, 43 imaging, 49–51 definition, 41 functions, 43 KV. See Kupffer’s vesicle

L Laser-induced thrombosis future perspectives, 202–203 normal values and larval stages in, 202 procedure of larvae preparation, 200–201 laser ablations, 201–202 laser setup, 199–200 Laser-scanning confocal microscopy, 13 of GFP and Alexa 647, 8–9 with one-photon excitation, 14 vs. spinning disc confocal microscopy, 14 Laser setup, 199–200 Latent learning in zebrafish, 261 Learning definition, 250 and memory, 250 model organism for studying, 251 zebrafish for studying, 251–252 tasks design principles, 253 Lectin binding to WKM scatter fractions, 95, 96 Left–right asymmetry defect analysis heart positioning, 54–56 using in situ hybridization, 56–57 Lipid metabolism in developing zebrafish bile production and release, 114–115 triacylglycerol, 115 yolk lipids, 114 genetic screens for, 218 larval zebrafish as model of, 113–114 need for whole animal studies of, 112–113 visualization in zebrafish using BODIPY-cholesterol, 126 BODIPY fatty acid analogs, 125–126 fluorescent lipid reporters, 126–131 lipophilic dyes, 122–125 Lipid signaling, during early zebrafish development cPLA2 mRNA levels and PLA2 enzymatic activity, 118–120 PGC migration, 120–122 PGE2 role in, 120 Lipophilic dyes, 122–125 nile red, 125 oil red O, 123 statins, 124 sudan black B, 122, 123 uses of, 122, 123 Live-imaging, cytoskeleton in zebrafish embryos comparison of microscopic techniques for, 12–14

277

Index factors influencing, 3–4 microfilaments in cleaving embryos. See Microfilament imaging, in cleaving zebrafish embryos microtubules in cleaving embryos. See Microtubule imaging, in zebrafish embryos mounting embryos for, 5–6 probes for, 14–15 Liver development analysis protocols chemical screens, 210–212 fluorescence-activated cell sorting, 212–213 histological assessment, 212 epigenetic factors regulating, 209 Liver enzyme tests, 218 Liver function assessment lipid metabolism, 218–219 liver enzymes, 218 Liver injury, models of genetic ablation, 216–217 resection of adult zebrafish liver, 215–216 transient genetically induced apoptosis, 217 zebrafish model of APAP toxicity, 214–215 Liver regeneration, 213 zebrafish and mammalian, parallels between, 214 Lmo2+ cells, full fate potentials of, 88 LSCM. See Laser-scanning confocal microscopy Lymphatic vessel development, 184–185 Lysochromes. See Lipophilic dyes M Mammalian primitive red blood cells, 78 MBT. See Midblastula transition MCC. See Multiciliated cells Mechanical forces and VEGF signaling, 190 Mechanosensory hair cells characterization of, 45 sound wave detection by, 45 Meis1 (marker of eye primordium), 21 MEK1/2 pathway inactivation and anthrax lethal toxin, 192 Memory, 265 and learning, 250 strength of, 266 Methylcellulose clonal assays, 103–104 Metronidazole, 216 Microangiography, 188 Microbiota role in gut development, 146 Microfilament imaging, in cleaving zebrafish embryos FP-tagged probes for, 10–11

problems associated with, 10 by rhodamine-labeled phalloidin, 10 Utr-CH-GFP expression of F-actin imaging and, 12 probes for, 11 Microtubule imaging, in zebrafish embryos by Alexa-647-labeled tubulin, 7 by rhodamine-labeled tubulin, 6–7 by transgenic zebrafish line, 7–8 Midblastula transition, 20 Mitotic marker, whole mount immunohistochemistry with, 23–25 Mitotic spindle detection protocol, 25–26 MO. See Morpholinos Monoclonal markers, 47 Morpholinos, 117–118 Morphological integration assessment by GM and PCA, 240–241 definition, 239 Mouse hematopoietic development, timing of, 77 Mouse monoclonal antibodies, stem cell isolation using, 94, 95 Mowat-Wilson syndrome, 154 Multiciliated cells, 44 Multilineage hematopoiesis, 78 Mural cells progression of, 191 role in blood vessel function, 191 in zebrafish model, 191 Murine YS blood islands, 78 Mutational screens, 82 Myocardial differentiation phases of, 166 timing of, 166 transgenic assays to monitor, 166–168 Myocardial markers, 170 Myocardium–endoderm and myocardium–ECM interactions, 168 N N-acetyl-p-benzoquinone imine, 214 NAPQI. See N-acetyl-p-benzoquinone imine 22-NBD-cholesterol vs. PED6, 127 Negative reinforcers, associative learning, 253–255 Neural crest cells formation, basic mechanisms of, 144 migration into zebrafish gut, 147 Neural keel, 47 Neuromasts, 45–46 Nile red, 125

278

Index Nitroreductase expression and metronidazole, 216, 217 22-[N-(7-nitronbenz-2-oxa-1,3-diazol-4-yl) amino]- 23,24-bisnor-5-cholen-3-ol. See 22NBD-cholesterol Nodal signaling pathway, 206 Nonassociative learning, 267. See also Associative learning Nonhydrolyzable microspheres, 131 Notch signaling EMP specification and, 78 neural crest induction and, 144 role in zebrafish biliary specification, 210 O Oil-red-O (ORO) staining, 123, 212 Olfactory sensory neurons, 46, 67 labeling by DiI incorporation, 68 Onecut factors and biliary differentiation, 210 Ontogeny, morphological development over, 238 P PCA. See Principal component analysis PDGF (platelet-derived growth factor) role in mural cell development, 191 Peanut agglutinin, 96 PED6 (phosphoethanolamine analog) chemical structures of, 128 cleavage by phospholipase A2, 129–131 labeling of fat-free(ffr) larvae, 127, 129 Perivascular support cells. See Mural cells Phalloidin, 64 Phenotype-driven mutagenesis screens, 243 pH3 (mitotic marker), whole mount immunohistochemistry with, 23–25 Phospholipase A2 enzymatic activity and zebrafish development, 118–119 functions of, 118 Photoreceptor cells monitoring cilia-related defects in, 58–59 of vertebrate retina, 44 of zebrafish retina, 44–45 Photoreceptor cells, monitoring cilia-related defects in by immunohistochemistry, 62–63 by plastic histological sections, 58–59, 61 PLA2. See Phospholipase A2 PLCg 1 downstream of VEGFR signaling, 190 PNA. See Peanut agglutinin Positive reinforcement, 254

Postembryonic mutagenesis screens, 228–230 Potato lectin, 96 Primitive hematopoiesis primitive blood precursors and definitive hematopoietic cells, 88 primitive erythroid cell generation, 78 primitive macrophage development, 76, 78 Primordial germ cell (PGC) migration in embryos treated with GGTI-2166, 121–122 loss of HMGCoAR and, 120–121 visualization of, 120 Principal component analysis, 234 Pronephric duct imaging, 51–52 Pronephric glomerulus differentiation of, 43 Pronephros, origin of, 43 PTL. See Potato lectin R RA signaling. See Retinoic acid signaling Reagents and supplies, 34–35 Regeneration studies, zebrafish fin amputation for, 187 Regenerative angiogenesis blockade by VEGFR inhibitor, 187–188 evaluation using angiography assay, 189 Relational learning, 250, 259, 264, 265 Reporter transgenes, 163 Tg(cmlc2:egfp), 166 Tg(cmlc2:kaede), 166 RET coding sequence, mutations in, 154 Retinoic acid signaling, 209 Reverse chemical approach, 183 Rhodamine-labeled phalloidin, 10 Rhodamine-labeled tubulin, 6–7 S SCC. See Single ciliated cells Scl-interrupting locus, 22 SDCM. See Spinning disc confocal microscopy Seahorse, ciliary localization of, 53 Selective plane illumination microscopy, 173 Semitransparent caudal fin, 185 Senescence-associated b-galactosidase, detection of, 28–29 Shape analysis, geometric approach for, 234–235 Shuttle box learning performance, 263 Single ciliated cells, 44 Sip1 transcription factor, mutations in, 154 Skeletal morphogenesis, zebrafish model for, 229

279

Index sm22a (vascular smooth muscle actin gene), 191 Southpaw (spaw), 56 Sox32 overexpression and endoderm development, 206 Spatial learning performance, 259 SPIM. See Selective plane illumination microscopy Spinal central canal imaging, 51–52 Spinning disc confocal microscopy, 13, 14 Statins, 124 Stereocilia, 45 Sterol analogs, 126 Stmn2 (stathmin-like protein) and Kif1-binding protein, interaction between, 154 Stromal culture assays, 97 in vitro proliferation and differentiation, 100–101 ZKS cells, 98–99 Sudan black B, 122, 123 Suppressor screening, 189–190 T Tg(cmlc2:DsRed2-nuc), 163 Thin-plate spline analysis, 234 Thrombocytes, 198 Thrombosis, 198 Time taken to occlude assay, 199 TL lines of zebrafish. See AB and TL lines of zebrafish TPS analysis. See Thin-plate spline analysis Transgelin. See sm22a (vascular smooth muscle actin gene) Transgenic reporter lines, 213 Transgenic zebrafish line generation of, 7–8 for hematopoietic studies, 79 lineage-affiliated expression patterns in, 85–86 vascular function analysis using. See Vascular function analysis, in transgenic zebrafish line Transient genetically induced apoptosis, 217 Triple screening method exocrine pancreas assessment by, 133 larval digestive function assessment by, 131–133 regulation of phospholipase and protease activity by, 133 TTO assay. See Time taken to occlude assay TUNEL staining, apoptosis detection by, 30 Two-photon microscopy (2PM), 13 U Utr-CH-GFP expression, 11 Utrophin-based labeling, 11

V Vascular endothelial growth factor functions of, 182 Vascular function analysis, in transgenic zebrafish line advantages of, 184 blood vessel formation, 184 endothelial cells, 185 lymphatic vessels development, 185 Vascular mutation analysis, 185–186 Vascular occlusion causes of, 199 procedure of larvae preparation, 200–201 laser ablations, 201–202 laser setup, 199–200 laser thrombosis, 202 Vascular permeability and extravasation, fluorescent bead assay for, 188 Vascular remodeling and blood flow, 190–191 Vascular smooth muscle cells, 191 Vasculature development, zebrafish and chemical approaches adult angiography, 189 blood and lymphatic vessels, 183–184 chemical library screening, 187 chemical treatments, 186–188 fluorescent bead assay, 188 microangiography, 188 vascular events, 184–185 vascular mutations, 185–186 hemodynamic forces effects on, 190–191 similarities with other vertebrates, 183 VEGF. See Vascular endothelial growth factor VEGFR inhibitor PTK787/ZK222584, 183 regenerative angiogenesis blockade by, 187–188 VEGFR signaling pathway downstream components of, 183 PLCg 1 downstream of, 190 VEGF signaling pathway aortic arch blood vessels remodeling and, 190 and grl, connection between, 185–186 mural cell development and, 191 reverse chemical approach to investigate, 183 strategy to block, 182 Venous thrombosis, 202 Vertebrate genomes, 251 Visual stimuli, 256–257 vlad tepes mutant and WKM transplantation, 88

280

Index W Whole kidney marrow analysis by FACS blood lineage, 84–85 fluidics and beam size, 84 FSC and SSC, 82, 84 scatter profile, 84–85 cellular compositions of, 83 differential binding of lectins, 94, 95 immune response, 94, 96 transplantation, 92–93 and vlad tepes mutant, 88 WKM. See Whole kidney marrow WNT signaling, 155 inhibition and regenerative index, 215 role in liver development, 208 X Xenopus laevis, 10 Y Yolk metabolism, early vertebrate development apoc2 functions in, 118 gene expression and, 115 lipid metabolism genes in, 117–118 zebrafish YSL and, 115, 116 Yolk syncytial layer, 114 lipid metabolism genes role in, 117–118 YSL. See Yolk syncytial layer Z Zebrafish breeding competence, maintaining, 4 caf1b in, 21 caudal or tail fin, 187 cell cycle in, 20 cell-cycle machinery role in tissue differentiation, 21 in chemical screens, 32 cilia function studies using, 41 for developmental embryology studies, 20 inner ear of, 45 live-imaging of cytoskeletal dynamics in. See Live-imaging, cytoskeleton in zebrafish embryos mechanosensory hair cells, 45–46 neural keel, 47 olfactory system, 46 studies of developing, 20

Zebrafish blood vessels in larval development, 184 Zebrafish craniofacial mutants, 227 Zebrafish definitive hematopoietic differentiation, 84 Zebrafish development PGE2 role in, 120 PLA2 enzymatic activity role in, 118–119 Zebrafish embryo cell-cycle protocols apoptosis detection by acridine orange staining, 30–31 by TUNEL staining, 29–30 cell proliferation analysis BrdU, 26–27 DNA content, 22–23 mitotic spindle and centrosomes, 25–26 whole mount immunohistochemistry, 23–25 DNA damage analysis, 27–28 senescence analysis, 28–29 Zebrafish embryogenesis analysis of, 41 developmental stages of, 21 Zebrafish embryos cell-cycle regulation in, 20–21 techniques for study of, 24 characterization of, 3 cytoskeletal dynamics in live-imaging. See Live-imaging, cytoskeleton in zebrafish embryos heart positioning evaluation in, 54–56 microinjection, 8 for molecular–genetic analysis, 2–3 mounting of methods for, 5–6 rationale for, 5 phases of heart formation in, 163 in situ hybridization of antisense probes in, 31 Zebrafish kidney stromal (ZKS) cells, 97 Zebrafish larvae concurrent feeding of, 131–133 lipid uptake visualization, 127, 128 preparation for laser injury, 201 Zebrafish models of human dyslipidemias, 134–135 Zebrafish mutants, digestive tract fluorescence in, 131 Zebrafish oocytes cytoskeletal components of, 3 maternal products of, 3 Zebrafish orthologs of cell-cycle regulatory genes, 31 Zebrafish pronephric duct, 44 Zebrafish sensory organs mechanosensory hair cells, 45–46

281

Index photoreceptors, 44–45 Zebrafish spleen, 82 Zebrafish vasculature development and chemical approaches adult angiography, 189 blood and lymphatic vessels, 183–184 chemical library screening, 187 chemical treatments, 186–188 fluorescent bead assay, 188 microangiography, 188

vascular events, 184–185 vascular mutations, 185–186 hemodynamic forces effects on, 190–191 similarities with other vertebrates, 183 ZKS cells generation of, 98 maintenance and culture of, 98–99 in vitro proliferation and differentiation assays, 100–101 Zpr-1 antibody stains, 59

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Plate 1 (Figure 1.2 on page 7 of this volume)












The Zebrafish: Cellular and Developmental Biology, Part B, Volume 101, Third Edition (Methods in Cell Biology)

The Zebrafish: Cellular and Developmental Biology, Part A, Volume 100, Third Edition (Methods in Cell Biology)

Methods in Cell Biology Volume 25 The Cytoskeleton Part B

Methods in Cell Biology Volume 50 Methods in Plant Cell Biology, Part B

The Zebrafish: Genetics, Genomics and Informatics 3rd Edition-, Volume 104, Third Edition (Methods in Cell Biology)

Methods in Cell Biology Volume 32 Vesicular Transport Part B

Molecular Methods in Developmental Biology. Xenopus and Zebrafish

Molecular Methods In Developmental Biology: Xenopus and Zebrafish

Methods in Cell Biology Volume 52 Methods in Muscle Biology

B Cell Protocols (Methods in Molecular Biology)

B Cell Protocols (Methods in Molecular Biology)

Methods in Cell Biology Volume 24 The Cytoskeleton Part A

Medical Cell Biology, Third Edition (MEDICAL CELL BIOLOGY (GOODMAN))

Molecular Methods in Developmental Biology: Xenopus & Zebrafish (Methods in Molecular Biology Vol 127)

Methods in Cell Biology Volume 63 Cytometry, Part A

Methods in Cell Biology Volume 20

Methods in Cell Biology Volume 11

Methods in Cell Biology Volume 65 Mitochondria

Methods in Cell Biology Volume 70

Methods in Cell Biology Volume 15

Methods in Cell Biology Volume 14

Methods in Cell Biology Volume 31 Vesicular Transport Part A

Developmental Vascular Biology, Volume 62 (Current Topics in Developmental Biology)

Cell Biology, Four-Volume Set: Cell Biology, Volume 1, Third Edition: A Laboratory Handbook

Developmental Biology (9th Edition)

Methods in Nano Cell Biology, Volume 90 (Methods in Cell Biology)

Zebrafish: Methods and Protocols (Methods in Molecular Biology)

Methods in Cell Biology Volume 87 Avian Embryology 2nd Edition

Recent Advances in Cytometry, Part A, Volume 102, Fifth Edition: Instrumentation, Methods (Methods in Cell Biology)

The Zebrafish: Cellular and Developmental Biology, Part B, Volume 101, Third Edition (Methods in Cell Biology)

The Zebrafish: Cellular and Developmental Biology, Part A, Volume 100, Third Edition (Methods in Cell Biology)

Methods in Cell Biology Volume 25 The Cytoskeleton Part B

Methods in Cell Biology Volume 50 Methods in Plant Cell Biology, Part B

The Zebrafish: Genetics, Genomics and Informatics 3rd Edition-, Volume 104, Third Edition (Methods in Cell Biology)

Methods in Cell Biology Volume 32 Vesicular Transport Part B

Molecular Methods in Developmental Biology. Xenopus and Zebrafish

Molecular Methods In Developmental Biology: Xenopus and Zebrafish

Methods in Cell Biology Volume 52 Methods in Muscle Biology

B Cell Protocols (Methods in Molecular Biology)

B Cell Protocols (Methods in Molecular Biology)

Methods in Cell Biology Volume 24 The Cytoskeleton Part A

Medical Cell Biology, Third Edition (MEDICAL CELL BIOLOGY (GOODMAN))

Molecular Methods in Developmental Biology: Xenopus & Zebrafish (Methods in Molecular Biology Vol 127)

Methods in Cell Biology Volume 63 Cytometry, Part A

Methods in Cell Biology Volume 20

Methods in Cell Biology Volume 11

Methods in Cell Biology Volume 65 Mitochondria

Methods in Cell Biology Volume 70

Methods in Cell Biology Volume 15

Methods in Cell Biology Volume 14

Methods in Cell Biology Volume 31 Vesicular Transport Part A

Developmental Vascular Biology, Volume 62 (Current Topics in Developmental Biology)

Cell Biology, Four-Volume Set: Cell Biology, Volume 1, Third Edition: A Laboratory Handbook

Developmental Biology (9th Edition)

Methods in Nano Cell Biology, Volume 90 (Methods in Cell Biology)

Zebrafish: Methods and Protocols (Methods in Molecular Biology)

Methods in Cell Biology Volume 87 Avian Embryology 2nd Edition

Recent Advances in Cytometry, Part A, Volume 102, Fifth Edition: Instrumentation, Methods (Methods in Cell Biology)

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