Maize (Zeamays): A Model Organism for Basic and Applied ... · expansive amplification of...

Emerging Model Organisms, Volume 2 33 Zea mays

1Corresponding author ([email protected]).Cite as: Cold Spring Harb Protoc; 2009; doi:10.1101/pdb.emo132 www.cshprotocols.org

Chapter 2

Maize (Zea mays): A Model Organism for Basic and AppliedResearch in Plant Biology

Josh Strable and Michael J. Scanlon1

Department of Plant Biology, Cornell University, Ithaca, NY 14853, USA

INTRODUCTION

Zea mays ssp. mays is one of the world’s most important crop plants, boasting a multibillion dollarannual revenue. In addition to its agronomic importance, maize has been a keystone model organismfor basic research for nearly a century. Within the cereals, which include other plant model speciessuch as rice (Oryza sativa), sorghum (Sorghum bicolor), wheat (Triticum spp.), and barley (Hordeumvulgare), maize is the most thoroughly researched genetic system. Several attributes of the maizeplant, including a vast collection of mutant stocks, large heterochromatic chromosomes, extensivenucleotide diversity, and genic colinearity within related grasses, have positioned this species as acenterpiece for genetic, cytogenetic, and genomic research. As a model organism, maize is thesubject of such far-ranging biological investigations as plant domestication, genome evolution, devel-opmental physiology, epigenetics, pest resistance, heterosis, quantitative inheritance, and compara-tive genomics. These and other studies will be advanced by the completed sequencing and annotationof the maize gene space, which will be realized during 2009. Here we present an overview of the useof maize as a model system and provide links to several protocols that enable its genetic and genomicanalysis.

BACKGROUND INFORMATION

Zea mays L. ssp. mays, commonly referred to as maize or corn, belongs to the grass tribeAndropogoneae of the family Gramineae (Poaceae). The grasses originated 55-70 million years ago(mya) and subsequently diversified to include all the major cereal crop species in addition to nearly10,000 nondomesticated relatives (Fig. 1A; Kellogg 2001; Bolot et al. 2009). The maize genome orig-inated 4.8 mya via the segmental allotetraploidization of two progenitor genomes that themselvesdiverged from a sorghum progenitor ~11.9 mya (Gaut and Doebley 1997; Swigonova et al. 2004).Although they differ in ploidy and overall genome size, cereal genomes exhibit a relatively high degreeof genic colinearity and sequence conservation (Gale and Devos 1998; Bennetzen and Ma 2003).Much of the size variation within the cereal genomes is attributed to genome duplication and theexpansive amplification of transposable elements (SanMiguel et al. 1996, 1998; Bennetzen 2000).

Deciphering the genetic history of maize in light of its domestication is pivotal to an understand-ing of its natural history (for review, see Doebley 2004). The closest wild relatives of domesticatedmaize are the teosintes, the annual and perennial grasses of the genus Zea that are indigenous toMexico and Central America (Fukunaga et al. 2005). The teosintes include four species: Z. luxurians,Z. diploperennis, Z. perennis, and Z. mays, with Z. mays composed of at least four subspecies (ssp.mays, ssp. mexicana, ssp. parviglumis, and ssp. huehuetenangensis) (Fig. 1B; Fukunaga et al. 2005;Vollbrecht and Sigmon 2005). Genetic studies of maize domestication have identified Z. mays ssp.parviglumis as the direct ancestor of modern maize, and, complemented by archeological data(Piperno and Flannery 2001), estimate that maize diverged from its teosinte ancestor between 6000and 9000 years ago (Matsuoka et al. 2002). In that same study, Matsuoka and colleagues (2002)

02_EVOv2_033-042_font.qxp:8.5 x 10.875 2/4/10 9:34 AM Page 33

Copyright 2010 Cold Spring Harbor Laboratory Press. Not for distribution. Do not copy without written permission from Cold Spring Harbor Laboratory Press


postulate a single domestication event, which likely occurred in southern Mexico’s central Balsas Riverregion. The subsequent diversification of maize took place in the Mexican highlands between Oaxacaand Jalisco provinces. Dispersal of modern maize then proceeded along two principle routes: a north-ern route stretching from northwestern Mexico, through the southwestern United States, and intothe eastern United States and Canada; and a southern route ranging from Mexico’s southwesternlowlands into Guatemala, the Caribbean, and South America (Matsuoka et al. 2002). Zea mays ssp.mays produced a multibillion dollar revenue from the 157 million hectares harvested globally in 2007(http://faostat.fao.org).

The origin of maize as a model organism traces back to early studies performed in 1869 by GregorMendel, where maize was used to corroborate the more renowned breeding experiments he per-formed previously in Pisum (pea) (Rhoades 1984; Coe 2001). Like pea, maize is a large plant, makingit well-suited for phenotypic analysis and thereby conferring a decided advantage in genetic analysisof morphological mutants. Some 30 years later, Correns and de Vries utilized maize extensively in theirinvestigations on xenia—the dominant influence exerted by the pollen parent on the phenotype ofthe endosperm. The results from these studies complemented and extended Mendel’s original geneticstudies and were integral to the rediscovery of Mendel’s landmark work.

While Correns and de Vries were among the early pioneers of maize research, R. A. Emerson ofCornell University and his thesis advisor E. M. East of Harvard University are widely regarded as thefounding fathers of modern maize genetics. Emerson served as mentor to an astoundingly energeticand influential first generation of maize geneticists, including George Beadle, Charles Burnham,Marcus Rhoades, and Barbara McClintock. Throughout the 1920s and 1930s, the Cornell group estab-lished a solid foundation in transmission genetics and cytology that provided a framework for the useof maize as a model genetic system. Detailed historical perspectives on this foundational research inmaize genetics are provided in articles by Rhoades (1984) and Coe (2001).

SOURCES AND HUSBANDRY

The Maize Genetics Cooperation Stock Center (MGCSC) located at the University of Illinois at Urbana-Champaign, is a primary source for maize mutant stocks used in research (Table 1). As a free serviceto the maize community, the MGCSC obtains, maintains, and disseminates seed stocks internationally

FIGURE 1. Evolutionary relationships of maize tocereal crops and within the genus Zea. (A) A par-tial phylogeny of the cereals with Arabidopsis asan outgroup. Divergence of the major cereal cropprogenitors is estimated to have occurred withinthe past 50 mya. Maize and rice diverged ~50mya, whereas maize and sorghum diverged morerecently, ~9 mya. (Reprinted, with permission,from Bolot et al. 2009. © 2009 Elsevier.) (B) Aphylogeny of the genus Zea. After divergingfrom the genus Tripsacum, the ancestral genusZea underwent a relatively recent radiation togive rise to four extant species and at leastfour subspecies. Molecular data indicate Z. maysssp. parivglumis is the closest relative to modernmaize (Z. mays ssp. mays) and that the twodiverged ~9000 yr ago (*). (Reprinted, withpermission, from Vollbrecht and Sigmon 2005.© 2005 The Biochemical Society.)

A

B

02_EVOv2_033-042.qxp:8.5 x 10.875 2/23/10 12:16 PM Page 34



Table 1. Online maize resources

Name Feature/resource URL

Arizona Genomics Institute Maize FingerPrinted Contig (FPC) map, http://www.genome.arizona.edu/fpc/maizeBAC contigs, BAC end sequences (BESs)

Functional Genomics of MuTAIL libraries http://pgir.rutgers.edu/endosperm.info/Maize Endosperm NewEndosperm.html

Gramene Grass comparative genome resources http://www.gramene.org

iMapDB Integrates maize IBM genetic and http://www.maizemap.org/iMapDB/iMap.htmlphysical maps

Maize Assembled Genomic Assembled gene-enriched and http://magi.plantgenomics.iastate.eduIslands (MAGI) whole genome shotgun genome

sample sequences (WGS GSSs)

Maize Fluorescent Protein Investigative resource for cellular http://maize.tigr.org/cellgenomics/Tagged Lines biology in maize; information on

fluorescent reporter lines

Maize Genetic Mapping Project Maize IDP and MMP markers Genetic http://maize-mapping.plantgenomics.iastate.edu

Maize Genetics and Genomics maps, quantitative trait loci (QTL), http://www.maizegdb.orgDatabase (MaizeGDB) molecular markers, EST contigs, gene

products, sequences, and much more

Maize Genetics Cooperation Seed distributor of maize http://maizecoop.cropsci.uiuc.eduStock Center (MGCSC) genetic stocks

Maize Genome Current sequence and annotation http://www.maizesequence.orgSequencing Project of maize genome

Maize Targeted Transposon library for targeted http://mtm.cshl.eduMutagenesis (MTM) selection of gene knockouts

Maize TILLING Project Resource for EMS-mutagenesis for http://genome.purdue.edu/maizetillingreverse genetics

Mu killer Genetic resource for silencing Mu activity http://plantbio.berkeley.edu/~mukiller

Mu Transposon Resource for Mutator-based research http://www.mutransposon.orgInformation Resource

Panzea—Molecular and Bioinformatic and genomics tools; http://www.panzea.orgFunctional Diversity of the resources for maize molecularMaize Genome diversity; maize phenotypic variation

Photosynthetic Mutant Library Library of Mu-induced photosynthetic http://pml.uoregon.edumutants in maize

Plant Genome Database Comparative plant genome http://www.plantgdb.org(PlantGDB) database; species-specific EST and

GSS database, many tools

PlantGDB Ac/Ds resources Resource of Ds lines for forward and http://www.plantgdb.org/prj/reverse genetics; Ds/sequenced BAC AcDsTagging/#Dissociationand annotated gene location files

Regional Mutagenesis Utilizing Current map of Ac elements http://bti.cornell.edu/Brutnell_lab2/Activator (Ac) in Maize Projects/Tagging/BMGG_pro_tagging.html

RescueMu Maize Mutant Phenotype and sequence database http://www.maizegdb.org/Phenotype Database for gene discovery using modified rescuemu-phenotype.php

transposon RescueMu

Sorghum genome sequence at Resources from the Sorghum http://www.phytozome.org/sorghumPhytozome genome project

TIGR Maize Database TIGR resources for maize http://maize.tigr.org/genomics and annotation

TIGR Rice Genome Sequence Rice genomics and annotation http://rice.tigr.org/

UniformMu Maize Project Resource for reverse genetics using http://uniformmu.uf-genome.orgMu-induced mutagenesis in uniforminbred lines

02_EVOv2_033-042_font.qxp:8.5 x 10.875 2/4/10 9:34 AM 5



and is a repository for information regarding all known allelic and cytological variations in maize.The MGCSC collection contains over 100,000 pedigrees, including several hundred gene mutantsand alleles (Neuffer et al. 1997). Also included in the collection are stocks harboring chromosomeaberrations and aneuploidy, inbred-specific ethyl methane sulfonate (EMS) mutant stocks, and aprodigious collection of well-characterized transposable element stocks that are a hallmark of maizegenetic research (see “Genetics and Genomics”).

Owing to its exceptional genetic diversity, maize is highly adaptable and responsive to selectivepressure. As a result, maize has been cultivated from the tropics to southern Canada, a wide bio-geographical range that encompasses tremendous diversity in soil composition, climate, day length,and elevation (Neuffer 1982). The length of the growing period and the quality of ambient light arethe major determinants of the geographical ranges suitable for field cultivation of specific maizeinbred lines.

Although hundreds of maize inbred lines are described, B73, Mo17, and W22 are the most widelyused laboratory accessions. These lines require a growing period of ~100 d; planting to pollinationspans 60-70 d, after which ~40 d are required for seed development and desiccation. In the conti-nental United States, the summer climate allows for field cultivation, whereas winter stocks must beraised in light-supplemented greenhouses. In addition, growing maize at lower latitudes during thewinter months permits the propagation of two large field crops per calendar year. For specific proto-cols and advice on pollinating and growing maize, see “Technical Approaches.”

Most field-grown maize in the continental United States (i.e., under long-day conditions) ispredominately day-neutral with respect to floral induction (Galinat and Naylor 1951). That is, afterthe vegetative meristem initiates a fixed number of vegetative nodes, it converts to floral or tasseldevelopment (Fig. 2). Therefore, for the majority of maize lines, floral induction occurs in responseto a developmental cue rather than day length. However, floral induction in some maize lines canbe influenced by environmental signals (Russell and Stuber 1983). Indeed, photoperiod limits thelatitudinal range of some tropical lines of maize, which are classified as quantitative short-day plants.

RELATED SPECIES

Enabled by the extensive synteny among cereal genomes, comparative studies provide a powerful toolfor gene discovery and analyses of genome evolution in the grasses (Bennetzen and Ma 2003; Devos2005). This extensive colinearity of gene order and orientation between maize, sorghum, rice, wheat,and barley is often exploited to circumvent experimental barriers inherent to one or more of thesespecies. The maize genome is moderately sized (~2.5 gigabase pairs [Gbp]) compared to many ofits grass relatives such as rice (0.4 Gbp), sorghum (0.75 Gbp), barley (6 Gbp), and wheat (17 Gbp)(Mullet et al. 2002; Rabinowicz and Bennetzen 2006). In addition, maize offers decided geneticadvantages in mutant stock collections, cytogenetics, transposon mutagenesis, and ease of controlledpollinations (see “Technical Approaches”). Advantages of rice include a relatively small, sequencedgenome and more efficient transformation technologies. As the first grass genome to be fullysequenced (Goff et al. 2002), rice has been used extensively during annotation of the emerging maizegenomic sequence.

USES OF MAIZE AS A MODEL SYSTEM

Like many other grasses, maize is wind pollinated and a natural cross-pollinator. However, maize is par-ticularly amenable to genetic analysis owing to its monoecious floral development, wherein unisexualmale and female flowers are borne on separate stems. Male (staminate) flowers develop in the tassel,a branched inflorescence that is positioned at the top of the main stem. Female (pistillate) flowers arefound in the ear, a compact inflorescence borne at the ends of reduced lateral branches and locatedat the midpoint of the stem ~five to six nodes below the tassel (Fig. 2). In maize, sexual identity isacquired through the programmed cell death of stamen primordia in female flowers and the corollaryabortion of carpel primordia in male florets (Cheng et al. 1983). The developmental partitioningof male and female flowers has implications for the discovery of genes controlling sex determinationin maize (Delong et al. 1993; Chuck et al. 2007, 2008). Moreover, this physical separation of maleand female flowers greatly facilitates controlled pollinations, in that ear and tassel shoots can beeasily covered to prevent pollen contamination and to collect pollen, respectively. Crossing maize isnot only simple, but also highly productive: Several hundred seed can be produced from a single




FIGURE 2. Zea mays ssp. mays. (A) The adult maize plant is comprised of the above-ground shoot and the undergroundroot. Nodes mark points of leaf attachment along the stem; the internode is the stem segment between successive nodes.The tassel, a branched male inflorescence found at the apex of the stem, contains the staminate flowers. Pistillateflowers are located in the ear, which terminates a short branch found in the axis of leaves near the middle of the stem.The strap-like foliar leaf, a lateral appendage positioned at each node on the stem, has two distinct parts—the distalblade and the proximal sheath that encircles the stem at its insertion. Shoot-borne crown and brace roots are formedfrom consecutive basal nodes. Adventitious roots form the bulk of the root system and give rise to highly branchedsecondary roots. (B) Tassel branches bear an ordered arrangement of small flower-producing branches called spikelets.Two spikelets are initiated at each branch point, and each spikelet produces two functional florets—a pedicellate floretborne on a moderate stem and a sessile floret borne on a short stem. Anthers, forced out of the flower at anthesis,dangle downward and shed pollen. (C) Unlike the unequal spikelets of the tassel, ear spikelets are equivalent in size,and only one floret per spikelet develops to maturity. The mature ear floret contains a single ovary terminated in anelongated style, or silk. During pollination, pollen shed from the anthers germinates on the silk and travels through thegrowing pollen tube to the ovule. The ovary enlarges after fertilization to produce the kernel, a one-seeded fruit. (D)A husk leaf is attached at each node on the stem of the ear shoot. (E) The mature kernel contains the embryo and isenclosed in the pericarp, a transparent tissue layer of maternal origin. Pigmented cells are localized to the aleurone,which form the outermost cell layer of the endosperm. At maturity, the embryo harbors five or six tiny leaf primordiaand a primary root. The endosperm accumulates starch reserves that are mobilized upon germination and transmittedthrough the scutellum to the growing seedling. (Original illustration by Dr. Walton Galinat; used with artist’s permission.)




pollinated ear. This feature sets maize apart from other cereals as an advantageous genetic system—genetic crosses in other cereals require emasculation, a laborious procedure that yields a single seedper flower.

Meiosis in maize is synchronized, such that the relative size of developing maize anthers correlateswith the meiotic stage of the microsporocytes within. This useful feature, coupled with the relativelylarge physical size of maize chromosomes, has placed maize at the forefront of plant cytogeneticresearch. Chromosomal characteristics such as knobs facilitate the tracking of chromosome pairingand segregation. Because of these traits, uncovering meiotic mutants in maize has been relativelystraightforward. In addition, chromosomal aberrations including translocations, inversions, and defi-ciencies were used to assign genes to chromosomal locations (see, e.g., Rhoades 1984). For example,the waxy-marked translocation stocks link this observable endosperm marker to each arm of theten haploid chromosomes in maize, thereby facilitating the placement of genes to chromosomearm (Laughnan and Gabay-Laughnan 1994). Recent, more sophisticated approaches to cytogeneticanalysis, such as the development of single-copy fluorescence in situ hybridization (FISH) andfiber-FISH technologies, have revolutionized studies on maize genomic organization (Jiang et al. 2003;Kato et al. 2004; Wang et al. 2006).

Maize kernel morphology and composition are quantitative traits of immense agronomic andnutritional importance. A single-seeded fruit, the large and prominent maize kernel has been the focusof hundreds of genetic analyses of morphological and biochemical mutants. Barbara McClintock’spioneering research on transposable elements exploited genetically mosaic sectors induced via trans-posable element activity in the aleurone, the anthocyanin-enriched outer cell layer of the maizeendosperm (McClintock 1950; Dooner and Kermicle 1971). Mutations affecting the developmentof the embryo or the accumulation of storage proteins and starch in the endosperm are especiallyabundant, and have contributed to our understanding of developmental and biosynthetic pathwaysoperating in the maize kernel (Laughnan 1953; Scanlon et al. 1994).

As a predominantly out-crossing species, maize has an extraordinary level of genotypic diversity.The frequency of nucleotide polymorphism observed when comparing the genomes of any twomodern maize inbred lines is equivalent to the sequence diversity between chimpanzees and humans(Buckler et al. 2006). Pivotal to the success of maize domestication, this nucleotide diversity continuesto be exploited in modern breeding programs. For example, the native heterozygosity of the maizegenome is exploited in association mapping strategies, wherein correlations between phenotypic andgenetic diversity are identified in analyses of complex, natural populations (Yu and Buckler 2006).Such studies have identified candidate genes associated with complex traits, such as flowering time(Thornsberry et al. 2001), starch biosynthesis (Wilson et al. 2004), and seed carotenoid content(Harjes et al. 2008), which can be manipulated by breeders for agronomic and nutritional improve-ment of maize varieties.

GENETICS AND GENOMICS

A comprehensive, standardized gene nomenclature for maize was established in the late 1990s.Guidelines for the naming of nuclear and organellar genes and gene products, allelic and nonallelic des-ignations, transposon-induced mutations, chromosomal rearrangements, and molecular genetic markerloci are outlined on the Maize Genome and Genetics Database website (http://www.maizegdb.org/maize_nomenclature.php).

Maize research has benefited from a vast collection of genetic mutants (Neuffer et al. 1997). Overthe past decade, high-throughput mutagenesis programs were launched to both expand this inven-tory of maize mutants and expedite the distribution of mutants to the research community (Table 1).For example, the Maize TILLING (Targeting Induced Local Lesions IN Genomes) project is a reverse-genetics approach utilizing gene-targeted screening of EMS-mutagenized inbred populations (Tillet al. 2004; Weil and Monde 2007). A second project involves the use of the Activator (Ac)/Dissociation(Ds) transposable element system in a strategy of targeted, regional mutagenesis. Exploiting thepropensity of Ac and Ds to transpose short distances, this strategy aims to saturate specific chromo-somal regions adjacent to mapped Ac/Ds transposons in an effort to generate new insertional muta-tions in genes closely linked to existing transposons (Bai et al. 2007). Once identified, new Ac/Dsinsertional mutations generate a molecular tag that is utilized for cloning targeted genes (Pohlman etal. 1984). Imprecise transposon excision/insertion events can create “footprint” alleles, which gener-ate allelic variation and contribute to genome evolution (Chen et al. 1987; Kolkman et al. 2005; Baiet al. 2007). Characterized by an increased transposon copy number and a correspondingly elevated




mutation rate, the Mutator (Mu) transposable element system has emerged as a popular methodfor mutagenesis among maize geneticists. The Maize-Targeted Mutagenesis (MTM) project utilizes apolymerase chain reaction (PCR)-based reverse genetic strategy to identify gene-specific, germinaltransposon insertions in a population of ~44,000 plants that are enriched for mobilized Mu elements.Following mutagenesis, Mu activity in the MTM lines is epigenetically silenced by crossing to Mu killer(Slotkin et al. 2003), a strategy that eliminates new Mu transpositions in the progeny (May et al.2003). Importantly, Mu silencing effectively eliminates new somatic Mu insertions, which are nottransmitted to the progeny and can lead to confounding false positives in PCR screens (May et al.2003). A steady-state mutagenesis system is utilized in the UniformMu collection, a forward- andreverse-genetic strategy wherein active Mu transposons are introgressed into the W22 inbred line andinsertion mutations are identified by massively parallel sequence technology (McCarty et al. 2005).Despite these outstanding genetic resources, maize research is slowed by the lack of cost-efficient,high-throughput approaches for genetic transformation, a technology that is still relatively slow andtechnically challenging in maize (Frame et al. 2002).

A diverse toolkit for genomic analyses is readily available to the maize research community.The intermated B73 × Mo17 recombinant inbred lines (IBMRIL; Lee et al. 2002) and their subsequentsaturation with molecular markers, such as restriction fragment length polymorphisms (RFLPs), simplesequence repeats (SSRs; Sharopova et al. 2002), insertion/deletion polymorphisms (IDPs; Fu et al.2006), and single-nucleotide polymorphisms (SNPs), have greatly increased the resolution of themaize genetic map. Capitalizing on the extensive heterogeneity within the maize gene pool, themaize nested associated mapping (NAM) population comprises 5000 RILs developed from crossesbetween B73 and each of 25 diverse maize inbred lines (Yu and Buckler 2006). The NAM populationcaptures much of the natural allelic variation present in Zea mays ssp. mays and represents a power-ful tool for the fine-scale resolution and molecular dissection of genes contributing to complex traitsin maize. Positional cloning is emerging as the most powerful tool for gene identification in maize, aprocedure that is greatly enhanced by the genomic sequencing effort (Bortiri et al. 2006). Links tothese tools and other related maize resources are summarized in Table 1.

Genomic sequencing of the maize inbred line B73 is nearing completion; full release of thegenome sequence is expected during 2009. B73 was selected as the reference genome because itperforms well throughout much of the continental United States, exhibits high yield, and is the mostutilized inbred line in basic research laboratories. Consequently, B73 is the source of the majority ofthe maize expressed sequence tags (ESTs) and commercially important germplasm, as well as the vastcollection of bacterial artificial chromosome (BAC) libraries (Bennetzen et al. 2001). Presently, anno-tation of the maize genome is incomplete. Annotation efforts are driven by the maize researchcommunity, and maize gene predictions are extensively derived from models of rice orthologs.

TECHNICAL APPROACHES

Maize genetic research is greatly facilitated by the physical separation of male and female flowers,making controlled pollinations simple and highly productive. A detailed protocol for maize pollination,complete with video demonstrations, helpful tips, and required materials can be accessed from thewebsite “Controlled Pollinations of Maize” (http://www.maizegdb.org/IMP/WEB/pollen.htm). Anefficient protocol for the isolation of maize DNA suitable for use in restriction digests, small-insert (i.e.,25 kb or less) cloning, and PCR analysis has been developed by Dr. Stephen Dellaporta and colleagues(http://www.agron.missouri.edu/mnl/57/25dellaporta.html). Maize has been extensively utilized inthe development of advanced cytogenetic protocols, including genomic in situ hybridization (GISH),single copy FISH (Wang et al. 2006), chromosome painting (Kato et al. 2004), and extended fiber FISH(Jiang et al. 2003). Protocols for GISH and fiber FISH can be accessed from Dr. Jiming Jiang’s labora-tory (http://www.hort.wisc.edu/jjiang/protocols_1.htm). Maize is also the first organism utilized for insitu hybridization of small RNA transcripts (21-24 nucleotides) utilizing locked nucleic acid (LNA)hybridization probes, as described by Drs. Catherine Kidner and Marja Timmermans (http://schnablelab.plantgenomics.iastate.edu/docs/resources/protocols/pdf/in_situ_protocol.2007.04.01.pdf); Kidner and Timmermans 2006).

Maize is a highly tractable system for developmental genetic and genomics. The relatively largesize of maize organs, such as the leaf primordia and shoot apical meristem (SAM), renders them espe-cially suitable for laser microdissection analyses. Laser microdissection permits the precise isolation ofdiscrete cells, tissues, or organs from thin sections of fixed plant tissue immobilized on glass slides.




RNA isolated from microdissected samples is suitable for use in expression analyses, such as quantita-tive real-time PCR (qRT-PCR), microarray hybridization, and massive parallel RNA sequencing(RNA-Seq). Provided here are links to detailed protocols from Dr. Patrick Schnable’s laboratory:

• Growing maize seedlings in controlled environmental conditions suitable for genomic analyses(http://schnablelab.plantgenomics.iastate.edu/docs/resources/protocols/pdf/Growing_maize_seedlings.2007.04.01.pdf);

• Tissue fixation, paraffin embedding, and microtome sectioning of maize shoots (http://schnablelab.plantgenomics.iastate.edu/docs/resources/protocols/pdf/Paraffin_sectioning_for_LCM.2007.10.19.pdf);

• Cryosectioning of maize tissues (http://schnablelab.plantgenomics.iastate.edu/docs/resources/protocols/pdf/Cryosectioning_for_LCM.2007.04.01.pdf);

• Linear amplification of microdissected RNA (http://schnablelab.plantgenomics.iastate.edu/docs/resources/protocols/pdf/RNA_amplification.2007.04.01.pdf);

• qRT-PCR analyses of maize cDNA (http://schnablelab.plantgenomics.iastate.edu/docs/resources/protocols/pdf/realtime_RT-PCR.2007.04.01.pdf);

• Maize Tissue Preparation and Extraction of RNA from Target Cells for Genotyping (Ohtsuand Schnable 2007a);

• T7-Based RNA Amplification for Genotyping from Maize Shoot Apical Meristem (Ohtsu andSchnable 2007b); and

• SNP Mining from Maize 454 EST Sequences (Ohtsu and Schnable 2007c).

ACKNOWLEDGMENTS

We acknowledge the vast contributions and friendly collegiality of the past and present community ofmaize researchers.

REFERENCES

Bai L, Singh M, Pitt L, Sweeney M, Brutnell TP. 2007. Generatingnovel allelic variation through Activator insertional mutagenesis inmaize. Genetics 175: 981–992.

Bennetzen JL. 2000. Comparative sequence analysis of plant nucleargenomes: Microcolinearity and its many exceptions. Plant Cell 12:1021–1029.

Bennetzen JL, Ma J. 2003. The genetic colinearity of rice and othercereals on the basis of genomic sequence analysis. Curr Opin PlantBiol 6: 128–133.

Bennetzen JL, Chandler VL, Schnable PS. 2001. National ScienceFoundation-sponsored workshop report. Maize genome sequenc-ing project. Plant Physiol 127: 1572–1578.

Bolot S, Abrouk M, Masood-Quraishi U, Stein N, Messing J, Feuillet C,Salse J. 2009. The ‘inner circle’ of the cereal genomes. Curr OpinPlant Biol 12: 119–125.

Bortiri E, Jackson D, Hake S. 2006. Advances in maize genomics: Theemergence of positional cloning. Curr Opin Plant Biol 9: 164–171.

Buckler ES, Gaut BS, McMullen MD. 2006. Molecular and functionaldiversity of maize. Curr Opin Plant Biol 9: 172–176.

Chen J, Greenblatt LM, Dellaporta SL. 1987. Transposition of Ac fromthe P locus of maize into unreplicated chromosomal sites.Genetics 117: 109–116.

Cheng PC, Greyson RI, Waldon DB. 1983. Organ initiation and thedevelopment of unisexual flowers in the tassel and ear of Zeamays. Am J Bot 70: 450–462.

Chuck G, Meeley R, Irish E, Hajime S, Hake S. 2007. The maizetasselseed4 microRNA controls sex determination and meristemcell fate by targeting Tasselseed6/indeterminate spikelet1. NatGenet 39: 1517–1521.

Chuck G, Meeley R, Hake S. 2008. Floral meristem initiation andmeristem cell fate are regulated by the maize AP2 genes ids1 andsid1. Development 135: 3013–3019.

Coe EH Jr. 2001. The origins of maize genetics. Nat Rev Genet 2:898–905.

DeLong A, Calderon-Urrea A, Dellaporta SL. 1993. Sex determinationgene TASSELSEED2 of maize encodes a short-chain alcoholdehydrogenase required for stage-specific floral organ abortion.Cell 74: 757–768.

Devos KM. 2005. Updating the ‘crop circle.’ Curr Opin Plant Biol8: 155–162.

Doebley J. 2004. The genetics of maize evolution. Annu Rev Genet 38:37–59.

Dooner HK, Kermicle JL. 1971. Structure of the Rr tandem duplicationin maize. Genetics 67: 427–436.

Frame BR, Shou H, Chikwamba RK, Zhang Z, Xiang C, Fonger TM,Pegg SEK, Li B, Nettleton DS, Pei D, et al. 2002. Agrobacteriumtumefaciens-mediated transformation of maize embryos using astandard binary vector system. Plant Physiol 129: 13–22.

Fu Y, Wen TJ, Ronin YI, Chen HD, Gou L, Mester DI, Yang Y, Lee M,Korol AB, Ashlock DA, et al. 2006. Genetic dissection of inter-mated recombinant inbred lines using a new genetic map ofmaize. Genetics 174: 1671–1683.

Fukunaga K, Hill J, Vigouroux Y, Matsuoka Y, Sanchez J, Liu K, BucklerES, Doebley J. 2005. Genetic diversity and population structure ofteosinte. Genetics 169: 2241–2254.

Gale MD, Devos KM. 1998. Comparative genetics in the grasses.Proc Natl Acad Sci 95: 1971–1974.

Galinat WC, Naylor AW. 1951. Relation of photoperiod to inflores-cence proliferation in Zea mays L. Am J Bot 38: 38–47.

Gaut BS, Doebley JF. 1997. DNA sequence evidence for the segmentalallotetraploid origin of maize. Proc Natl Acad Sci 88: 2060–2064.

Goff SA, Ricke D, Lan TH, Presting G, Wang R, Dunn M, Glazebrook J,Sessions A, Oeller P, Varma H, et al. 2002. A draft sequence of therice genome (Oryza sativa L. ssp. japonica). Science 296: 92–100.




Harjes CE, Rocheford TR, Bai L, Brutnell TP, Kandianis CB, SowinskiSG, Stapleton AE, Vallabhaneni R, Williams M, Wurtzel ET, et al.2008. Natural genetic variation in lycopene epsilon cyclasetapped for maize biofortification. Science 319: 330–333.

Jiang J, Birchler JA, Parrott WA, Dawe RK. 2003. A molecular view ofplant chromosomes. Trends Plant Sci 8: 570–575.

Kato A, Lamb JC, Birchler JA. 2004. Chromosome painting usingrepetitive DNA sequences as probes for somatic chromosomeidentification in maize. Proc Natl Acad Sci 101: 13554–13559.

Kellogg EA. 2001. Evolutionary history of the grasses. Plant Physiol125: 1198–1205.

Kidner C, Timmermans M. 2006. In situ hybridization as a tool tostudy the role of microRNAs in plant development. Methods MolBiol 342: 159–179.

Kolkman JM, Conrad LJ, Farmer PR, Hardeman K, Ahern KR, Lewis PE,Sawers RJH, Lebejko S, Chomet P, Brutnell TP. 2005. Distributionof Activator (Ac) throughout the maize genome for use in regionalmutagenesis. Genetics 169: 981–995.

Laughnan JR. 1953. The effect of the sh2

factor on carbohydratereserves in the mature endosperm of maize. Genetics 38: 485–499.

Laughnan JR, Gabay-Laughnan S. 1994. The placement of genesusing waxy-marked reciprocal translocations. In The maize hand-book (eds. M Freeling and V Walbot), pp. 255–257. Springer,New York.

Lee M, Sharopova N, Beavis WD, Grant D, Katt M, Blair D, HallauerA. 2002. Expanding the genetic map of maize with the inter-mated B73 × Mo17 (IBM) population. Plant Mol Biol 48: 453–461.

Matsuoka Y, Vigouroux Y, Goodman MM, Sanchez GJ, Buckler E,Doebley J. 2002. A single domestication for maize shown bymicrosatellite genotyping. Proc Natl Acad Sci 99: 6080–6084.

May BP, Liu H, Volbrecht E, Senior L, Rabinowicz PD, Roh D, Pan X,Stein L, Freeling M, Alexander D, et al. 2003. Maize-targetedmutagenesis: A knockout resource for maize. Proc Natl Acad Sci100: 11541–11546.

McCarty DR, Settles AM, Suzuki M, Tan BC, Latshaw S, Porch T, RobinK, Baier J, Avigne W, Lai J, et al. 2005. Steady-state transposonmutagenesis in inbred maize. Plant J 44: 52–61.

McClintock B. 1950. The origin and behavior of mutable loci inmaize. Proc Natl Acad Sci 36: 344–355.

Mullet JE, Klein RR, Klein PE. 2002. Sorghum bicolor—An importantspecies for comparative grass genomics and a source of beneficialgenes for agriculture. Curr Opin Plant Biol 5: 118–121.

Neuffer MG. 1982. Growing maize for genetic purposes. In Maize forbiological research (ed. WF Sheridan), pp. 19–30. University Press,University of North Dakota, Grand Forks, ND.

Neuffer MG, Coe EH, Wessler SR. 1997. Mutants of maize. ColdSpring Harbor Laboratory Press, Cold Spring Harbor, NY.

Ohtsu K, Schnable PS. 2007a. Maize tissue preparation and extrac-tion of RNA from target cells for genotyping. Cold Spring HarbProtoc doi: 10.1101/pdb.prot4784.

Ohtsu K, Schnable PS. 2007b. T7-based RNA amplification for geno-typing from maize shoot apical meristem. Cold Spring Harb Protocdoi: 10.1101/pdb.prot4785.

Ohtsu K, Schnable PS. 2007c. SNP mining from maize 454 ESTsequences. Cold Spring Harb Protoc doi: 10.1101/pdb.prot4784.

Piperno DR, Flannery KV. 2001. The earliest archaeological maize(Zea mays L.) from highland Mexico: New accelerator massspectrometry dates and their implications. Proc Natl Acad Sci 98:2101–2103.

Pohlman RF, Fedoroff NP, Messing J. 1984. The nucleotide sequenceof the maize controlling element Activator. Cell 37: 635–643.

Rabinowicz PD, Bennetzen JL. 2006. The maize genome as a modelfor efficient sequence analysis of large plant genomes. Curr OpinPlant Biol 9: 149–156.

Rhoades MM. 1984. The early years of maize genetics. Annu RevGenet 18: 1–29.

Russell WK, Stuber CW. 1983. Effects of photoperiod and tempera-tures on the duration of vegetative growth in maize. Crop Sci 23:847–850.

SanMiguel P, Tikhonov A, Jin YK, Motchoulskaia N, Zakharov D,Melake-Berhan A, Springer PS, Edwards KJ, Lee M, Avramova Z,et al. 1996. Nested retrotransposons in the intergenic regions ofthe maize genome. Science 274: 765–768.

SanMiguel P, Gaut BS, Tikhonov A, Nakajima Y, Bennetzen JL. 1998.The paleontology of intergene retrotransposons of maize. NatGenet 20: 43–45.

Scanlon MJ, Stinard PS, James MG, Myers AM, Roberston DS. 1994.Genetic analysis of 63 mutations affecting maize kernel develop-ment isolated from Mutator stocks. Genetics 136: 281–294.

Sharopova N, McMullen MD, Schultz L, Schroeder S, Sanchez-VilledaH, Gardiner J, Bergstrom D, Houchins K, Melia-Hancock S, MusketT, et al. 2002. Development and mapping of SSR markers formaize. Plant Mol Biol 48: 463–481.

Slotkin RK, Freeling M, Lisch D. 2003. Mu killer causes the heritableinactivation of the Mutator family of transposable elements in Zeamays. Genetics 165: 781–797.

Swigonova Z, Lai J, Ma J, Ramakrishna W, Llaca V, Bennetzen JL,Messing J. 2004. Close split of sorghum and maize genomeprogenitors. Genome Res 14: 1916–1923.

Thornsberry JM, Goodman MM, Doebley J, Kresovich S, Nielsen D,Buckler ES. 2001. Dwarf8 polymorphisms associate with variationin flowering time. Nat Genet 28: 286–289.

Till BJ, Reynolds SH, Weil C, Springer N, Burtner C, Young K, BowersE, Codomo CA, Enns LC, Odden AR, et al. 2004. Discovery ofinduced point mutations in maize by TILLING. BMC Plant Biol 4:12. doi: 10.1186/1471-2229-4-12.

Vollbrecht E, Sigmon B. 2005. Amazing grass: Developmental genet-ics of maize domestication. Biochem Soc Trans 33: 1502–1506.

Wang CJ, Harper L, Cande WZ. 2006. High-resolution single-copy gene fluorescence in situ hybridization and its use in theconstruction of a cytogenetic map of maize chromosome 9. PlantCell 18: 529–544.

Weil CF, Monde RA. 2007. Getting to the point—mutations in maize.Crop Sci 47: S60–S67.

Wilson LM, Whitt SR, Ibanez AM, Rocheford TM, Goodman MM,Buckler ES. 2004. Dissection of maize kernel composition andstarch production by candidate gene association. Plant Cell 16:2719–2733.

Yu J, Buckler ES. 2006. Genetic association mapping and genomeorganization of maize. Curr Opin Biotech 17: 155–160.



Emerging Model Organisms, Volume 2 365 Ciona intestinalis

1Present address: Center for Developmental Genetics, Department ofBiology, Faculty of Arts and Science, New York University, New York,NY 10003, USA.2Corresponding authors ([email protected]); ([email protected]).Cite as: Cold Spring Harb Protoc; 2009; doi:10.1101/pdb.emo138 www.cshprotocols.org

Chapter 13

The Sea Squirt Ciona intestinalis

Lionel Christiaen,1,2 Eileen Wagner, Weiyang Shi, and Michael Levine2

Molecular and Cell Biology Department, University of California, Berkeley, California 94720, USA

INTRODUCTION

Sea squirts (Ciona intestinalis) are tunicates (or urochordates), the closest living relatives of the verte-brates. Although the adults are simple, sessile filter feeders, the embryos and larvae possess clear chor-date features including a prominent notochord and dorsal, hollow neural tube. Tail-bud-stageembryos and mature swimming tadpoles are composed of approximately 1000 and 2600 cells,respectively, with complete lineage information. This cellular simplicity is coupled with a streamlinedgenome that has not undergone the duplications seen in vertebrates. A variety of molecular tools havebeen applied to understanding Ciona embryogenesis. Comparisons of the C. intestinalis genome andthe related but divergent Ciona savignyi genome have facilitated the identification of conserved non-coding DNAs, including regulatory DNAs such as tissue-specific enhancers. Systematic in situhybridization assays and gene-disruption experiments using specific morpholino antisense oligonu-cleotides have led to the elaboration of provisional gene regulatory networks underlying the specifi-cation of key chordate tissues, including the notochord, neural tube, and beating heart. Thesenetworks provide a foundation for understanding the mechanistic basis of more complex cell-specifi-cation processes in vertebrates, and for understanding the evolutionary origins of distinctive verte-brate characteristics such as the neural crest. Because tunicates and vertebrates are sister groups, thereis every indication that the developmental mechanisms revealed in the simple Ciona model will beapplicable to comparable processes in vertebrates.

RELATED INFORMATION

The Tunicate Web Portal (http://www.tunicate-portal.org) is the entry point of choice for all webresources concerning research on tunicates.

BACKGROUND INFORMATION

The tunicate (or urochordate) C. intestinalis (Linnaeus, 1767) is a solitary ascidian species that lives inshallow waters in the inter- to subtidal zones of most temperate coasts around the world. These ses-sile animals are usually found in groups attached to a variety of natural and artificial immersed mate-rials. They spend most of their life cycle as adult filter feeders that produce both oocytes and sperm,which are released on a daily basis (usually at dawn, upon sunrise).

The hermaphroditic adult looks like an elongated gelatinous bag surrounded by a translucenttunic containing cellulose, and it displays two conspicuous oral and atrial siphons delineated by acorona of yellow pigmentation (Fig. 1A). This typical invertebrate morphology led the French zoolo-gist Georges Cuvier (1769-1832) to classify ascidians as molluscs. However, gill slits are observed inthe adult pharynx, and larvae display key characteristics of a chordate tadpole: an elongated tail sup-ported by a central rod of vacuolized notochord cells and a dorsal hollow neural tube markedly pat-terned along the antero-posterior axis (Fig. 1B). These characteristics of the ascidian tadpole led theRussian embryologist Alexander Kovalevsky (1840-1901) to classify tunicates as bona fide chordates,




a phylogenetic position that has since been extensively confirmed by molecular phylogenetic analy-ses (Fig. 1C; e.g., Delsuc et al. 2006).

The subphylum Tunicata (or Urochordata), to which C. intestinalis belongs, includes numerous(more than 3000) extant species that have traditionally been classified into four major classes:Ascidiacea, Thaliacea, Appendicularia (or Larvacea), and Sorberacea. These classes share the uniqueability to synthesize cellulose as a result of an ancient horizontal transfer event involving the cellulosesynthase gene (Matthysse et al. 2004). The class Ascidiacea is traditionally subdivided into two orders,Stolidobranchia and Enterogona; the latter comprises the Aplousobranchia and Phlebobranchiasuborders. These clades constitute monophyletic groups, and the Thaliacea and Appendicularia classesare sister groups to the Phlebobranchia and Aplousobranchia, respectively (Fig. 1C). In this complexphylogeny, C. intestinalis is a founding member of the Cionidae family of Phlebobranchia ascidians.

SOURCES AND HUSBANDRY

Developmental studies using C. intestinalis rely on the availability of gravid adult animals. In the bestseason, usually from April to December in the northern hemisphere, thousands of oocytes and up to200 µL of concentrated sperm can be obtained by dissecting a single animal. Currently, the vastmajority of studies use animals collected from their natural habitats (e.g., coastal harbors in the Pacificand Atlantic Oceans). These animals can be kept in seawater tanks equipped with appropriate pumps,filters, and skimmers. Animals are usually maintained in short-term culture for 1-2 wk and do not needto be fed during this time.

Several attempts to grow C. intestinalis in the lab have been reported (Hendrickson et al. 2004;Joly et al. 2007). Such efforts circumvent the constraints on collecting fresh animals from the wild;furthermore, these animals have natural fluctuations in fecundity. Unfortunately, the most efficientcultivation depends on the availability of natural running seawater, which can only be achievedwith coastal infrastructures. Nonetheless, a standardized culturing system would permit the establish-ment and distribution of isogenic strains. Indeed, cryptic subspecies appear to coexist in the wild, andthe high natural polymorphism (~1%-2%) may interfere with systematic genetic studies (Sordinoet al. 2008).

To get started, we suggest using fresh animals and short-term maintenance in a simple saltwatertank in the lab. The following procedures apply to both C. intestinalis and C. savignyi, although C.intestinalis has a broader worldwide distribution.

FIGURE 1. C. intestinalis body plan and phylogeny. (A) Adult body plan; (o.s.) oral siphon; (a.s.) atrial siphon; (p.s.) pig-mented spot; (s.d.) sperm duct; (o.d.) ooduct. (B) Larval body plan (anterior is up, dorsal side is left). Note the pigmentedcells in the sensory vesicle (anterior ventral otolith and posterior dorsal ocellus). Different scales are used on the adultand larval pictures. (C) A phylogeny of tunicates. The tree shows the phylogenetic relationships between selected groupsof deuterostomes. Only the topology is relevant; the branch lengths do not represent evolutionary distances. Althoughtunicates presumably have an ancestral benthic lifestyle, members of the Thaliacea and Appendicularia, denoted withan asterisk (*), are pelagic animals. (For color figure, see doi: 10.1101/pdb.emo138 online at www.cshprotocols.org.)




Ciona adults are usually found growing on wooden docks and floats in marine harbors. Animalstightly adhere to their substrates, so take care when detaching them by using the base of the animalto gently pull. After detachment, place the animals in a cooler containing a large volume of freshseawater. If the research lab is not located near an ocean, it is possible to purchase animals from avariety of marine supply companies. Marine Research and Educational Products (M-Rep;http://www.m-rep.com/profile.htm), located in San Diego, CA, can ship animals to either the west orthe east coast of the United States. The Roscoff Biological Station (http://www.sb-roscoff.fr/ModBiol),affiliated with the University of Paris VI, supplies animals to laboratories located in Europe. There is aplentiful supply of gravid animals during the warmer months, from May through October, on bothcoasts of the United States as well as in many parts of Europe and Asia. From January to April, it is bestto purchase animals from warmer climates, such as San Diego, CA.

Animals can be shipped by FedEx, but it is important to minimize the transport time and keepthe temperature low (no more than ~10°C) with ample aeration. Ship animals in 3-4 L of seawater;this volume is sufficient to sustain 20-30 animals overnight. Use double plastic bags surrounded byice packs to keep the temperature down. Upon receipt of animals after shipment, slowly raise thetemperature by leaving the plastic bag in a seawater tank. Cut the bag open while incubating in orderto provide adequate levels of oxygen.

An aquarium with an open circulating seawater tank system that can hold ~80-100 gal of seawa-ter should be adequate for most lab uses. Basically, the system should contain a temperature-controlled cooling unit, circulator pump, denitrification unit, protein skimmer, and filter for solidwastes. Keep the water temperature at ~16°C-18°C. Use hanging baskets to hold the animals. It isimportant to provide constant illumination over the tank in order to prevent animals from spawningwith the normal day-night cycle. Use fluorescent lamps that emit a strong daylight spectrum.

RELATED SPECIES

The most relevant related species for studying C. intestinalis is C. savignyi. Its genome was sequencedshortly after that of C. intestinalis (Small et al. 2007) and was used to compute whole-genomealignments, providing streamlined phylogenetic footprinting results that have proven quite valuablefor the computational identification of non-coding DNAs, such as tissue-specific enhancers (Johnsonet al. 2005). In addition, spontaneous and induced mutations have been isolated and analyzed inC. savignyi, thereby providing a foundation for more comprehensive genetic studies (e.g., Nakataniet al. 1999).

The Japanese ascidian Halocynthia roretzi produces significantly larger eggs than C. intestinalis,permitting the elucidation of complete fate maps of early blastomeres until the onset of gastrulation(110-cell stage) (Nishida 1987). In addition, numerous studies using embryological manipulations andmicroinjection in this species have uncovered essential aspects of early ascidian development, such asthe identification of the classical muscle determinant, macho-1 (Nishida and Sawada 2001).

The reduced cell number constituting ascidian embryos offers an opportunity for detailed molec-ular imaging in live embryos. In this regard, the natural pigmentation of C. intestinalis embryos can beproblematic, a limitation that has been circumvented using Ascidiella aspersa and Phallusia mammillataembryos, which are absolutely transparent.

Other tunicate species are being used to provide insights into diverse biological phenomena. Thecolonial ascidian Botryllus schlosseri, which can be raised in the laboratory and is amenable to classicalgenetics, provides an increasingly attractive model for studying asexual reproduction, germ cellbiology, allorecognition, and the evolution of the immune system (e.g., De Tomaso et al. 2005). Theappendicularian Oïkopleura dioïca has an amazingly short life cycle (4 d) and constitutes an extremeexample of genomic compaction and rearrangements (the complete genome is just ~80 Mb), butnonetheless displays the basic chordate body plan (Seo et al. 2001).

USES OF THE CIONA MODEL SYSTEM: EMBRYONIC DEVELOPMENT

The use of ascidian embryos for experimental studies was initiated in the 19th century by the Frenchbiologist Laurent Chabry (1855-1894), who performed the first manipulations of living embryos usingA. aspersa. Subsequently, the American embryologist Edwin Conklin reported a comprehensivedescription of early cell lineages in the species Styela clava and defined the nomenclature for ascidianblastomeres that is still in use.

13_EVOv2_365-396.qxp:8.5 x 10.875 2/19/10 2:54 PM Page 367



Ascidian embryos became a paradigm of so-called mosaic development, whereby individualblastomeres are endowed with maternal determinants specifying diverse cell fates such as muscle,endoderm, or epidermis. Since the early 1980s, the increasing use of molecular tools helped deter-mine the nature and function of such determinants and refine the traditional view by showing thatcell-to-cell interactions are also essential for the specification of the basic chordate tissues, such as thenotochord and the central nervous system (CNS). The classical era of ascidian “mosaic” developmentreached closure when Nishida and colleagues reported the identification of the zinc finger transcrip-tion factor macho-1 as the classical maternal muscle determinant originally described by Chabry andConklin (Nishida and Sawada 2001).

Numerous ascidian species are used in developmental studies; however, C. intestinalis reigns as thedominant model because of its worldwide distribution, the availability of a complete genome assem-bly (Dehal et al. 2002), the establishment of post-genome tools such as the Agilent and Affymetrixmicroarrays (Azumi et al. 2003; Christiaen et al. 2008), and the determination of comprehensive geneexpression profiles and gene disruption assays (Imai et al. 2004, 2006) in this species. The number ofdevelopmental studies using C. intestinalis has increased exponentially in the last 20 yr, and they havebeen the subject of numerous excellent review articles (Nishida 2002; Satoh 2003; Satoh et al. 2003;Passamaneck and Di Gregorio 2005). Here, we briefly summarize a few key landmark studies, alongwith some of the emerging general findings relevant to chordate development.

Gene Regulatory Networks and Cleavage Patterns in the Early Embryo

The Ciona system is well suited for the rapid identification of cis-regulatory DNAs, particularlytissue-specific enhancers. Initial efforts along these lines depended on traditional microinjection assays.However, the advent of a simple electroporation method (see Electroporation of Transgenic DNAsin the Sea Squirt Ciona [Christiaen et al. 2009a]) permitted the simultaneous transformation ofhundreds or even thousands of synchronously developing embryos. This method was initially used forthe identification of a notochord-specific enhancer from the 5′ regulatory region of the CionaBrachyury gene (Corbo et al. 1997). The green fluorescent protein (GFP) reporter gene was used forthis analysis; this reporter works exceptionally well for tracing individual cells, tissues, and lineagesduring Ciona embryogenesis. The subsequent characterization of the Brachyury enhancer led to theelucidation of a provisional gene network for notochord specification, including distinctive regulatoryinputs for the primary and secondary notochord lineages. Similar methods were used to dissectthe Otx enhancer, which is an early marker for primary neural induction in the Ciona embryo. Thecharacterization of this enhancer led to the identification of fibroblast growth factor (FGF) signaling(FGF9/16/20), Ets/pointed2, and GATAa as the key mediators of Otx induction in the presumptiveCNS (Bertrand et al. 2003).

Morphogenesis and Differentiation in a Chordate Body Plan

A provisional “blueprint” for the early Ciona embryo was determined by systematically analyzing 76different regulatory genes that encode either sequence-specific transcription factors (TFs) or cell-signaling components (STs) that impinge on the activities of these factors (Imai et al. 2006). Each ofthese 76 genes (53 TFs and 23 STs) is expressed in a specific subset of blastomeres between thecritical 16- and 110-cell stages of embryogenesis. About 30 of these genes were systematicallydisrupted by microinjecting antisense morpholino oligonucleotides (see Microinjection ofMorpholino Oligos and RNAs in Sea Squirt (Ciona) Embryos [Christiaen et al. 2009b]), and theexpression profiles of the remaining genes were examined by quantitative reverse transcriptasepolymerase chain reaction (RT-PCR) assays and/or by in situ hybridization (see Whole-Mount In SituHybridization on Sea Squirt (Ciona intestinalis) Embryos [Christiaen et al. 2009c]). These studiesprovided a foundation for understanding the specification of a variety of different cell types, such asthe progenitors of the cardiomyocytes comprising the beating heart of adult sea squirts.

The adult heart arises from a single pair of blastomeres, the B7.5 blastomeres, at the 110-cell stageof embryogenesis. Mesp, which encodes a bHLH transcription factor, is a key determinant of thecardiac mesoderm (Satou et al. 2004). Mesp is required for the induction of the heart fate by FGFsignaling in the anterior descendants of the B7.5 lineage (Davidson et al. 2006). A variety of targetgenes required for heart cell differentiation and directed migration are activated downstream fromMesp and FGF signaling in the Ciona tadpole. One such target gene is the forkhead transcriptionfactor FoxF, which mediates directed precardial migration by regulating a variety of cellular effectors,




such as RhoDF (Beh et al. 2007; Christiaen et al. 2008). This example illustrates one of the greatstrengths of the Ciona system, namely, the ability to link regulatory networks to cellular morphogen-esis in the developing embryo.

GENETICS

Developmental studies involving the manipulation of wild-type embryos potentially outpace geneticstudies. C. intestinalis has a generation time of several months, even though its eggs develop intoswimming larvae within 24 h. This relatively long generation time, coupled with the culturing restric-tions described above, has slowed the development of classical genetic tools for studying C. intesti-nalis. However, there has been notable progress in the creation of genetic maps coupled to bacterialartificial chromosome (BAC) clones for each of the 14 C. intestinalis chromosomes (Shoguchi et al.2006; Kano 2007).

Forward genetic approaches have been performed using the sibling species C. savignyi, and therecent development of stable transgene insertion methods for both species has led to the generationof enhancer-trap and mutant lines (Deschet et al. 2003; Matsuoka et al. 2004, 2005; Sasakura et al.2008). C. savignyi has a shorter generation time than C. intestinalis (6 wk rather than a few months),and it has been possible to exploit the self-fertilization of hermaphrodites to perform genetic screensfor patterning mutants. This approach has been used with considerable success for the identificationof a variety of genes required for the morphogenesis of the Ciona notochord. As discussed earlier, itis entirely feasible to use both C. savignyi and C. intestinalis for the concerted analysis of specificdevelopmental processes. For example, C. intestinalis Brachyury-GFP fusion genes work just fine inC. savignyi, permitting the detailed cellular visualization of notochord patterning mutants.

GENOMIC AND POST-GENOME TOOLS

The C. intestinalis genome is ~160 Mb in size and contains about 16,000 protein-coding genes, whichis a similar gene density to that seen in Drosophila. The C. intestinalis genome was the seventh animalgenome to be completely assembled and annotated (Dehal et al. 2002). This assembly, along withthe C. savignyi genome assembly, permits investigators to use PCR-based methods for the isolationof specific genomic DNAs, either coding or non-coding. The JGI (http://genome.jgi-psf.org/ciona4/ciona4.home.html) was the first repository for C. intestinalis genomic sequences; version 1is the favorite of the research community as agreed on at the International Tunicate meeting held inVillefranche-sur-Mer (France) in 2007. The ENSEMBL database hosts version 2 of the C. intestinalisgenome, allowing integrated interfacing with genomes from other species for comparative genomics(http://www.ensembl.org/Ciona_intestinalis/Info/Index). A mirror of the UCSC browser for C. intesti-nalis (http://genome.ciona.cnrs-gif.fr/cgi-bin/hgGateway?org=C.+intestinalis&db=ci1) is hosted byGif-sur-Yvette (France) and displays additional tracks (e.g., Affymetrix probe sets).

There is extensive annotation of the C. intestinalis genome, mainly owing to the comprehensiveexpressed sequence tag (EST) collections, including full-length cDNAs, that are publicly available(Satou et al. 2002, 2008). An additional cDNA library has been introduced into the versatile Gatewayvectors by Patrick Lemaire and coworkers at the Institute for Developmental Biology of Marseilles(France) (Roure et al. 2007). BAC clones are available upon request from the Satoh lab (KyotoUniversity), and an arrayed cosmid library is available (Burgtorf et al. 1998), although it has not beenmapped to the genome. Sequence identification is achieved through hybridization on membranereplicates of the library. Finally, both Agilent and Affymetrix microarrays are available for transcriptomeprofiling (Azumi et al. 2003; Christiaen et al. 2008).

The rapidly expanding molecular information concerning the organization and function of theC. intestinalis genome is available on several publicly available websites. The Ghost Database main-tained by Yutaka Satou and colleagues (http://hoya.zool.kyoto-u.ac.jp/cgi-bin/gbrowse/ci) hosts mostof the expression data and annotations currently available for C. intestinalis. The ANISEED database(http://crfb.univ-mrs.fr/aniseed) hosts most of the genomic and expression information knownabout C. intestinalis in a very integrated manner. The virtual early embryo can also be downloadedfrom this database. The Four Dimensional Body Atlas (FABA) database for C. intestinalis (http://chordate.bpni.bio.keio.ac.jp/faba/1.2/top.html) constitutes the most comprehensive resource forC. intestinalis morphological information through the juvenile stage.




Azumi K, Takahashi H, Miki Y, Fujie M, Usami T, Ishikawa H, KitayamaA, Satou Y, Ueno N, Satoh N. 2003. Construction of a cDNAmicroarray derived from the ascidian Ciona intestinalis. Zoolog Sci20: 1223–1229.

Beh J, Shi W, Levine M, Davidson B, Christiaen L. 2007. FoxF is essen-tial for FGF-induced migration of heart progenitor cells in theascidian Ciona intestinalis. Development 134: 3297–3305.

Bertrand V, Hudson C, Caillol D, Popovici C, Lemaire P. 2003. Neuraltissue in ascidian embryos is induced by FGF9/16/20, acting via acombination of maternal GATA and Ets transcription factors. Cell115: 615–627.

Burgtorf C, Welzel K, Hasenbank R, Zehetner G, Weis S, Lehrach H.1998. Gridded genomic libraries of different chordate species:A reference library system for basic and comparative genetic stud-ies of chordate genomes. Genomics 52: 230–232.

Christiaen L, Davidson B, Kawashima T, Powell W, Nolla H, VranizanK, Levine M. 2008. The transcription/migration interface in heartprecursors of Ciona intestinalis. Science 320: 1349–1352.

Christiaen L, Wagner E, Shi W, Levine M. 2009a. Electroporation oftransgenic DNAs in the sea squirt Ciona. Cold Spring Harb Protoc(this issue). doi: 10.1101/pdb.prot5345.

Christiaen L, Wagner E, Shi W, Levine M. 2009b. Microinjection ofmorpholino oligos and RNAs in sea squirt (Ciona) embryos. ColdSpring Harb Protoc (this issue). doi: 10.1101/pdb.prot5347.

Christiaen L, Wagner E, Shi W, Levine M. 2009c. Whole-mount in situhybridization on sea squirt (Ciona intestinalis) embryos. ColdSpring Harb Protoc (this issue). doi: 10.1101/pdb.prot5348.

Christiaen L, Wagner E, Shi W, Levine M. 2009d. Isolation of seasquirt (Ciona) gametes, fertilization, dechorionation, and devel-opment. Cold Spring Harb Protoc (this issue). doi: 10.1101/pdb.prot5344.

Christiaen L, Wagner E, Shi W, Levine M. 2009e. X-gal staining ofelectroporated sea squirt (Ciona) embryos. Cold Spring HarbProtoc (this issue). doi: 10.1101/pdb.prot5346.

Christiaen L, Wagner E, Shi W, Levine M. 2009f. Isolation of individ-ual cells and tissues from electroporated sea squirt (Ciona)embryos by fluorescence-activated cell sorting (FACS). Cold SpringHarb Protoc (this issue). doi: 10.1101/pdb.prot5349.

Corbo JC, Levine M, Zeller RW. 1997. Characterization of a noto-chord-specific enhancer from the Brachyury promoter region ofthe ascidian, Ciona intestinalis. Development 124: 589–602.

Davidson B, Shi W, Beh J, Christiaen L, Levine M. 2006. FGF signalingdelineates the cardiac progenitor field in the simple chordate,Ciona intestinalis. Genes & Dev 20: 2728–2738.

Dehal P, Satou Y, Campbell RK, Chapman J, Degnan B, De Tomaso A,Davidson B, Di Gregorio A, Gelpke M, Goodstein DM, et al. 2002.

The draft genome of Ciona intestinalis: Insights into chordate andvertebrate origins. Science 298: 2157–2167.

Delsuc F, Brinkmann H, Chourrout D, Philippe H. 2006. Tunicatesand not cephalochordates are the closest living relatives ofvertebrates. Nature 439: 965–968.

Deschet K, Nakatani Y, Smith WC. 2003. Generation of Ci-Brachyury-GFP stable transgenic lines in the ascidian Ciona savignyi. Genesis35: 248–259.

De Tomaso AW, Nyholm SV, Palmeri KJ, Ishizuka KJ, Ludington WB,Mitchel K, Weissman IL. 2005. Isolation and characterization of aprotochordate histocompatibility locus. Nature 438: 454–459.

Hendrickson C, Christiaen L, Deschet K, Jiang D, Joly JS, Legendre L,Nakatani Y, Tresser J, Smith WC. 2004. Culture of adult ascidiansand ascidian genetics. Methods Cell Biol 74: 143–170.

Imai KS, Hino K, Yagi K, Satoh N, Satou Y. 2004. Gene expressionprofiles of transcription factors and signaling molecules in theascidian embryo: Towards a comprehensive understanding ofgene networks. Development 131: 4047–4058.

Imai KS, Levine M, Satoh N, Satou Y. 2006. Regulatory blueprint fora chordate embryo. Science 312: 1183–1187.

Johnson DS, Zhou Q, Yagi K, Satoh N, Wong W, Sidow A. 2005.De novo discovery of a tissue-specific gene regulatory module ina chordate. Genome Res 15: 1315–1324.

Joly JS, Kano S, Matsuoka T, Auger H, Hirayama K, Satoh N, Awazu S,Legendre L, Sasakura Y. 2007. Culture of Ciona intestinalis inclosed systems. Dev Dyn 236: 1832–1840.

Kano S. 2007. Initial stage of genetic mapping in Ciona intestinalis.Dev Dyn 236: 1768–1781.

Matsuoka T, Awazu S, Satoh N, Sasakura Y. 2004. Minos transposoncauses germline transgenesis of the ascidian Ciona savignyi.Dev Growth Differ 46: 249–255.

Matsuoka T, Awazu S, Shoguchi E, Satoh N, Sasakura Y. 2005.Germline transgenesis of the ascidian Ciona intestinalis by electro-poration. Genesis 41: 67–72.

Matthysse AG, Deschet K, Williams M, Marry M, White AR, SmithWC. 2004. A functional cellulose synthase from ascidian epider-mis. Proc Natl Acad Sci 101: 986–991.

Nakatani Y, Moody R, Smith WC. 1999. Mutations affecting tail andnotochord development in the ascidian Ciona savignyi.Development 126: 3293–3301.

Nishida H. 1987. Cell lineage analysis in ascidian embryos by intra-cellular injection of a tracer enzyme. III. Up to the tissue restrictedstage. Dev Biol 121: 526–541.

Nishida H. 2002. Specification of developmental fates in ascidianembryos: Molecular approach to maternal determinants andsignaling molecules. Int Rev Cytol 217: 227–276.

These databases contain extensive spatiotemporal expression information for thousands of genesbased on systematic whole-mount in situ hybridization screens (Satou et al. 2002; Imai et al. 2004;Tassy et al. 2006). This information is not only useful to the Ciona research community, but can pro-vide potential insights into the activities of orthologous genes in vertebrates.

TECHNICAL APPROACHES

A variety of molecular tools have been applied to the understanding of Ciona embryogenesis.Protocols describing the practical use of Ciona for developmental studies include Isolation of SeaSquirt (Ciona) Gametes, Fertilization, Dechorionation, and Development (Christiaen et al. 2009d),Electroporation of Transgenic DNAs in the Sea Squirt Ciona (Christiaen et al. 2009a), X-galStaining of Electroporated Sea Squirt (Ciona) Embryos (Christiaen et al. 2009e), Microinjection ofMorpholino Oligos and RNAs in Sea Squirt (Ciona) Embryos (Christiaen et al. 2009b), Whole-Mount In Situ Hybridization on Sea Squirt (Ciona intestinalis) Embryos (Christiaen et al. 2009c),and Isolation of Individual Cells and Tissues from Electroporated Sea Squirt (Ciona) Embryos byFluorescence-Activated Cell Sorting (FACS) (Christiaen et al. 2009f).

REFERENCES




Nishida H, Sawada K. 2001. macho-1 encodes a localized mRNA inascidian eggs that specifies muscle fate during embryogenesis.Nature 409: 724–729.

Passamaneck YJ, Di Gregorio A. 2005. Ciona intestinalis: Chordatedevelopment made simple. Dev Dyn 233: 1–19.

Roure A, Rothbacher U, Robin F, Kalmar E, Ferone G, Lamy C, MisseroC, Mueller F, Lemaire P. 2007. A multicassette Gateway vectorset for high throughput and comparative analyses in Cionaand vertebrate embryos. PLoS ONE 2: e916. doi: 10.1371/journal.pone.0000916.

Sasakura Y, Konno A, Mizuno K, Satoh N, Inaba K. 2008. Enhancerdetection in the ascidian Ciona intestinalis with transposase-expressing lines of Minos. Dev Dyn 237: 39–50.

Satoh N. 2003. The ascidian tadpole larva: Comparative moleculardevelopment and genomics. Nat Rev Genet 4: 285–295.

Satoh N, Satou Y, Davidson B, Levine M. 2003. Ciona intestinalis:An emerging model for whole-genome analyses. Trends Genet19: 376–381.

Satou Y, Takatori N, Fujiwara S, Nishikata T, Saiga H, Kusakabe T,Shin-i T, Kohara Y, Satoh N. 2002. Ciona intestinalis cDNAprojects: Expressed sequence tag analyses and gene expressionprofiles during embryogenesis. Gene 287: 83–96.

Satou Y, Imai KS, Satoh N. 2004. The ascidian Mesp gene specifiesheart precursor cells. Development 131: 2533–2541.

Satou Y, Mineta K, Ogasawara M, Sasakura Y, Shoguchi E, Ueno K,Yamada L, Matsumoto J, Wasserscheid J, Dewar K, et al. 2008.Improved genome assembly and evidence-based global genemodel set for the chordate Ciona intestinalis: New insight intointron and operon populations. Genome Biol 9: R152. doi:10.1186/gb-2008-9-10-r152.

Seo HC, Kube M, Edvardsen RB, Jensen MF, Beck A, Spriet E, GorskyG, Thompson EM, Lehrach H, Reinhardt R, et al. 2001. Miniaturegenome in the marine chordate Oikopleura dioica. Science 294:2506. doi: 10.1126/science.294.5551.2506.

Shoguchi E, Kawashima T, Satou Y, Hamaguchi M, Sin IT, Kohara Y,Putnam N, Rokhsar DS, Satoh N. 2006. Chromosomal mappingof 170 BAC clones in the ascidian Ciona intestinalis. Genome Res16: 297–303.

Small KS, Brudno M, Hill MM, Sidow A. 2007. A haplome alignmentand reference sequence of the highly polymorphic Ciona savignyigenome. Genome Biol 8: R41. doi: 10.1186/gb-2007-8-3-r41.

Sordino P, Andreakis N, Brown ER, Leccia NI, Squarzoni P, Tarallo R,Alfano C, Caputi L, D’Ambrosio P, Daniele P, et al. 2008. Naturalvariation of model mutant phenotypes in Ciona intestinalis. PLoSONE 3: e2344. doi: 10.1371/journal.pone.0002344.

Tassy O, Daian F, Hudson C, Bertrand V, Lemaire P. 2006. A quantita-tive approach to the study of cell shapes and interactions duringearly chordate embryogenesis. Curr Biol 16: 345–358.




1Present address: Center for Developmental Genetics, Department ofBiology, Faculty of Arts and Science, New York University, New York,NY 10003, USA.2Corresponding authors ([email protected]); ([email protected]).Cite as: Cold Spring Harb Protoc; 2009; doi:10.1101/pdb.prot5344 www.cshprotocols.org

Protocol

Isolation of Sea Squirt (Ciona) Gametes, Fertilization,Dechorionation, and Development



INTRODUCTION

This protocol is the starting point for most manipulations that are used to study the sea squirt (Ciona)embryo, including in situ hybridization, the microinjection of morpholino oligos, and the electropo-ration of transgenic DNAs. Ciona eggs and embryos are exquisitely sensitive to even trace amounts ofdetergent; therefore, it is strongly advised to designate a soap-free workspace for embryo culture. Anysolutions that come into contact with embryos should be prepared in absolutely clean glassware withthe highest quality water.

RELATED INFORMATION

See The Sea Squirt Ciona intestinalis (Christiaen et al. 2009a) for an introduction to Ciona intestinalisas a model organism. Embryos that have been dechorionated as described here can be subsequentlyelectroporated or microinjected (see Electroporation of Transgenic DNAs in the Sea Squirt Ciona[Christiaen et al. 2009b] and Microinjection of Morpholino Oligos and RNAs in Sea Squirt (Ciona)Embryos [Christiaen et al. 2009c], respectively).

MATERIALS

CAUTIONS AND RECIPES: Please see Appendices for appropriate handling of materials marked with <!>, andrecipes for reagents marked with <R>.

Reagents

<!>Ampicillin (100 mg/mL) (for cultivation of larvae and juveniles only; see Steps 26-30)<!>Alternatively, use penicillin-streptomycin (5000 U/mL penicillin plus 5000 g/mL streptomycin).

Ciona animals, gravid<R>Filtered artificial seawater buffered with TAPS (FASW-T)

FASW-T is the only seawater that should come into contact with eggs and embryos! Large amounts ofFASW-T are consumed for each experiment, so have 2-3 L ready before beginning. It is desirable to have oneor two plastic squirt bottles filled with FASW-T available as well.

Prepare FASW-T containing 0.05% (w/v) bovine serum albumin (BSA) for cultivating larvae and juveniles (seeSteps 26-30).

Glycine (10 mg/mL, prepared in FASW-T) (for dechorionation only; see Steps 14-25)<!>NaOH (10 N)Pronase (Pronase E, embryo-tested) (for dechorionation; see note at beginning of “Fertilization”

and Steps 14-25)




METHOD

Isolation of Eggs and Sperm

In Steps 1-8, eggs are collected from each individual into separate wells of a six-well plate. This will limit precocious fer-tilization in the event that the sperm duct is accidentally pierced during egg retrieval. Sperm are collected separatelyin 1.5-mL tubes.

1. Choose at least two healthy, gravid animals; such animals will respond to a gentle touch by con-tracting their siphons.

2. Expose the egg duct (Fig. 1) of each animal as follows:

i. Hold the animal upside down with the atrial siphon opening between the fingertips, pullinggently to elongate the siphon.

Prepare Pronase ahead of time by dissolving the lyophilized powder in FASW-T at a final concentration of 2.5%(w/v). Aliquot in 400- or 800- L portions and store at −80°C.

<!>Sodium thioglycolate (for dechorionation; see note at beginning of “Fertilization” and Steps14-25)

Equipment

<R>Agarose-coated Petri plates (60 × 15 mm) containing FASW-TCiona embryos that develop from dechorionated eggs require a thin layer of agarose on which to settle, to pre-vent irreversible adherence to the plastic. Uncoated plates may be used to raise juveniles (see Step 28).

Aluminum foilConical tubes (50 mL)

After use, rinse the 50-mL conical tubes thoroughly in double-distilled H2O. They can be reused indefinitely.

Dissecting microscopeFilter basket

This is a handy contraption used to contain eggs while they are thoroughly rinsed with FASW-T. It consists ofa small cell strainer (pore size 70-80 m) attached to the end of a conical tube (Fig. 1). To make it, remove thecap from a 50-mL tube, and use a flame to slightly melt the plastic around the open end. Then affix the strainerto the melted surface, ensuring even contact, and hold until it is set. Finally, use a flame to heat a scalpel andremove the conical portion of the tube at about the 20-mL mark. This filter basket is reusable indefinitely; thor-oughly rinse in double-distilled H

2O after each use.

Forceps (#5), sharpGlass jars

Each glass jar should be able to hold a 50-mL conical tube and should be about two-thirds the depth of the50-mL conical. We use 8-oz glass jars from Fisher.

Ice bucket with iceIncubator(s) at 13°C and/or 16°C, equipped with fan (e.g., Vinotemp wine cellar) (see Step 7)Lab tissuesMicrocentrifuge tubes (1.5 mL)Microscope slides (glass) or plastic Petri dish lids (optional; see Steps 11 and 13)Pasteur pipettes and bulbsPlastic bags, sealablePlastic beakers (1 or 2 L)Scissors (small, sharp ones for dissecting)Temperature-controlled room at 18°C (for cultivation of larvae and juveniles only; see Steps 26-30)

A temperature-controlled room or small incubator is necessary to ensure synchronous development of embryos.Small fluctuations in temperature can cause embryos to develop at different rates; whether or not this is a majorissue depends on the nature of the experiment.

Timer (see Step 12 and “Dechorionation”)Tissue culture plates (six-well)

After use, rinse the six-well plates thoroughly in double-distilled H2O. They can be reused indefinitely.




ii. Position the scissors to cut through the body wall muscle to the tunic.

iii. Use the squirt bottle to rinse away waste products from the intestine and occasionalamphipods.

iv. Carefully cut away any connective tissue, such as body wall muscle, from the surface of thegonad ducts.

Do this over a large plastic beaker to collect water waste.

3. When the egg duct is exposed, position the animal over a well of a six-well plate that is half full ofFASW-T (~5 mL FASW-T/well). Gently pierce the surface of the egg duct with a sharp #5 forceps,or the sharp end of scissors.The proximity of the sperm duct can make this challenging, especially if the egg duct is not stretched to capac-ity. It helps to have a sharp tool to make the smallest hole possible and avoid puncturing the sperm duct at thesame time. Because the freshly harvested eggs have an intact chorion, they will not adhere to the plastic ofthe six-well plate.

4. Apply gentle pressure to the egg duct with the flat end of the scissors, guiding the eggs into thewell. Continue until the duct is depleted.The eggs exist as a cohesive unit in mucilage; minimize disturbance of the egg mass.

5. (Optional) Look at the eggs under the dissecting microscope.Healthy eggs are deeply pigmented and surrounded by oblong follicle cells, which resemble rays of the sun.You will see a golden ring around the egg; this is the chorion.

6. Blot the surface of the ducts with a tissue to absorb excess seawater; it is desirable to collect thesperm “dry.”

7. Pierce the sperm duct with a sharp forceps or scissors, and collect the sperm with a Pasteur pipette.Keep sperm in a 1.5-mL tube on ice, and pool from each individual.At this point, sperm can be stored on ice or at 4°C-13°C for several hours or even a day or two. Eggs can rest inFASW-T for up to 12 h at 16°C.

8. Collect the spent animals in a sealable plastic bag. Freeze the animals to reduce foul odors untilproper disposal is convenient.

Fertilization

If fertilization will be followed by dechorionation in Steps 14-25, have 0.4 mg of sodium thioglycolate in a 50-mLconical tube ready and 800 L of Pronase thawing on ice before beginning the fertilization.

9. Activate the sperm at an elevated pH in the presence of eggs as follows:

i. Prepare basic seawater by adding 10 L of 10 N NaOH to 40 mL of FASW-T in a labeled50-mL conical tube. Invert several times to mix.

ii. Add 20-50 L of sperm, and mix again.

iii. Use a Pasteur pipette to add a small number of eggs (a small drop of 10-20 eggs issufficient) to the sperm.

This is not necessary if you see that eggs are present in your collected sperm samples.

FIGURE 1. (A) Dissected C. intestinalis adult. Individualeggs within the duct of the C. intestinalis adult can bediscerned (red) and lie adjacent to the sperm duct(white). (Arrow) Body wall muscle that overlies gameteducts and should be carefully cut away prior to gameteretrieval. (B) The glass jars and homemade basketused to work with eggs. (For color figure, see doi:10.1101/pdb.prot5344 online at www.cshprotocols.org.)




iv. Let activation proceed for 5-10 min.

10. While the sperm are activating, wash the eggs as follows:

i. Place the filter basket in a glass jar and fill halfway with FASW-T.

ii. Transfer the eggs en masse with a Pasteur pipette from each well of the six-well plate to thesubmerged basket.

iii. Squirt FASW-T along the side of the tube, and swirl the basket, keeping the eggs sub-merged. Do not squirt directly onto the eggs, because this may damage them.

iv. Discard the water as the jar becomes full, and continue adding more water and swirling thebasket to distribute the eggs.

v. Gently pipette to help separate and wash the eggs, but be careful not to expel them withforce against the mesh filter.

11. (Optional) Check the sperm activation by transferring a drop to a plastic Petri dish lid or a micro-scope slide, and look under a dissecting microscope.The sperm are very small, but you should be able to see their extremely rapid motion.

12. Discard water from the egg jar, leaving just enough to keep the eggs submerged. Pour the dilutedsperm onto the eggs, and start the timer, counting up.The time of sperm addition is the “zero” point; developmental time is usually measured in hours post-fertiliza-tion (hpf). Begin dechorionation (starting at Step 14) 5 min after fertilization. If dechorionation is not required,the eggs can be incubated to the desired stage (see Steps 26-30). For most research purposes, however, dechori-onation is required.

13. (Optional) Check the fertilization after a few minutes by pipetting a drop of the mixture onto aslide or plastic Petri dish lid and examining under a dissecting microscope.The eggs will spin in circles if the sperm is very abundant. Subtle changes in cell shape due to ooplasmic segre-gation may also be apparent.

Dechorionation

Dechorionation is an essential component of Ciona research, as it is a prerequisite for both electroporation and microin-jection. The ease with which it is accomplished can vary from animal to animal, or from differences in activity frombatch to batch of Pronase. Dechorionated eggs must be washed several times to remove cellular debris.Dechorionation and washing (Steps 14-25) should ideally be completed by 15-25 min post-fertilization if the eggsare to be used for electroporation.

14. Prepare the dechorionation solution as follows:

i. Add FASW-T to the 0.4 mg of sodium thioglycolate prepared in the 50-mL conical tube,to almost 40 mL. Invert to mix.

ii. Add 168 L of 10 N NaOH, and mix again.

iii. Add 800 L of 2.5% Pronase and mix.

iv. Bring the final volume to 40 mL with FASW-T.

15. Pour roughly two-thirds of the dechorionation solution into a glass jar.

16. Quickly transfer the egg basket from the fertilization solution to the dechorionation solution. It isimportant not to let the eggs dry.

17. Pour the remaining dechorionation solution into the basket, on top of the eggs.

18. Empty the FASW-T from one agarose-coated Petri dish, and prop one end up on the lid.The plate at this angle allows the eggs to be collected in a small surface area to limit the direct contact of eggswith the agarose and subsequent cell lysis.

19. Transfer the eggs from the basket to the tilted plate with a Pasteur pipette. Take care to gentlyrelease the eggs with the pipette tip submerged. As more liquid is added, lay the plate flat.

20. Gently and steadily pipette the dechorionation solution to form a current:




i. Hold the pipette at a low angle, and aim the water flow tangentially to the wall of the plate.

ii. Create circular movement of the liquid containing the eggs, but do not draw the eggsthemselves into the pipette.

This provides mechanical force that works in conjunction with the chemical treatment to facilitate chorionremoval.

21. Monitor the dechorionation reaction under the dissecting microscope, and continue pipetting tomaintain the current.When the eggs start to dechorionate, the water may become cloudy and yellow, usually after 2-3 min. The firsteggs to dechorionate will typically lie in the center of the egg pile and may become elongated. Within 3-5 min,most of the eggs will start to lose the chorion.

See Troubleshooting.

22. Add 1 mL of 10 mg/mL glycine to stop the reaction, dilute by squirting a bit of FASW-T, andpipette a bit more for thorough mixing.It is better to stop the reaction a bit early, with decreased dechorionation efficiency, than to over-dechorionatethe eggs. The dechorionated eggs will be washed and isolated away from those that are not dechorionated inStep 25. Overexposure to the Pronase/thioglycolate solution can have adverse effects on embryo development.

23. Collect the dechorionated eggs by swirling the plate with gentle pressure in small circles on thebenchtop.This will create subtle movement of the water, and the eggs will coalesce in the center of the plate.Dechorionated eggs appear brown to purple in color and settle on the agarose. Eggs with the chorion still intactappear yellow to orange and tend to float on the surface and around the edge of the dechorionated egg mass.

24. Use a Pasteur pipette to transfer the dechorionated eggs to a fresh plate with FASW-T. Ensure thatthe pipette is submerged, and gently distribute the eggs throughout the plate.Dechorionated eggs are extremely fragile and will explode upon surface contact. Care must be taken when pipet-ting dechorionated eggs to gently release them under the surface of the water.

25. Swirl to collect the eggs again, and continue to transfer to fresh plates with FASW-T, six to eightmore times.It is helpful for the beginner to look in the microscope to monitor the eggs at each successive transfer. Try totransfer as many of the dechorionated eggs as possible, while leaving the non-dechorionated eggs and cell debrisbehind. After six to eight washes, you should have a uniform population of dechorionated eggs.

If dechorionated eggs are to be electroporated or microinjected, proceed immediately to the appropriate proto-col (Electroporation of Transgenic DNAs in the Sea Squirt Ciona [Christiaen et al. 2009b] or Microinjectionof Morpholino Oligos and RNAs in Sea Squirt (Ciona) Embryos [Christiaen et al. 2009c], respectively).Otherwise, continue to Step 26; fertilized eggs keep developing and cannot be arrested or stored.

Cultivation of Larvae and Juveniles

26. Because culture of larvae and juveniles occurs over a longer time period, plate the embryos inagarose-coated Petri plates containing FASW-T supplemented with penicillin-streptomycin (at finalconcentrations of 50 U/mL and 50 g/mL, respectively) or ampicillin (at a final concentration of100 g/mL). Distribute the eggs from Step 25 among several plates. Leave the dechorionated eggsto develop to the desired stage in a temperature-controlled room at 18°C.It is critical to achieve uniform distribution of the eggs, such that direct contact among them is minimized. Theeggs have a natural tendency to cluster, which results in aggregates of malformed embryos caused by cell fusion.This is not a concern if the eggs have intact chorions.


27. When the tadpoles begin to swim, dilute them into many separate plates.The tadpole larva forms ~18 hpf at 18°C. Note that when the larvae begin to swim, the collision between indi-viduals results in their irreversible adherence to one another, and can result in large clumps of tadpoles withtwitching tails. The only practical way to limit tadpole clumping is to dilute the tadpoles into many separateplates. In addition, add 0.05% BSA (w/v) to the FASW-T to minimize clumping between the larvae. Tadpolesthat hatch from eggs with intact chorions do not have this problem, and can swim unfettered at high density.

28. If raising juveniles, decide whether to raise them on uncoated plates or on agarose-coated plates.Swimming larvae will settle on the bottom of uncoated plates and attach tightly, allowing easier water changeand long-term culturing over periods of weeks. If early juveniles (i.e., 1-4 d after hatching) are desired, an




Christiaen L, Wagner E, Shi W, Levine M. 2009a. The sea squirt Cionaintestinalis. Cold Spring Harb Protoc (this issue). doi: 10.1101/pdb.emo138.

Christiaen L, Wagner E, Shi W, Levine M. 2009b. Electroporation oftransgenic DNAs in the sea squirt Ciona. Cold Spring Harb Protoc

(this issue). doi: 10.1101/pdb.prot5345.Christiaen L, Wagner E, Shi W, Levine M. 2009c. Microinjection of

morpholino oligos and RNAs in sea squirt (Ciona) embryos. ColdSpring Harb Protoc (this issue). doi: 10.1101/pdb.prot5347.

alternative way is to grow juveniles on agarose-coated plates. In this case, the larvae will not attach to agarosebut to each other or to other debris in the dish. Because the juveniles then float in seawater, it is easier to removethem for experimental manipulations like fixation or live imaging.

29. To raise juveniles, use a Pasteur pipette to transfer late-tailbud-stage embryos (Stage 24-25) tofresh plates (either uncoated plates or agarose-coated plates) with antibiotic-supplementedFASW-T containing 0.05% (w/v) BSA.The tadpoles are now swimming larvae until they settle on a substrate and commence metamorphosis. Thisdramatic rearrangement of the body plan culminates in a juvenile tunicate. Most of the tadpoles will settle andcommence metamorphosis within a couple of days.


30. Cover the plates in aluminum foil to shield them from light. Change the water daily.

TROUBLESHOOTING

Problem: Dechorionation is ineffective.[Step 21]Solution: Consider the following:

1. Try a different batch of Pronase and/or fresh sodium thioglycolate.

2. Make sure that the eggs are washed thoroughly.

Problem: Embryos do not develop.[Step 26]Solution: Consider the following:

1. Ensure that the sperm are activated.

2. Make sure that the eggs are fertilized.

Problem: Tadpoles do not settle.[Step 29]Solution: The tadpoles may settle more securely on plates coated with 0.05% (w/v) BSA in FASW-T

rather than agarose. This is a matter of preference for the experimenter.

REFERENCES





Protocol

Electroporation of Transgenic DNAs in the Sea Squirt Ciona



INTRODUCTION

The electroporation method described here is probably the mainstay of sea squirt (Ciona) research,because the utility of transgene expression in staged embryo populations enables a wide array of bio-logical questions to be addressed. It allows rapid identification and characterization of cis-regulatoryDNA, such as tissue-specific enhancers. Electroporation of plasmids expressing fluorescent reportergenes permits live imaging and lineage tracing. Finally, structure-function relationships can be exam-ined by expressing dominant-negative or constitutively active forms of specific proteins using appro-priate cell-specific enhancers.

RELATED INFORMATION

See The Sea Squirt Ciona intestinalis (Christiaen et al. 2009a) for an introduction to Ciona intestinalisas a model organism. For related methods, see Isolation of Sea Squirt (Ciona) Gametes, Fertilization,Dechorionation, and Development (Christiaen et al. 2009b).

MATERIALS


Reagents

Batch of dechorionated Ciona eggs on agarose-coated plates containing FASW-T (see Isolationof Sea Squirt (Ciona) Gametes, Fertilization, Dechorionation, and Development[Christiaen et al. 2009b])The optimal egg density in a 60- × 15-mm dish is probably 300-500. Up to 10 electroporations can be per-formed during the same session (i.e., from the same batch of eggs).

D-Mannitol (0.96 M)Dissolve the D-mannitol in highly purified H

2O (e.g., GIBCO/Invitrogen) to a final concentration of 0.96 M;

gentle heating is required for complete solubilization. Prepare 50-500 mL at a time and filter through 0.22-µmmesh. Aliquots can be stored at −20°C. (It is also stable at room temperature but tends to crystallize over time.)

Plasmid DNAs of interest (e.g., Macherey-Nagel midi-preps)Highly purified plasmid DNA must be prepared in large quantities and at high concentrations (2-5 µg/µL) foruse in Step 1.

Equipment

<R>Agarose-coated plates (60 × 15 mm) containing FASW-T




METHOD

Label the tubes and plates for each sample you wish to electroporate ahead of time. Work quickly; the goal is to haveeach sample electroporated before 30 min post-fertilization. This is roughly 30 min before the time of the first cleav-age, which bisects the egg into left and right blastomeres.

Plasmid Preparation

For each sample you wish to electroporate, you will need a 1.5-mL tube containing the plasmids prepared as describedhere.

1. Combine the desired plasmids in a single 1.5-mL tube, and add H2O to a final volume of 100 µL.

A typical electroporation sample contains one to three different constructs, 20-120 µg of each, in a total volumeof 100 µL.

2. Add 400 µL of 0.96 M D-mannitol to the tube, mix well, and centrifuge at 12,000g for 10 sec.This is your electroporation mix. The final D-mannitol concentration should be 0.77 M.

Electroporation

3. Swirl to collect the dechorionated eggs in the center of the plate.Set aside a small amount of eggs to develop as a “fertilization-only” control. The quality of your electroporatedembryos can be compared to this control sample.

4. Distribute the eggs among the 2-mL low-adhesion tubes with a Pasteur pipette. Make sure thatthe pipette tip is always submerged. Let the eggs settle to the bottom of the tube; this occursquickly.The number of tubes used will vary, depending on egg availability and the requirements of the experiment.

5. Use a vacuum aspirator with a fine tip to reduce the volume of FASW-T to 0.2 mL; graduated tubesare thus essential here.

6. Using a Pasteur pipette, transfer the electroporation mix to one tube of eggs. Gently mix.The high conductivity of seawater necessitates its dilution prior to electroporation. In this step, osmotic balance(and thus cell integrity) is maintained by replacing a portion of the FASW-T with a concentrated D-mannitolsolution.

7. With the same pipette, transfer the egg-plasmid-D-mannitol mixture to a cuvette. Position thecuvette as necessary in the electroporation apparatus. Press the button, and release when you hearthe beep. Note the time constant.The ideal time constant is 16-18 msec. Smaller numbers will result in decreased electroporation efficiency,whereas larger numbers will destroy more eggs and can delay development.


If handling more than 300-500 eggs, use Petri dishes that are larger than 60 × 15 mm because settledembryos have a tendency to adhere to each other during development.

Electroporation apparatus (e.g., Gene Pulser II, 0.4-cm cuvettes, and capacitance extender fromBio-Rad)Set the electroporator voltage to 50 V and the capacitance to 1.00 F. Have one clean cuvette ready for eachsample. Cuvettes can be rinsed thoroughly in H

2O and reused five to 10 times.

Low-adhesion tubes (2-mL, graduated)Embryos do not seem to stick to the 2-mL tubes with catalog number 1480-2700 from USA Scientific.

These low-adhesion tubes should be rinsed after use, and they can be reused indefinitely. The eggs are lessprone to sticking to previously used tubes, so reuse is strongly advised.

MicrocentrifugeMicrocentrifuge tube rack with 1.5-mL tubesPasteur pipettes with bulbsVacuum aspiration unit




8. Retrieve the eggs from the cuvette, and divide evenly between two 60-mm agarose-coated platescontaining FASW-T. Take care to minimize aggregation of the eggs.

9. Repeat Steps 6-8 for each sample, using a clean pipette for each.

10. Let the embryos develop to the desired stage (for details, see Isolation of Sea Squirt (Ciona)Gametes, Fertilization, Dechorionation, and Development [Christiaen et al. 2009b]).Mosaic inheritance of plasmid DNA is a common occurrence and does not generally present problems. Theexperimenter should keep this in mind, however, when analyzing the results.


TROUBLESHOOTING

Problem: There are variable time constants during electroporation.[Step 7]Solution: Consider the following:

1. Use new cuvettes.

2. Check that the salinity of FASW-T is 3.4%.

Problem: Embryos fail to develop to advanced stages.[Step 10]Solution: Consider the following:

1. Compare electroporated embryos to the fertilization control. If the problem is specific to the elec-troporated embryos, try a different preparation of D-mannitol.

2. Ensure that the DNA is clean.

3. If the fertilization control embryos also fail to develop, then there is probably a problem with thedechorionation. Use fresh chemicals and animals.

REFERENCES


Christiaen L, Wagner E, Shi W, Levine M. 2009b. Isolation of sea

squirt (Ciona) gametes, fertilization, dechorionation, and devel-opment. Cold Spring Harb Protoc (this issue). doi: 10.1101/pdb.prot5344.



X-gal Staining of Electroporated Sea Squirt (Ciona) Embryos



INTRODUCTION

This protocol describes the fixation, staining, and mounting of sea squirt (Ciona intestinalis) embryosand larvae that have been electroporated with a plasmid DNA containing a cis-regulatory DNA (e.g.,tissue-specific enhancer) fused to the lacZ reporter gene. Green fluorescent protein (GFP) reportergenes can be directly visualized in living embryos and larvae, although fixed preparations can useaspects of this protocol.

RELATED INFORMATION

See The Sea Squirt Ciona intestinalis (Christiaen et al. 2009a) for an introduction to C. intestinalis asa model organism. For related methods, see Electroporation of Transgenic DNAs in the Sea SquirtCiona (Christiaen et al. 2009b).



Protocol

MATERIALS


Reagents

C. intestinalis embryos, electroporated with plasmid DNA containing a cis-regulatory elementfused to a lacZ reporter gene as described in Electroporation of Transgenic DNAs in theSea Squirt Ciona (Christiaen et al. 2009b)

<R>LacZ buffer (LZB)<R>MEM-GAMounting medium (e.g., 50%-80% glycerol in 1X PBS)Nail varnish, transparent (optional; see Step 15)<R>Phosphate-buffered saline (PBS) (1X) containing 0.05%-0.1% Tween 20 (PBT)

The stock solution of PBS should be prepared without CaCl2or MgCl

2.

<!>PBT containing 4% paraformaldehyde (PFA)X-gal stock solution (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside) (40 mg/mL)

<!>Prepare the stock solution in DMF (dimethylformamide).

<!>Alternatively, use DMSO (dimethylsulfoxide) as the solvent for preparing the X-gal stock solution.

Equipment

Dissecting microscope




METHOD

Fixation

1. For each experiment, label one 1.5-mL tube and fill it with 0.5 mL of MEM-GA. Transfer theembryos to the tube using a Pasteur pipette, and let them settle for 2-5 min.See Troubleshooting.

2. Remove the supernatant, and replace it with 1 mL of fresh MEM-GA. Nutate for 20 min at roomtemperature.

3. Let the embryos settle for 5 min.

4. Replace the supernatant with 1 mL of PBT, and nutate for 10 min.

5. Repeat Steps 3 and 4 three to five times.

Staining

6. Replace the PBT with 0.75 mL of LZB, and let the embryos settle.

7. While the embryos are settling, prepare the staining solution by adding 2 µL of 40 mg/mL X-galstock solution to 1 mL of LZB, for a final concentration of 80 µg/mL X-gal.

8. Replace the LZB with the staining solution. Transfer the embryos to the staining dish.

9. Incubate the embryos in the staining dish at 37°C.

10. Monitor staining development on a dissecting microscope (typically 30 min to 2 h).See Troubleshooting.

Post-fixation and Mounting

11. Transfer the embryos to the original tubes. Rinse the embryos once with PBT, and then replace thePBT with 1 mL of PBT containing 4% PFA. Nutate the embryos in PBT containing 4% PFA for 1 hat room temperature.

12. Rinse the embryos with PBT three times.

13. Replace the PBT with mounting medium. Let the embryos equilibrate for 3-5 min.

14. Mount the embryos as follows:

i. Position narrow (~2-mm) stripes of double-sided tape on the long sides of the glass slide.

ii. Pipette ~150-200 µL of mounting medium containing the stained embryos onto the slide.

iii. Carefully position the coverslip.

iv. Gently press down on the sides with double-sided tape.

v. Remove excess mounting medium.

15. (Optional) Seal the coverslip with transparent nail varnish for long-term storage.

Double-sided tapeGlass slides and coverslipsIncubator preset to 37°CMicrocentrifuge tubes (1.5 mL)NutatorPasteur pipettes and bulbsStaining dish (e.g., a 24-well cell culture dish rinsed with hot tap water and then with distilled

H2O)



TROUBLESHOOTING

Problem: The embryos stick to the Pasteur pipette during transfer.[Step 1]Solution: Consider the following:

1. Pipette seawater several times before transferring the embryos, or use pretreated pipettes (e.g.,treated overnight in distilled H

2O and dried).

2. Alternatively, raise embryos in seawater supplemented with 0.05% bovine serum albumin (BSA).

Problem: Larvae do not settle quickly to the bottom of the tube.[Step 1]Solution: Centrifuge at very low speed (100g) for 30 sec.

Problem: No staining appears, even after overnight incubation.[Step 10]Solution: Most constructs will yield ectopic staining (e.g., in the mesenchyme). Complete absence of

staining can be due to one of the following:

1. An inappropriate fixative (paraformaldehyde instead of glutaraldehyde) has been used.

2. There is a frameshift in the reporter construct (e.g., some coding sequence from the gene ofinterest has been inadvertently included with the cis-regulatory DNA but is not in the same frameas the lacZ coding sequence).

3. The construct is not active. In this case, in situ hybridization with a lacZ-specific antisense probe orreverse transcriptase polymerase chain reaction (RT-PCR) using lacZ-specific primers can be usedto confirm that the construct is actively being transcribed.

REFERENCES


Christiaen L, Wagner E, Shi W, Levine M. 2009b. Electroporation oftransgenic DNAs in the sea squirt Ciona. Cold Spring Harb Protoc(this issue). doi: 10.1101/pdb.prot5345.






Protocol

Microinjection of Morpholino Oligos and RNAs in Sea Squirt(Ciona) Embryos



INTRODUCTION

This protocol describes microinjection of morpholino oligos (MOs) and RNAs into sea squirt (Ciona)embryos. This is the method of choice for gene disruption assays. MOs that target the initiating ATGcan be used, in addition to those that target splice donor and acceptor sites. The latter method per-mits the selective inhibition of zygotic mRNAs in cases in which the gene in question is expressed inboth the egg and embryo. Although injection is usually performed at the one-cell stage, it is possibleto target individual blastomeres, up to the eight-cell stage, thereby permitting lineage-specific knock-down of pleiotropic genes. Injection can also be performed in unfertilized eggs to inhibit maternalgenes. Microinjection also permits DiI labeling and lineage tracing.

RELATED INFORMATION

See The Sea Squirt Ciona intestinalis (Christiaen et al. 2009a) for an introduction to Ciona intestinalisas a model organism. For related methods, see Isolation of Sea Squirt (Ciona) Gametes,Fertilization, Dechorionation, and Development (Christiaen et al. 2009b) and X-gal Staining ofElectroporated Sea Squirt (Ciona) Embryos (Christiaen et al. 2009c).

MATERIALS


Reagents

Ciona embryos or eggs and sperm (see Isolation of Sea Squirt (Ciona) Gametes, Fertilization,Dechorionation, and Development [Christiaen et al. 2009b])

Injection mix<!>The injection mix consists of nucleic acids (e.g., MOs, DNA, or RNA) and dye (e.g., Fast Green FCF orAlexaFluor 555). Prepare and use the components as follows:

For MOs, resuspend 300 nmol of the specific MO in 100 µL of RNase-free H2O (not diethyl pyrocarbonate

[DEPC]-treated) to make a 3 mM stock solution. Store at room temperature because MOs seem to go out ofsolution when stored at −80°C. Inject these MOs at 0.5-1.5 mM.

Use capped mRNA (in vitro transcription product, stored at −80°C) at 0.2-1 µg/µL.

Use plasmid DNA at 5-20 ng/µL.




METHOD

Preparation of Injection Device

1. Place the capillaries in aluminum foil with an open tube of Sigmacote in a vacuum desiccator.Apply vacuum for 10 min and leave the capillaries overnight to evenly coat the glass walls.This step siliconizes the needles to reduce the amount of DNA or RNA sticking to the side of the glass capillaries.

2. Prepare the needles as follows:

i. Pull the needles with a micropipette puller. Use the automatic test program on the machineto determine the optimal heat constant of the particular batch of glass needles. Experimentwith other parameters to produce a thin needle with enough stiffness.

ii. Cut the needle ~3 cm from the tip using a sand stone. Hold the needle tight when cuttingto avoid breaking off the tip.

iii. Store the needles in a dust-free, vibration-free container.

3. Pull thin loading capillaries (0.1 mm) over a Bunsen burner.

4. Fill the injection needle as follows:

i. Connect the loading capillary to a 1-mL syringe with PVC tubing.

ii. Centrifuge the injection mix before loading to prevent needle clogging.

iii. Load 0.2-0.5 µL of injection mix to the tip of the injection needle.

<!>For Fast Green (Fast Green FCF; Sigma), prepare a 5 mg/mL stock in H2O, filter (0.2 µm), and store at 4°C.

Use at a final concentration of 0.2-0.5 mg/mL.

For fluorescent dextran dye (e.g., AlexaFluor 555 [10 kDa]; Molecular Probes/Invitrogen), prepare a 10 mMstock in H

2O, filter (0.2 µm), and store at −20°C. Use at a final concentration of 0.3 mM.

Mineral oil<!>Sigmacote (Sigma)

Equipment

<R>Agarose-coated plates containing FASW-T (60- × 15-mm plates for growing embryos; 100- ×15-mm plates for injection)

Aluminum foilBunsen burnerCentrifugeContainer for storing needlesDissecting microscope (e.g., Olympus SZX-12)Egg transfer pipettes (pulled from 3-mm glass tube)Fluorescence microscopeGlass coverslips (25 × 25 mm)Glass syringe (5 mL)Hot metal needle (for blastomere injection or ablation only; see Steps 15-17)Injection holder (Narishige)Micromanipulator (3D) and microscope mounting adaptor (Narishige)Micropipette puller (Sutter Instruments)Oil hydraulic micromanipulator (Narishige)Polyvinyl chloride (PVC) tubing (2 mm)Sand stoneSyringes (disposable, 1 mL)Thin-wall glass capillary with filament (e.g., World Precision Instruments TW100F-4)Vacuum desiccator




iv. Set up a similar loading capillary and syringe filled with mineral oil.

v. Fill the rest of the injection needle with mineral oil and avoid air bubbles.

5. Prepare the injection device on the micromanipulator as follows:

i. Connect the 5-mL injection syringe to the injection holder with PVC tubing. Fill with dou-ble-distilled H

2O.

ii. Insert the injection needle into the rubber gasket, and tighten the holder. Make sure theairway is completely filled with H

2O and avoid air bubbles.

iii. Position the injection device on the micromanipulator at an angle of ~30° relative to thesurface.

Injection

6. Place a 25- × 25-mm glass coverslip in the center of a 100-mm agarose-coated Petri dish.

7. Transfer approximately 100 eggs to the Petri dish and crush them against one side of the cover-slip to coat the glass.This step prevents the eggs from sticking.

8. Break open the injection needle by pushing it against the edge of the coverslip. Then, push the 5-mL syringe to make sure that the colored solution oozes from the needle tip.This produces a faint cloud that quickly disappears.

9. Use a transfer pipette to dispense approximately 100 eggs along the coated edge of the coverslip.Slide the coverslip toward the eggs to align them against the edge.

10. Inject the egg as follows:

i. Position the needle against the middle of the first egg at an ~30° angle.

ii. Push the needle into the egg using the hydraulic micromanipulator, and pull back thesyringe to break the seal between the needle tip and egg cortex.

iii. Push the syringe to dispense the injection mix into the egg.A small smear of dye will appear at the site of the needle tip but quickly disperse.


11. Pull out the needle.See Troubleshooting.

12. Move to the next egg, and repeat Steps 10 and 11.

13. After injecting the aligned eggs, transfer them to a 60-mm agarose-coated dish for incubation.If injecting unfertilized eggs, add diluted sperm to the dish, and let them fertilize for 10 min before rinsing offthe sperm.

14. Screen the embryos as follows:

For fluorescent dextran-injected embryos, screen the embryos under a fluorescence microscopeand remove uninjected embryos.

For embryos co-injected with a DNA plasmid, screen for the marker (green fluorescent protein[GFP], lacZ, etc.) after embryo fixation/staining. (see X-gal Staining of Electroporated Sea Squirt(Ciona) Embryos [Christiaen et al. 2009c]).

Blastomere Injection or Ablation

Use the following steps in place of Steps 6-9 for blastomere injection or ablation.

15. Use a hot metal needle to poke small pits about the size of an embryo into the agarose layer in aPetri dish.

16. Transfer individual embryos to the pits, and orient them so that the blastomere of interest isfacing the surface.





Christiaen L, Wagner E, Shi W, Levine M. 2009b. Isolation of seasquirt (Ciona) gametes, fertilization, dechorionation, and devel-

opment. Cold Spring Harb Protoc (this issue). doi: 10.1101/pdb.prot5344.

Christiaen L, Wagner E, Shi W, Levine M. 2009c. X-gal staining ofelectroporated sea squirt (Ciona) embryos. Cold Spring HarbProtoc (this issue). doi: 10.1101/pdb.prot5346.

17. For blastomere injection, proceed as for injecting 1-cell embryos. For cell ablation experiments, jabthe blastomere until it bursts.

TROUBLESHOOTING

Problem: Embryos develop too fast, resulting in insufficient time for injection.Solution: Consider the following:

1. To extend the working time, either inject in a temperature-controlled room or use a water-cooledmetal stage to slow development.

2. Alternatively, inject unfertilized eggs and fertilize them later.

Problem: The needle tip gets clogged during injection.[Step 10]Solution: Break the needle at the tip or use a new needle.

Problem: The embryo sticks to the needle.[Step 11]Solution: Use a sudden pull to shake off the egg.

REFERENCES





Protocol

Whole-Mount In Situ Hybridization on Sea Squirt (Cionaintestinalis) Embryos



INTRODUCTION

This protocol describes whole-mount in situ hybridization on sea squirt (Ciona intestinalis) embryos.This method is a mainstay of the sea squirt (Ciona) research community. It permits the detailed visu-alization of gene expression at single-cell resolution. It has been used to detect localized maternalmRNAs in the fertilized egg and to identify restricted patterns of gene expression within individualcells of the developing central nervous system at advanced stages of development, including swim-ming tadpoles.

RELATED INFORMATION

See The Sea Squirt Ciona intestinalis (Christiaen et al. 2009a) for an introduction to C. intestinalis asa model organism. For related methods, see Isolation of Sea Squirt (Ciona) Gametes, Fertilization,Dechorionation, and Development (Christiaen et al. 2009b).

MATERIALS


Reagents

Antibody solution (anti-DIG Fab fragments, diluted 1/2000 in TNB)For best results, pre-absorb a 1/10 dilution of the antibody with blocked embryos and/or larvae prior to prepar-ing the final (1/2000) dilution and incubating with hybridized samples.

Dechorionated Ciona embryos from Isolation of Sea Squirt (Ciona) Gametes, Fertilization,Dechorionation, and Development (Christiaen et al. 2009b)

<!>Digoxigenin (DIG)-labeled RNA probe<!>Synthesize labeled RNA probes using commercially available kits. Best results are obtained by using lin-earized plasmid DNA as templates for in vitro transcription and by purifying the synthesized RNA probes usingcommercially available RNA clean-up kits. Store RNA probes in 50% formamide at -80°C. The probe can varyin concentration (the stock concentration is ~100 ng/µL, and the working concentration is ~0.5 ng/µL).

Ethanol (100%, 75%, and 50%)Glycine solution (2 mg/mL, prepared in PBT)<R>Hybridization buffer for Ciona (HB)<R>MEM-PFA




METHOD

Unless otherwise stated, all rinses, washes, and replacements should be performed in a volume of 1 mL, and incubationsshould be at room temperature. For Steps 14-27, the volume should be 0.4-0.5 mL (embryos are in the baskets inthe 48-well plates, which cannot accommodate more than this volume).

Fixation and Dehydration

1. Transfer dechorionated embryos to 1.5-mL tubes containing 0.5 mL of MEM-PFA. Let the embryossettle.

<!>Methanol (100%)Also prepare methanol:PBT solutions at 7:3, 1:1, and 3:7 (v:v).

<!>NaOH solution (1 N)<R>NBT-BCIP-PVA staining solution<R>PBS-EDTA stop solution<!>PBT containing 4% paraformaldehyde (PFA)<!>Permount<R>Phosphate-buffered saline (PBS) (1X) containing 0.05%-0.1% Tween 20 (PBT)<R>Phosphate-buffered saline (PBS) without CaCl

2or MgCl

2(1X) (ice cold for Step 3)

<R>Prehybridization buffer for Ciona (Pre-HB)<!>Proteinase K (2-4 µg/mL [for embryos] or 5-10 µg/mL [for larvae], prepared in PBT)

The optimal proteinase K concentration must be determined empirically with every new batch of proteinase K.

<R>Solution A for Ciona<!>Solution A is required only for an optional procedure described in the Troubleshooting section. If thisprocedure is performed, Solution A must be prepared alone and with 20 µg/mL RNase A.

<R>TMN (Tris-MgCl2-NaCl buffer)

<R>TNB (Tris-NaCl-blocking buffer)<R>TNT (Tris-NaCl-Tween buffer)<R>Wash buffer 1 for Ciona (WB1) (prewarmed to 55°C)<R>Wash buffer 2 for Ciona (WB2) (prewarmed to 55°C)

A 1:1 (v:v) solution of WB1:WB2 is also required.

<R>Wash buffer 3 for Ciona (WB3) (prewarmed to 55°C)<!>Xylene:ethanol solution (5:1, v:v)

Equipment

Aluminum sealing sheetCulture dish (48 well)Dissecting microscopeForcepsGlass slides and large coverslipsHeat block or polymerase chain reaction (PCR) machine set at 80°CHomemade baskets

To construct a homemade basket designed to fit into the well of a 48-well dish, cut off the bottom of a 0.5-mLtube where it becomes conical. Remove the central part of the lid, keeping the outer sleeve. Close the lid of thetube with nylon mesh (40 µm) between the lid and the tube. Make sure nylon mesh is not folded. Recut theborder of the lid/mesh to make the basket fit in the 48-well plate.

Hybridization oven preset to 55°CIncubator preset to 37°CMicrocentrifuge tubes (1.5 mL)PCR tubes (0.2 mL)RockerShaking platforms at room temperature and at 4°CStaining dish




Approximately 80 µL of settled embryos should be transferred to each tube, but this varies a great deal fromexperiment to experiment.

2. Replace the MEM-PFA with 1 mL of fresh MEM-PFA, and incubate the embryos for 14-18 h at 4°Cwith agitation.

3. Let embryos settle, and replace the solution with 1 mL of ice-cold 1X PBS.

4. Repeat Step 3 three times.

5. Add 0.5 mL of 100% ethanol to the embryos (in 1 mL of 1X PBS), and rock for 15-20 min at roomtemperature.

6. Remove 1 mL of solution (leaving 0.5 mL), add 0.75 mL of 50% ethanol and 0.25 mL of 100%ethanol, and rock for 15-20 min.

7. Replace 1 mL of solution with 1 mL of 50% ethanol, and rock for 15-20 min.

8. Replace 1 mL of solution with 0.75 mL of 75% ethanol and 0.25 mL 100% ethanol, and rock for15-20 min.

9. Replace 1 mL with 1 mL of 75% ethanol, and rock for 15-20 min.

10. Wash the embryos two to three times with 75% ethanol, rocking 15-20 min each time.Store the dehydrated embryos at -20°C.

Preparation of Dishes and Baskets

11. Use a forceps to distribute homemade baskets to separate wells of a 48-well dish. Dispense ~1 mLof 1 N NaOH to each well, and incubate overnight or longer.

12. Remove the 1 N NaOH. Rinse the baskets and wells successively with:

i. H2O (three times)

ii. 1X PBS (once)

iii. 75% ethanol (twice)

Rehydration and Permeabilization

13. Distribute the embryos in 0.5-mL baskets held on a 48-well plate containing 75% ethanol.Approximately 10-20 µL of settled embryos or ~10-500 embryos should be transferred to each basket, depend-ing on the experiment. Once the embryos are in the baskets, perform each buffer replacement by removing theold buffer from the outer side of the basket and adding fresh buffer gently to the inner side of the basket.

14. Rinse the embryos with 100% methanol by replacing the ethanol with 100% methanol, and thenrehydrate the embryos in each of the following solutions (15 to 20 min each):

i. Methanol:PBT (7:3, v:v)

ii. Methanol:PBT (1:1, v:v)

iii. Methanol:PBT (3:7, v:v)

iv. PBT (three times)

15. Replace the PBT with proteinase K solution, and incubate for 25 min at 37°C.

16. Replace the proteinase K solution with glycine solution, and rinse twice with PBT. Post-fix for 1 hat room temperature with PBT containing 4% PFA.

17. Rinse three times with PBT.

Hybridization

18. Replace PBT with pre-HB, and incubate for 5-10 min at room temperature.

19. Replace the pre-HB with 0.5 mL of HB, and incubate for 1 h at 55°C.




20. Dispense 50 µL of HB into 0.2-mL PCR tubes and add DIG-labeled RNA probe to a final probeconcentration of 0.1-1 ng/µL. Incubate at 80°C for 2-5 min.

21. Add 52.5 µL probe/HB mix to the embryos, seal the plate with an aluminum sealing sheet, andincubate at 55°C for 14-18 h.

Post-hybridization Washes

22. Carefully remove the probe/HB mix and keep for reuse. Replace with WB1 once, then wash in thefollowing sequence using prewarmed buffers:

i. WB1 for 20 min at 55°C (twice)

ii. WB1:WB2 (1:1, v:v) for 20 min at 55°C (twice)

iii. WB2 for 20 min at 55°C (twice)

iv. WB3 for 20 min at 55°C (twice)

Blocking and Antibody Incubation

23. Wash three times in TNT for 2-5 min each at room temperature.

24. Block with 0.5 mL of TNB for 1 h at room temperature.

25. Replace the TNB with antibody solution. Incubate for 2 h at room temperature and then for14-18 h at 4°C with shaking.

Antibody Washes and Staining

26. Replace the antibody solution with TNT, once as a quick rinse and then six times with a 20-minincubation each.

27. Wash with TMN three times for 10 min each. Transfer the embryos to 1.5-mL tubes after thesecond wash.

28. Replace the TMN with the NBT-BCIP-PVA staining solution, and transfer the embryos to a stainingdish. Keep the staining dish in the dark.

29. Regularly monitor color development using a dissecting microscope.The time required for staining depends on the probe; it can range from 30 min to several hours.

Dehydration and Mounting

30. Add 0.5-1 mL of PBS-EDTA stop solution to stop the staining reaction. Transfer the embryos to a1.5-mL tube and replace the PBS-EDTA several times.

31. Post-fix the embryos with PBT containing 4% PFA for 1 h.

32. Rinse the embryos with 1X PBS three times.

33. Add 0.5 mL of 100% ethanol to 1 mL of 1X PBS, and rock for 15-20 min.

34. Remove 1 mL (leaving 0.5 mL) of solution, add 0.75 mL of 50% ethanol and 0.25 mL of 100%ethanol, and rock for 15-20 min.


36. Replace 1 mL of solution with 0.75 mL of 75% ethanol and 0.25 mL of 100% ethanol, and rockfor 15-20 min.


38. Wash the embryos two to three times with 100% ethanol, rocking 15-20 min each time.

39. Wash quickly with xylene:ethanol (5:1, v:v), remove the xylene:ethanol mix, and mount inPermount.

40. Store the slide horizontally overnight until the resin has dried.




The results can vary greatly, depending on the target gene of interest and stage of the embryos.

See Troubleshooting for typical problems and solutions.

TROUBLESHOOTING

Problem: No staining is visible above background, even after a long period of staining.[Step 40]Solution: Consider the following:

• Some low levels of expression cannot be detected by in situ hybridization. Use conventionalreverse transcriptase (RT)-PCR to determine whether the gene of interest is expressed at thestage examined.

• Determine that the probe is of the expected size (using an RNA gel) and has been labeled withDIG (using a dot-blot assay).

• Run positive controls for (1) the experiment (i.e., use a probe for a gene that is known to beexpressed at the stage of interest) and (2) the specific probe being tested (i.e., by hybridizingtissues or embryos that are known to express the gene).

• Reduce the hybridization temperature to 42°C; this may also increase the background.

• Adjust the proteinase K treatment (excess or insufficient treatment reduces the sensitivity of theassay).

• Use moderate alkaline lysis to improve the penetration of the probe.

• Mount the embryos in 50%-80% glycerol instead of a xylene-based resin.

• Replace the 0.1% SDS in WB1 with 0.1% Tween 20; this may also increase the background.

Problem: There is high background.[Step 40]Solution: Consider the following:

• In Ciona embryos, maternal transcripts can be present throughout the embryo, which results indiffuse staining resembling background. To address this possibility, hybridize with a sense probe,which should not produce any staining.

• Increase the stringency of the washes by increasing the concentration of SDS (up to 1%) inbuffers WB1 and WB2. This might also reduce the sensitivity of the assay.

• Add an RNase treatment procedure during the post-hybridization washes. This reduces the sen-sitivity with most probes. To do this, proceed with the following steps after the wash with WB2(Step 22.iii): (1) Wash three times with solution A at room temperature; (2) incubate in solutionA supplemented with RNase A (20 µg/mL final concentration) for 20 min at 37°C; (3) wash twoto three times with WB3 at room temperature; and (4) continue with the WB3 wash as describedin Step 22.iv.

REFERENCES


Christiaen L, Wagner E, Shi W, Levine M. 2009b. Isolation of sea

squirt (Ciona) gametes, fertilization, dechorionation, and devel-opment. Cold Spring Harb Protoc (this issue). doi: 10.1101/pdb.prot5344.



Isolation of Individual Cells and Tissues from ElectroporatedSea Squirt (Ciona) Embryos by Fluorescence-Activated CellSorting (FACS)



INTRODUCTION

This protocol describes the dissociation of electroporated sea squirt (Ciona) embryos and the subse-quent isolation of individual cells, cell lineages, and tissues, using fluorescence-activated cell sorting(FACS). Sorted samples can be used for RNA extraction and reverse transcriptase polymerase chainreaction (RT-PCR) or microarray analysis. Alternatively, dissociated samples can be used for the subse-quent culturing and/or chemical treatment of isolated embryonic cells. This method permits the iden-tification of effector genes responsible for changes in cell shape, polarity, or motility in a given tissue.

RELATED INFORMATION

See The Sea Squirt Ciona intestinalis (Christiaen et al. 2009a) for an introduction to C. intestinalis asa model organism. For related methods, see Electroporation of Transgenic DNAs in the Sea SquirtCiona (Christiaen et al. 2009b).



Protocol

MATERIALS


Reagents

<R>Calcium- and magnesium-free artificial seawater (CMF-ASW)<!>Add antibiotics (e.g., 25-50 µg/mL of penicillin and streptomycin) for a long period of storage. For Steps 5and 8, supplement the CMF-ASW with 0.05% (w/v) embryo-tested bovine serum albumin (BSA), filter-sterilize,and chill on ice.

Ciona embryos

For FACS purification, embryos must have been electroporated with a construct producing green fluorescentprotein (GFP) expression in the cells of interest (see Electroporation of Transgenic DNAs in the Sea SquirtCiona [Christiaen et al. 2009b]). Because most constructs yield ectopic expression upon electroporation inCiona, counter-selection can be achieved by using another construct driving, for example, red fluorescentprotein (RFP) in the cells that would also express GFP but are not desired. In any case, negative controls mustbe prepared and used to define the parameters for FACS.

RNAlater (Ambion)RNAqueous-Micro Kit (Ambion)




METHOD

Collection and Dissociation

1. Add 1 mL of CMF-ASW to a borosilicate glass tube. Transfer the electroporated embryos (severalhundred to ~2000) to the tube and let them settle.

2. Remove the supernatant and add 1 mL of CMF-ASW. Transfer the embryos to 2-mL tubes. Rinseonce more with CMF-ASW.

3. Dilute the stock solution of trypsin to a final concentration of 0.1% in CMF-ASW.

4. Replace the supernatant covering the embryos with 1 mL of 0.1% trypsin and pipette with aPasteur pipette for 2-3 min.

5. Add 1 mL of ice-cold CMF-ASW supplemented with 0.05% BSA. Place the tubes on ice.

6. Filter once through a 40-µm cell strainer.

7. Centrifuge at 3000 rpm (~800g) for 2 min at 4°C for embryos older than the neurula stage or at1000 rpm for younger embryos.

8. Remove the supernatant and add 1 mL of ice-cold CMF-ASW-BSA. Pipette thoroughly but gentlyto resuspend the cells.

9. Repeat the centrifugation and rinse steps (Steps 7 and 8) at least once.

Cell Sorting and RNA Extraction

10. Bring the samples and collection tubes to the flow-cytometry facility.The detailed parameters and conditions used will vary with each facility and should be determined with the staffthat runs the facility.

11. Just before sorting, filter the cell suspension again on the cell-strainer cap of a 5-mL round-bottomtube used for sorting.

12. Collect cells in 0.5-mL of RNAlater buffer. Mix well and proceed in one of the following ways:

i. Store the cells at 4°C overnight.

ii. Freeze the cells at -80°C (indefinitely).

iii. Proceed immediately to RNA extraction using the RNAqueous-Micro Kit.See Troubleshooting.

<!>Trypsin (cell culture grade)Prepare a 2.5% stock solution in CMF-ASW, make 200-µL aliquots, and store at -20°C.

Equipment

Borosilicate glass tubes (12 × 75 mm)Rinse them overnight with distilled H

2O and dry before use (otherwise embryos tend to stick to the glass).

Cell strainers (40 µm)CentrifugeFlow cytometerIce bucket with iceLow-adhesion tubes (2 mL graduated)

Embryos do not seem to stick to the 2-mL tubes with catalog number 1480-2700 from USA Scientific.

Pasteur pipettes equipped with 2-mL bulbsAt least one Pasteur pipette per sample is required.

Polystyrene round-bottom tubes (5 mL) with cell-strainer cap (40 µm)

13_EVOv2_365-396_font.qxp:8.5 x 10.875 2/4/10 9:40 AM 4


TROUBLESHOOTING

Problem: There is contamination with cells from other tissues.[Step 12]Solution: The construct used for selection probably “leaks” in other tissues. Counter-selection with

another construct and different color must be used.

Problem: Very few events are actually sorted.[Step 12]Solution: This may arise from poor dissociation, which should be monitored visually to ensure thor-

ough dissociation. Larvae are more difficult to dissociate than embryos. Tissues representing a smallfraction of the embryos can be sorted, but more embryos will be required to obtain significantamounts of cells.

REFERENCES


Christiaen L, Wagner E, Shi W, Levine M. 2009b. Electroporation oftransgenic DNAs in the Sea Squirt Ciona. Cold Spring Harb Protoc(this issue). doi: 10.1101/pdb.prot5345.




Maize (Zeamays): A Model Organism for Basic and Applied ... · expansive amplification of...

Documents

Transcript of Maize (Zeamays): A Model Organism for Basic and Applied ... · expansive amplification of...