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Gene Expression Patterns 8 (2008) 237–247

Changes of c-tubulin expression and distribution in the zebrafish(Danio rerio) ovary, oocyte and embryo

Jianxiong Liu, Charles A. Lessman *

Department of Biology, The University of Memphis, 201 Life Science Building, Memphis, TN 38152-3560, USA

Received 19 June 2007; received in revised form 18 December 2007; accepted 20 December 2007Available online 31 December 2007

Abstract

The tubulin gene family is important for individual zebrafish development from the oocyte through to hatching. This involves oftenrapid, complex changes in the gametes and embryonic cells that are reflected in underlying gene expression changes. Tubulin dynamics,i.e., the interchange of polymeric and soluble forms in zebrafish oogenesis and embryogenesis, is important for microtubule (MT) cellularfunctions. Nevertheless, our understanding of how tubulin gene expression changes during zebrafish development is not clear. Previousdata showed that soluble a-tubulin and c-tubulin are associated with large molecular weight complexes (>2 MDa) which are reduced bythe blastula stage, with a concomitant decrease in soluble tubulin amount. Complexes (<2 MDa) then increased in the gastrula with anincrease in soluble tubulin. Microarray revealed similar patterns of tubulin gene product expression for zebrafish ovary and eggs whileboth differed from day 4 larva. In situ hybridization with c-tubulin oligonucleotide probes revealed diffuse label in oocytes, with amarked localization to the primordial blastodisc upon maturation. These findings, together with recent work on c-tubulin ring complexesin other species, suggest that c-tubulin (protein complexes) may be involved in regulating tubulin dynamics, thus is important for zebra-fish oogenesis and embryogenesis.� 2007 Elsevier B.V. All rights reserved.

Keywords: c-Tubulin; Microarray; In situ hybridization; Tubulin gene family; Ovary; Oocyte

Microtubules are hollow polymers of a-, b-tubulin thatshow GTP-dependent assembly dynamics and comprise acritical part of the eukaryotic cytoskeleton. The main micro-tubule organizing center (MTOC) of animal cells, the cen-trosome, is composed of a pair of centrioles surroundedby a fibrous pericentriolar material acting as a scaffold thatconcentrates both the microtubule nucleation machineryand its regulatory factors. In dividing cells, duplicated cen-trosomes separate at the onset of mitosis to establish thepoles of the mitotic spindle. Concomitantly, the pericentrio-lar material will progressively develop and then disassembleas cells exit mitosis (Dictenberg et al., 1998).

The mechanism of in vivo microtubule nucleation and itsrelationship to the various molecular components of thecentrosome remains unclear. Genetic and biochemical

1567-133X/$ - see front matter � 2007 Elsevier B.V. All rights reserved.

doi:10.1016/j.gep.2007.12.004

* Corresponding author. Tel.: +1 901 678 2963; fax: +1 901 678 4457.E-mail address: clessman@memphis.edu (C.A. Lessman).

approaches have led to the discovery of c-tubulin, a newsubtype of tubulin. The c-tubulin gene was first found inthe fungus Aspergillus nidulans as an extragenic suppressorof a mutation in the b-tubulin gene (Oakley and Oakley,1989). It has since been found almost ubiquitously ineukaryotic cells but at lower levels, when compared to a-and b-tubulin. Immunofluorescence and immunoelectronmicroscopy studies have revealed the presence of c-tubulinin every major MTOC that has been examined includingthe spindle pole bodies of Schizosaccharomyces pombe

and A. nidulans (Oakley et al., 1990; Horio et al., 1991),the centrosomes of Drosophila, Xenopus and mammaliancells (Stearns et al., 1991; Zheng et al., 1991; Debec et al.,1995; Sunkel et al., 1995), the MTOCs of differentiated cellssuch as neurons, chicken retinal epithelium and post-mito-tic ciliated cells (Baas and Joshi, 1992; Joshi and Besharse,1993) as well as the MTOCs of higher plants (Liu et al,1993).

238 J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247

Recently, the zebrafish (Danio rerio), a small tropicalfreshwater teleost, has emerged as an ideal model to studyvertebrate early development because of its high fecundity,short generation time and rapid development of the exter-nally fertilized and translucent embryos (Driever et al.,1994). MT arrays in zebrafish embryos are vital to manydevelopmental processes. Preliminary data from our labshowed that the MDa tubulin complexes extracted fromzebrafish ovaries contain a-, b- and c-tubulin as well asother proteins and these MDa tubulin complexes changedynamically during zebrafish early development (Liu andLessman, 2007). In addition, the soluble tubulin complexescontaining a-tubulin may have some relationship betweenc-tubulin and/or c-tubulin ring complexes as has beendemonstrated in Rana oocytes (Lessman and Kim, 2001)and in Xenopus oocytes (Zheng et al., 1995). Using immu-nofluorescence microscopy technique, we revealed thedynamic distribution of c-tubulin protein during zebrafishearly development (our unpublished data). Furthermore,futile cycle (fue) mutant zebrafish embryos show apparentlynormal centrosomes during cleavage, as deduced by c-tubulin staining, yet produce anuclear blastomeres priorto arrest (Dekens et al., 2003). This intriguing paper sug-gests that centrosome duplication, aster formation and cen-trosome assembly as well as cytokinesis may occur duringcleavage without spindle assembly. Thus, the dynamic dis-tribution of c-tubulin mRNA, and the physiological rolesc-tubulin play during zebrafish oogenesis and embryogene-sis deserve further investigation.

Over the past several years, the new technology of DNAmicroarray has attracted tremendous interest among biolo-gists. This technology promises to monitor the whole gen-ome on a single chip so that researchers can have a betterpicture of the interactions among thousands of genessimultaneously. There are two major applications for theDNA microarray technology: (1) identification of sequence(gene/gene mutation); and (2) determination of expressionlevel (abundance) of genes (Shi, 1998–2002). In this study,to test the expression levels of tubulin-related genes duringzebrafish early development, total RNAs from zebrafishwhole ovary, mature oocytes (eggs), and day 4 larvae wereisolated and analyzed using GeneChip and microarraytechniques. The abundance of each tubulin-related geneproduct was compared among these three stages of zebra-fish development. Thus this study will provide some funda-mental data for the future study of zebrafish, anincreasingly important model organism.

In situ hybridization is a widely accepted method tovisualize mRNA expression in tissues and cells. In thisstudy, to determine the dynamic distribution of c-tubulinmRNA during zebrafish oogenesis and embryogenesis, afluorescein-labeled DNA probe with a complimentarysequence to zebrafish c-tubulin mRNA was hybridized tothe fixed tissues of zebrafish ovary, oocyte, and differentstaged embryos. The resultant signal, based on alkalinephosphatase conjugated anti-fluoroscein antibody, wasimaged with a microscope. Our data showed that c-tubulin

mRNA was actively expressed during zebrafish early devel-opment, and its cellular distribution changed dynamicallyfrom ovarian oocytes to late embryos (i.e., from corticallyarrayed in oocytes to localized in the blastodisc and blasto-meres of mature eggs and embryos, respectively).

1. Results

1.1. Microarray data

To test the expression levels of a variety of tubulin-relatedgenes, especially c-tubulin, in zebrafish early development,we isolated total RNA from zebrafish whole ovary, matureoocytes (eggs), and day 4 larvae, respectively, using eitherTrizol or Stat-60 method. The RNAs were then analyzedwith GeneChip using microarray. The abundance of eachtubulin-related gene among those three stages was comparedand listed as signal units in Table 2. The data showed thatthere was a broad range of tubulins and associated mRNAsexpressed in whole ovary, eggs and day 4 larvae, includingdifferent isoforms of a- and b-tubulins.

During the process of oocyte maturation, c-tubulininteracting protein (yeast SPC98 homolog) mRNAincreased nearly 2.5-fold from oocytes to matured egg,while c-tubulin 1 and c-tubulin complex protein 2remained relatively constant. In contrast to the decreaseof a 1 mRNA (almost 2-fold), b 2 and b 5 mRNAs wereall shown to be increased at least 2-fold.

The data also revealed that ovary and eggs had similarabundance of tubulin gene product expression while differ-ing from day 4 larvae. Although c-tubulin 1 and c-tubulincomplex protein 2 were expressed in all stages: wholeovary, eggs and day 4 larvae, the abundance decreased sig-nificantly in day 4 larvae. In addition, c-tubulin interactingprotein (yeast SPC98 homolog) mRNA was present in bothovary and eggs but it was undetectable in day 4 larvae.Meanwhile, the amount of a 4 like tubulin and b-tubulincofactor C decreased dramatically during embryogenesis.In contrast, a 1 and 8; b, b 2 and b 5 were all found tobe increased from oocytes to larvae.

1.2. In situ hybridization assay

Since mRNAs of c-tubulin and its associated protein weredetected in microarray assay, and it is well-known that c-tubulin plays a variety of important roles in zebrafish embry-onic development (Table 1), the localization of c-tubulinmRNA during zebrafish early embryogenesis was investi-gated. To visualize the c-tubulin gene products in zebrafishearly development, the c-tubulin mRNAs were detected spe-cifically by in situ hybridization assays. Fig. 1 illustrated thelocalization of c-tubulin gene expression, i.e., mRNAs, indifferent stages of zebrafish oogenesis and embryogenesis.The data showed that the c-tubulin mRNAs were diffuselylocalized along the cortex of the fully-grown oocytes(Fig. 1A and inset). In contrast, the c-tubulin mRNAs wereaggregated in the animal pole primordial blastodisc area in

Table 1Functions of c-tubulin in somatic, embryonic and germ-cells

References Organism

General somatic cells

Component of pericentriolar material Stearns et al. (1991) Xenopus, yeast, maize, diatoms

Microtubule (MT) nucleating material with cell cycle-specific variation Zheng et al. (1991) Drosophila

Cell viability Oakley et al. (1990) Aspergillus nidulans

Stearns et al. (1991) Xenopus, yeast, maize, diatomsHorio et al. (1991) Schizosaccharomyces pombe

MT assembly in vivo Oakley et al. (1990) Aspergillus nidulans

Horio et al. (1991) Schizosaccharomyces pombe

Joshi et al. (1992) Mammalian cells

Self-assembly into novel tubular structures Shu and Joshi (1995) Mammalian cells

Regulation in MT dynamics and organization Paluh et al. (2000) Schizosaccharomyces pombe

Anchoring the medial cytokinetic actin ring Pardo and Nurse, 2003 Schizosaccharomyces pombe

Spindle orientation, cleavage site specification Venkatram et al. (2004) Schizosaccharomyces pombe

Coordination of mitotic events and checkpoint control Prigozhina et al. (2004) Aspergillus nidulans

Organization of kinetochore and interpolar MT Strome et al., 2001 Caenorhabditis elegans

MT nucleation in eukaryotic flagellum McKean et al. 2003 Trypanosoma brucei

Differentially upregulation to the synthesis of tubulin Zhou et al. (2002) Mammalian cells

Formation and maintenance of basal bodies Shang et al. (2002) Tetrahymena thermophila

Oogenesis/oocyte

Bicoid mRNA localization Schnorrer et al. (2002) Drosophila

Oocyte differentiation and female germ-cell proliferation Tavosanis and Gonzalez (2003) Drosophila

Establishment or maintenance of A–V axis Gard (1993) Xenopus

Spermiogenesis

Maintenance of juxtaposed spindles in spermatocytes Barbosa et al. (2003) Drosophila

Embryogenesis

Reconstitution of zygotic centrosome Shin and Kim (2003) Calf

Establishment of spindle bipolarity Prigozhina et al. (2004) Aspergillus nidulans

Anucleate cleavage in fue mutant Dekens et al. (2003) Danio rerio

Fig. 1. In situ hybridization (probed with FITC-oligo c-tubulin probe) of different stages of zebrafish oocytes (A and G are fully grown, immature oocytes;B and H are mature oocytes or eggs) and embryos (C and I are 32 cell; D and J are 256 cell; E and K are �30% epiboly gastrula; F and L are pharyngula).The top panels are ‘‘with probe” treatments (A–F) and the lower panels are corresponding ‘‘without probe” treatments (G–L). The primordial blastodisc(b) that forms at the animal pole of the egg during oocyte meiotic maturation labels intensely with the probe (B). Inset: immature oocyte probed afterhemisection showing cortical label. Anti-FITC second antibody conjugated to alkaline phosphatase and the substrate DAB were used to develop color.The specimens were dehydrated in 100% MeOH and placed in clearing media benzyl benzoate:benzyl alcohol (2:1) prior to mounting on slides. Scale barequals 250 lm.

J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247 239

240 J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247

DHP-treated mature oocytes (Fig. 1B). In zebrafish cleavageembryos, the c-tubulin mRNAs were found to be localizedprimarily in blastomeres and not in yolk cells (Fig. 1C andD). The c-tubulin mRNAs were barely detectable in gastrulastage embryos (Fig. 1E). But in late pharyngula stageembryos, the c-tubulin mRNAs were enriched along zebra-fish anterior–posterior axis, from head to tail (Fig. 1F).

We treated some immature oocytes with both actinomy-cin D (transcription inhibitor) and DHP. We found thatc-tubulin mRNAs were still aggregated in the animal poleprimordial blastodisc area of the treated DHP-maturedoocytes (Fig. 2H). These data suggested that the bulk ofthe c-tubulin mRNAs were synthesized and stored mater-nally before maturation. In addition, the translocation ofc-tubulin mRNAs from cortex of immature oocytes to pri-mordial blastodisc area in DHP-treated mature oocyteswas not dependent on an intact microtubule network, sincec-tubulin mRNAs could still be detected predominately inthe animal pole primordial blastodisc in demecolcine-treatedDHP-matured oocytes (microtubule de-polymerizingreagent, Fig. 2C), similar to DHP-treatment alone (Fig. 2Aand F). If the oocytes were treated with RNase prior tohybridization, all the signals were diminished (Fig. 2J). Inaddition, all the oocytes incubated in the absence of probehad signal approaching background (Fig. 1: lower panel;Fig. 2B, D, G and I). All these results demonstrated the integ-rity of this in situ hybridization assay.

2. Discussion

2.1. c-Tubulin and microtubules

Microtubule (MT) nucleation and tubulin dynamics areimportant in a variety of cell functions during oogenesis

Fig. 2. c-Tubulin in situ hybridization of 17 a 20 b dihydroxyprogesterone (DHin the animal pole primordial blastodisc (b). (A, D, G, J and M) incubatedpreincubated with RNase prior to hybridization. (G and H) preincubated withK preincubated with actinomycin D (1 lg/ml) and DHP prior to fixation and h(b) formation at the animal pole after DHP incubation. (A, B and C) are ani

and embryogenesis. In the African frog Xenopus laevis,MTs contribute to the establishment and maintenance ofthe animal–vegetal polarity (Wylie et al., 1985; Gard,1991, 1995, 1999; Gard et al., 1995; Palecek and Ubbels,1997). In the fertilized egg, MTs of the sperm aster andthe cortical MT array are involved in pronuclear migrationand the specification of the dorsal–ventral axis (Elinsonand Rowning, 1988; Schroeder and Gard, 1992; Larabellet al., 1996). Therefore, MT dynamics and organizationmust be regulated both temporally and spatially. The rela-tively large size of Xenopus eggs and the rapid assembly ofthe sperm aster pose unique problems for MT growth andorganization, suggesting Xenopus eggs and early embryosmight contain novel factors that regulate MT dynamicsand organization during early development (Becker andGard, 2000). In addition, members of the tubulin proteinfamily are present in oocytes and eggs of both Rana pipiens

(Lessman, 1993; Wang and Lessman, 1997, 2002) and X.

laevis (Zhou et al., 1991) in soluble MDa complexes.Similarly, MT arrays are vital to many developmental

processes in zebrafish embryos. Besides their obvious rolein spindle formation, MTs are required for epiboly (Sol-nica-Krezel and Driever, 1994; Strahle and Jesuthasan,1993), furrow formation (Pelegri et al., 1999) and the cohe-sion of post-cytokinesis blastomeres (Jesuthasan, 1998).Dorsal maternal determinants in teleost embryos arethought to be present in the vegetal mass of the yolk cellsoon after fertilization (Mizuno et al., 1997). Recent workimplicates microtubules in the transport of dorsal determi-nants from the vegetal pole of the zygote towards the blas-todisc. As transport lines for regulatory substances andmaternal mRNAs, MTs are also required for axisdetermination and establishment of embryonic polarity inzebrafish (Jesuthasan and Strahle, 1997). This microtu-

P 1 lg/ml) matured oocytes. c-Tubulin message was found predominatelywith probe; (B, E, H, K, and N) incubated without probe. (M and N)demecolcine (1 lg/ml) and DHP prior to fixation and hybridization. J andybridization. (C, F, I, L, and O) are live cells showing primordial blastodiscmal pole views, while all others are side views. Scale bar equals 200 lm.

J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247 241

bule-dependent transport might play a key role in theestablishment of embryonic polarity in the zebrafish (Jesu-thasan and Strahle, 1997). Thus the study of MT functionin zebrafish embryos is consequently important for under-standing the molecular mechanisms underlying numerousdevelopmental processes.

Microtubule nucleation in vivo is thought to involve theinteraction of tubulin subunits (a-, and/or b-tubulin) withcentrosome components. It was proposed that c-tubulininteracts with MTs via a physical interaction with b-tubulinsubunit of the tubulin heterodimer (Mandelkow and Man-delkow, 1994; Oakley, 1994). In addition, evidence gainedfrom the binding of GTP-analogs covalently attached tofluorescent beads suggests that b-tubulin is the terminalsubunit at the plus end of the MT; however, it was alsoproposed that the minus end may have an alternative struc-ture, perhaps consisting of b:c heterodimer (Mitchison,1993). Initiation of new microtubules in vivo requires c-tubulin, organized as an oligomer within the 2.2 MDa cTuRC, of higher eukaryotes. At 2.7 A crystal structure res-olution of the human protein, c-tubulin was reported to bebound to GTP-gammaS (a non-hydrolysable GTP ana-logue). A curved conformation for c-tubulin-GTP-gam-maS is found to be similar to that seen for GDP-bound,unpolymerized a-, b-tubulin (Mitchison, 1993). Thus thislateral contact between a- and b-tubulins in the microtu-bule probably forms the basis for c-tubulin oligomerizationwithin the cTuRC. Laterally associated c-tubulins in thecTuRC might promote microtubule nucleation by provid-ing a template that enhances the intrinsically weak lateralinteraction between a-, b-tubulin heterodimers. Our micro-array data showed that there was a broad range of tubulinmRNAs expressed in whole ovary, eggs and day 4 larvae,including different isoforms of a- and b-tubulins. Table 2demonstrated that after oocyte maturation, yeast SPC98homolog, b 2 and 5 increased up to 2-fold while c-tubulin1 and c-tubulin complex protein 2 remained relativelyconstant.

Table 2 also demonstrated that in day 4 larvae, theabundance of c-tubulin 1 and c-tubulin complex protein2 decreased significantly. Furthermore, c-tubulin interact-ing protein (yeast SPC98 homolog) mRNA was undetect-able in the larval stage. These data strongly agreed withour previous hypothesis (Liu and Lessman, 2007): ovariantubulin complexes were sequestered and maintained in atemporary ‘‘oligomeric” state and appear to be relativelystable storage forms of soluble tubulins, which subse-quently may play important roles in microtubule dynamicsand centrosome formation in zebrafish oogenesis andembryogenesis. The whole process of maintenance, sub-division, relocalization of these ovarian tubulin complexesmay be regulated by c-tubulin and/or other associatedproteins. In contrast, a broad range of isoforms of a- andb-tubulin mRNAs were greatly increased in later embryo-genesis, such as a1, a8 and b5. These data suggest thatthe oocytes, eggs, embryos and larvae may have differentdemands for microtubule nucleation, or even further, they

adopt different regulating systems at different stages duringzebrafish early development.

2.2. The physiological role of c-tubulin in animal cells

Evidence accumulated to date has shown that c-tubulin,the evolutionarily conserved and ubiquitously expressedprotein, is very crucial for microtubule (MTs) nucleation(Oakley et al., 1990; Stearns and Kirscher, 1994; Zhenget al., 1991) and other cellular functions (Table 1). Thecurrently favored model is that centrosome-associatedc-TuRCs serves as templates to nucleate the polymeriza-tion of a-, b-tubulin dimers into microtubules. This conclu-sion is based on experimental manipulations which includeantibody blocking and over-expression in mammalian cells(Joshi et al., 1992; Julian et al., 1993; Ahmad et al., 1994;Shu and Joshi, 1995); immunodepletion and biochemicalmanipulation of in vitro reconstituted centrosomes fromXenopus egg extracts (Felix et al., 1994; Stearns and Kir-scher, 1994); automated electron tomography (Moritzet al., 1995); biochemical purification and functional assays(Zheng et al., 1995); and genetic analysis in S. pombe

(Horio et al., 1991), A. nidulans (Oakley et al., 1990) andDrosophila (Sunkel et al., 1995). In Aspergillus, mixed het-erokaryons are viable but those spores that contain thetransformed nuclei and therefore lack the c-tubulin genecontain only few cytoplasmic microtubules and do notdivide. In vertebrate cells, injection of a polyclonal anti-body directed against a synthetic peptide of the c-tubulinsequence prevents microtubule regrowth after nocodazoleor cold-induced depolymerization, and mitotic spindle for-mation, and mitotic spindle formation (Joshi et al., 1992;Felix et al., 1994).

In addition, c-tubulin is required for several processes atdifferent stages of germ-cell development and oogenesis inDrosophila, including oocyte determination, differentiation,and female germ-cell proliferation (Tavosanis andGonzalez, 2003). Furthermore, the function performed byc-tubulin seems to be highly conserved across different spe-cies since the expression of human c-tubulin restores viabil-ity to S. pombe cells mutant for c-tubulin (Horio andOakley, 1994).

c-Tubulin has an important function in the coordinationof late mitotic events and that it has a microtubule-inde-pendent role in establishing or maintaining a mitotic check-point block (Prigozhina et al., 2004), such as chromosomaldisjunction, anaphase A, anaphase B, and chromosomaldecondensation. For mitosis to be completed successfully,multiple mitotic events or processes must be coordinatedcorrectly. Vardy and Toda (2000) proposed that check-point pathways-dependent on functional c-tubulin com-plexes exist and are evolutionarily conserved even tovertebrates. These data were recently extended and clarified(Vardy et al., 2002).

During Drosophila oogenesis, it was found that c-tubu-lin37C and c-tubulin ring complex protein 75 are essentialfor bicoid RNA (one of Drosophila embryonic determi-

Table 2Microarray data for tubulin-related genes and mRNAs in wild-type zebrafish ovary, mature egg and day 4 larva, presented as relative abundance

Ovary Egg Day 4 larva Target description Probe Accession No.

Signal from sample

609 783 477 Tubulin, c 1 Dr.11201.1 gb:BC045486.1443 370 121 c-Tubulin complex protein 2 Dr.3637.1 gb:BG302753241 620 c-Tubulin interacting protein (yeast SPC98 homolog) Dr.15107.1 gb:BM182344

2310 2207 14460 a-Tubulin Dr.11310.4 gb:BQ6150909012 Tubulin, a 1 Dr.11310.1 gb:BC042319.1

770 371 4109 Tubulin, a 1* Dr.11310.2 gb:AF029250.11329 514 2956 Tubulin, a 1* Dr.11310.2 gb:AF029250.1

6633 Tubulin, a 1** Dr.11310.3 gb:BG3062122386 Tubulin, a 1** Dr.11310.3 gb:BG306212

1440 1556 Tubulin a-1 chain, brain-specific Dr.25680.1 gb:AL7161502267 3018 41077 Tubulin, a 1 Dr.664.3 gb:BI326418179 199 307 Tubulin, a 2 isoform 1 Dr.25456.1 gb:BI673876

1866 Tubulin, a 2 Dr.26381.1 gb:BI86629912510 15701 214 Tubulin, a 4 like Dr.23436.1 gb:BC046889.111188 13743 26200 Tubulin, a 8 like 4 Dr.664.1 gb:BC045847.14407 3631 6783 Tubulin, a 8 like Dr.20010.2 gb:AW115602

1028 Tubulin, a 8 like 3 Dr.20214.1 gb:AI7937085065 5227 14362 Tubulin b*** Dr.5605.3 gb:AI4772424897 5424 13503 Tubulin b*** Dr.5605.3 gb:AI477242554 431 14108 Tubulin b Dr.5605.4 gb:BQ26391792 135 489 b-Tubulin Dr.6173.1 gb:AW07744454 78 3155 Tubulin b-2 chain Dr.24902.1 gb:AL717250

7670 10441 11886 Tubulin, b, 2 Dr.5605.1 gb:AI3971048078 11396 14653 Tubulin, b, 2 Dr.5605.2 gb:BM141612105 324 1587 Tubulin, b 5 Dr.4416.1 gb:AF528096.1

9885 10364 1220 Tubulin b-5 chain Dr.24758.1 gb:BQ0784427641 Tubulin b-5 chain Dr.24758.2 gb:BQ078442

1913 1978 2445 Tubulin cofactor a b-tubulin cofactor C; Dr.1163.1 gb:BC046032.11873 1852 485 Tubulin-specific chaperone c Dr.16676.1 gb:BI980285

Data presented as normalized signal intensity; items with a similar number of asterisks (*) have the same accession number and probe set (i.e., they areinternal replicates on the microarray).

242 J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247

nants required for anterior–posterior polarity) localization(Schnorrer et al., 2002). Mutations in these genes specifi-cally affect bicoid RNA localization, whereas other micro-tubule-dependent processes during oogenesis are notimpaired.

In contrast, other lines of evidence suggest that c-tubu-lin is not absolutely required to seed microtubule growth incells. Null c-tubulin mutations in Saccharomyces cerevisiae,S. pombe and Drosophila impair but do not completelyblock the assembly of mitotic spindles (Horio et al., 1991;Sobel and Snyder, 1995; Sunkel et al., 1995; Wilson andBorisy, 1998). And c-tubulin is not absolutely requiredfor microtubule nucleation in Caenorhabditis elegans butis required for the normal organization and function ofkinetochore and interpolar microtubules (Strome et al.,2001). In summary, the physiological role of c-tubulin inanimal cells is more complex than initially assumed.

2.3. mRNA localization in zebrafish oocytes and embryos

The zebrafish ovary consists of a jumbled array ofoocytes ranging through four different stages of oogenesis(Selman et al., 1993). At stage IV the oocyte undergo mat-uration, marked by both the migration of GV to the futureanimal pole of the embryo and the break down of the

nuclear envelope, which is one of the first definitive mor-phological signs of polarity in the oocyte/egg and futureembryo. Patterns of mRNA localization are analyzed onzebrafish ovary sections by performing in situ hybridiza-tions using probes to several maternally expressed zebrafishgenes (Howley and Ho, 2000). They found that the zebra-fish egg is polarized along the animal/vegetal axis, as seenthrough dynamic mRNA localization patterns. Many tran-scripts become localized to the animal pole during oogene-sis. Some mRNAs (e.g. b-catenin) localize only uponactivation (fertilization) and others (e.g. vasa and zDazl)redistribute themselves through cytoplasmic streamingmovements, provide alternative later pathways for mRNAsto be distributed in the embryo. In this study, by perform-ing in situ hybridizations using probes specific for c-tubu-lin, we found that c-tubulin mRNA was diffuselydistributed in oocytes. But when the oocytes were maturedwith DHP, c-tubulin mRNA relocated and accumulated tothe primordial blastodisc area. Thus according to the tran-script relocation types indicated by Howley and Ho (2000),c-tubulin mRNA relocation represented a new type com-bining the cortical class (e.g. vasa) with the animal poleclass (e.g. cyclin B). During the 6-h maturation process,c-tubulin mRNAs were found to be accumulated as‘‘patches” and moved together toward animal pole along

J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247 243

cortex surface. They were finally cleared from vegetal poleafter 5-h incubation and reached primordial blastodisc areaafter 6-h treatment (our unpublished data). The exactmechanism by which the c-tubulin mRNA is translocatedto the primordial blastodisc during oocyte maturation isunclear. Since actinomycin D failed to block the relocation,it seems unlikely to be due to new transcription, in additionthis drug was previously reported to have no effect onDHP-induced oocyte maturation (Kohli et al., 2005). Dem-ecolcine did not block the relocalization suggesting thatmicrotubules were not involved in the mechanism. Previouswork indicated that other microtubule drugs, includingtaxol and nocodazole, did not inhibit oocyte maturationinduced by DHP (Kondo et al., 2001). While testing cyto-chalasins for possible effects seems a logical next step, itwas demonstrated that cytochalasin B itself induces a formof oocyte maturation in the zebrafish oocyte by elicitingcyclin B mRNA translation, thus confounding this test(Kondo et al., 2001). Presumably ooplasmic movementsassociated with oocyte maturation, including germinal ves-icle migration (GVM), germinal vesicle dissolution (GVD)and primordial blastodisc formation, reported earlier(Lessman et al, 2007) contribute to the rearrangement ofthe mRNA in the zebrafish oocyte–egg transition. Theexact dynamics and rate of the primordial blastodisc for-mation during zebrafish oocyte maturation has not beencarefully studied to date. Thus, variations in primordialblastodisc size or extent, including the in situ label, maybe a function of the particular point at which the prepara-tion was made. The possibility, that demecolcine may affectthe final size or extent of the primordial blastodisc, cannotbe ruled out at this point. It may be that certain mRNAs,including c-tubulin mRNA and that for axial determinates,are transported en masse into the primordial blastodiscarea and that these RNAs are in fact linked somehow.

2.4. Regulation of c-tubulin synthesis

Shu and Joshi (1995) have found that c-tubulin, whenoverexpressed, accumulates throughout the cytoplasmand forms numerous ectopic microtubule nucleation sitesin mammalian cells. Subsequently it was shown that over-expression of c-tubulin differentially up-regulates the syn-thesis of a- and b-tubulins in mammalian cells (Zhouet al., 2002). Surprisingly, despite a dramatic increase inthe level of c-tubulin protein in transfected cells, there isno obvious alteration in the level of endogenous c-tubulinmRNA, suggesting that synthesis of c-tubulin mightemploy a regulatory mechanism other than the autoregu-latory pathway shared by a- and b-tubulins. Interestingly,the transfected cells fail to form normal bipolar mitoticspindle during mitosis. Numerous microtubules occur inthe cytoplasm intermingled with the condensed chromo-somes. These data indicated that the number of microtu-bule nucleation sites, or even c-tubulin itself, might playan important role in the regulation of tubulin synthesisas well as cell cycle progression. The co-localization of

c-tubulin and phosphorylated MAP kinase with microtu-bule assembly in both control and taxol-treated pigoocytes suggests that chromosomes are always co-local-ized with microtubules and that emanation of microtu-bules from chromosomes may be regulated/directed bymicrotubule-organizing material including c-tubulin andphosphorylated MAP kinase in pig oocytes (Sun et al.,2001).

Previous data from our lab demonstrated that c-tubulinand/or its associated proteins may play some roles in regu-lating the changes of soluble tubulin complexes during zeb-rafish oogenesis and embryogenesis (Liu and Lessman,2007). Both soluble and insoluble c-tubulin decreased inamount until mid-blastula transition stage, and then re-increased after gastrula stage. In this study, the cellular dis-tribution of c-tubulin mRNA changed dynamically duringzebrafish oogenesis, from cell cortex to primordial blasto-disc area as oocytes matured. And during later embryogen-esis, both the relative abundance and distribution of c-tubulin mRNA changed dynamically. It was undetectableat the gastrula stage but it increased again by the pharyn-gula stage. c-Tubulin mRNA accumulated in and was lim-ited to blastomere cells during zebrafish cleavage stage.This apparent change in abundance may be due to deple-tion of maternal mRNA stores during cleavage and replen-ishment by zygotic gene transcription after the midblastulatransition. After the gastrula stage, c-tubulin mRNA wasfound to be enriched along the anterior–posterior axis,from head to tail. Based on the above data, c-tubulinand its associated protein complexes turn out to be a strongcandidate in regulating tubulin dynamics during zebrafishoogenesis and embryogenesis.

2.5. Isoforms of c-tubulin

Current data showed that c-tubulin is differentiallyrequired for the maintenance of different MTOCs (Shanget al., 2002). During gametogenesis and early developmentin Drosophila, two c-tubulin isoforms, gTub37CD andgTub23C, are differentially expressed. Both isoformscontain structural features of c-tubulins essential forlocalization to centrosomes (Wilson et al., 1997). Posttrans-lational modifications of c-tubulin might have an impor-tant role in the regulation of microtubule nucleation(Sulimenko et al., 2002). Interestingly, in various evolu-tionarily diverse organisms such as Arabidopsis thaliana

(Liu et al., 1994), Zea mays (Lopez et al., 1995), Physarumpolycephalum (Lajoie-Mazene et al., 1996), Euplotes crassue

(Tan and Heckmann, 1998) and Paramecium tetraurelia

(Ruiz et al., 1999), two or three isoforms were identifiedfor c-tubulin. Among vertebrates, only human (Wiseet al., 2000) and mouse (Yuba-Kubo et al., 2005) havetwo closely related c-tubulin genes. However, neither theirphylogenetic evolution nor the isoform-specific expression/function has been well characterized. There are two iso-forms of c-tubulin present in Drosophila and they are func-tionally equivalent during female germ-cell development

244 J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247

and oogenesis (Tavosanis and Gonzalez, 2003). The expres-sion of c-tubulin isoforms in Drosophila oocytes is undertight developmental regulation. The two c-tubulins foundin Drosophila show slightly more than 80% homology,which is much less than that shown between the humanand Xenopus c-tubulin (98%), and about as much asbetween any of them and their Xenopus homolog. This rel-atively large divergence between the two Drosophila iso-forms may indicate functional divergence (Oakley, 1994).It suggests that, superimposed on their basic role in micro-tubule polymerization, they may have acquired functionswhich are specific for the microtubule organization require-ments of the developmental stages at which they areexpressed. This point is substantiated further by the verydifferent phenotypes displayed by mutant alleles of thesetwo genes (Tavosanis et al., 1997). The expression of 37Cisoform is restricted to ovaries and early embryos, andmutation in this gene has no major consequence other thanfemale sterility. In addition, c-tubulin 37C and c-tubulinring complex protein 75 are essential for bicoid RNA local-ization during Drosophila oogenesis (Schnorrer et al.,2002). In contrast, the 23C isoform is ubiquitouslyexpressed and mutations in this gene impairs viability (Sun-kel et al., 1995). Aside from these findings, the physiologi-cal relevance of the existence of other c-tubulin isoforms inother species is far from understood. These data providerationales for determining whether c-tubulin is requireddifferentially in zebrafish oogenesis and embryogenesis.With our microarray and in situ hybridization data, wecan conclude that the synthesis and distribution of c-tubu-lin mRNA changed dynamically during zebrafish earlydevelopment. Whether there are several isoforms of c-tubulin in zebrafish oocytes and embryos, as reported inArabidopsis (Liu et al., 1994), Drosophila (Wilson et al.,1997), human (Wise et al., 2000) and mouse (Yuba-Kuboet al., 2005); and whether c-tubulin isoforms are requireddifferentially in zebrafish early development is not known.In addition, the regulating mechanisms for c-tubulinmRNA synthesis and maintenance in oocytes, and its redis-tribution during activation, fertilization and later embryo-genesis need further investigation.

3. Experimental procedures

3.1. Zebrafish

Wild-type zebrafish were obtained from a local distributor and housedin a dedicated, temperature (28 �C) and photoperiod (14L) controlled fishroom in racks of food-quality plastic containers (6 and 12L) with flow-through dechlorinated water. Fish were fed 1–2 times daily a diet of Tetr-amin and dried brine shrimp flakes.

3.2. Whole ovary collection

Donor female zebrafish were decapitated, the spinal cord pithed, thewhole ovary was removed, washed with Cortland’s solution (NaCl7.25 g, CaCl2–2H2O 0.23 g, KCl 0.38 g, MgSO4–7H2O 0.23 g, NaHCO3

1.0 g, Penicillin 30 mg and Streptomycin 50 mg in 1000 ml double distilledwater) and pooled into a 1.5 ml microcentrifuge tube.

3.3. Fully-grown oocytes collection

After the whole ovary was removed and placed in a Petri dish halffilled with fresh Cortland’s solution, the opaque fully-grown oocytes werethen separated from the others and pooled into a 1.5 ml microcentrifugetube.

3.4. Mature oocyte (egg) collection

The opaque fully-grown oocytes were separated from the others fol-lowing the above procedure and pooled into a Petri dish half filled withfresh Cortland’s solution with 17 a–20 b-dihydroxyprogesterone (DHP;final concentration 1 lg/ml) to induce meiotic maturation (Lessmanet al., 2007). After the oocytes were incubated at 28 �C for 6 h, the trans-lucent, mature oocytes (eggs) were collected and pooled into a 1.5 mlmicrocentrifuge tube. For drug treatment, the opaque fully-grown oocyteswere separated and pre-incubated with 10 ml Cortland’s solution contain-ing both DHP and different drugs: demecolcine, or actinomycin D dis-solved in steroid vehicle (with final working concentration of 1 lg/ml)prior to fixation and hybridization or RNase added after fixation and per-meabilization. The same concentration of steroid vehicle (0.1%) was usedas a negative control.

3.5. Embryo collection

Male and female fish were maintained in separated tanks, and com-bined to collect embryos just prior to mating. Crosses were set up in theafternoon using 3 females and 4–5 males per breeding tank. After mating,males and females were separated again and allowed to rest for at least oneweek before the next cross. Embryos were collected next morning andwere washed to remove any debris (Grinblat et al., 1999). Standard fishwater (Westerfield, 1995) supplemented with methylene blue (final concen-tration 2 ppm to reduce fungal contamination) was used for the washesand maintained at 28 �C. Washed embryos were grouped according tothe zebrafish stage tables (Westerfield, 1995) and collected into separate1.5 ml microcentrifuge tubes.

3.6. Total RNA extraction

3.6.1. TRIzol method

Tissue samples (50–100 mg) were homogenized in 1 ml of TRIzolreagent using a glass–Teflon homogenizer. The sample volume did notexceed 10% of the volume of TRIzol reagent used for homogenization.The homogenized sample was incubated at room temperature for 5 minto permit the complete dissociation of nucleoprotein complexes. Chloro-form (0.2 ml) was then added per 1 ml of TRIzol reagent. The sampletubes were capped securely, shaken vigorously by hand for 15 s and incu-bated at room temperature for 2–3 min. The samples were then centri-fuged at 10,000g for 15 min at 2 �C. Following centrifugation, thecolorless upper aqueous phase (containing RNA) was transferred to afresh tube, and the lower red phenol–chloroform phase was saved forfuture DNA and protein isolation. The total RNA was precipitated fromthe aqueous phase by mixing with isopropyl alcohol (0.5 ml of isopropylalcohol per 1 ml of TRIzol reagent was used for the initial homogeniza-tion). The samples were incubated at room temperature for 10 min andcentrifuged at 10,000g for 10 min at 2 �C. The RNA precipitate waswashed once with 75% ethanol and air dried at room temperature. Thenthe RNA was dissolved into RNase-free water by passing the solution afew times through a pipette tip and incubated for 10 min at 55 �C. TheRNA solution could then be stored at �20 �C for later microarrayanalysis.

3.6.2. STAT-60 method

Tissue samples (100 mg) were homogenized in 1 ml of STAT-60reagent using a glass–Teflon homogenizer. The sample volume did notexceed 10% of the volume of STAT-60 reagent used for homogenization.The homogenate was stored under room temperature for 5–10 min and

J. Liu, C.A. Lessman / Gene Expression Patterns 8 (2008) 237–247 245

then mixed with 200 ll of chloroform. The sample tube was cappedtightly, shaken vigorously for 15 s and stored at room temperature for5 min. Then the mixture was centrifuged at 12,000g (max) for 45 min at4 �C. The colorless upper aqueous phase was transferred to a fresh tubeand mixed with isopropanol (500 ll isopropanol per 1 ml STAT-60 usedin the original homogenization). The sample was stored at room temper-ature for 30 min and centrifuged at 12,000g for 45 min at 4 �C. The super-natant was carefully removed and the white RNA pellet was washed oncewith 80% ethanol (1 ml ethanol per 1 ml STAT-60 used in the originalhomogenization). The washed RNA pellet was collected by centrifugationat 7500g (max) for 10 min at 4 �C. After the ethanol was discarded, theRNA pellet was air dried, dissolved in RNase-free water and stored at�20 �C for future analysis.

3.7. In situ hybridization

Samples (oocytes or embryos) were fixed in 4% paraformaldehyde at4 �C overnight. Then the samples were gradually dehydrated by addingmethanol (MeOH) to bring the sample to 100% MeOH incubation andstored at �20 �C for 2 h to overnight. The samples were then re-hydratedby successive incubation in 75% MeOH–25% PBS, 50% MeOH–50% PBSand 25% MeOH–75% PBS, 5 min for each time. The chorions of theembryos were manually removed before next step. The oocytes or embryoswere then incubated 4 � 5 min with 100% PBST (PBS plus 0.1% Tween-20). The following procedures were modified from the ZFIN Fish BookMolecular Method (http://zfin.org/zf_info/zfbook/chapt9/9.82.html). Allthe receipts for the related reagents could be found at http://www.fhcrc.org/labs/breeden/Methods/NorthernBlot.html. Briefly, prehy-bridization was done by incubating the samples with prehybridizationmix solution at 37 �C for 3 h. Prehybridization mix solution was thenremoved, discarded and replaced by 200 ll of hybridization mix contain-ing 100–200 ng of FITC-antisense DNA probe (50?30: CCAGGAGATGTAGTCGGGACGTGTGGCGGCGCTGTACTCF, Sigma). Hybrid-ization was performed overnight, in a water bath at 37 �C. On the follow-ing day, the samples were washed by successive incubation inhybridization mix (fast wash, 37 �C), 75% HM-25% 2� SSC (37 �C,15 min), 50% HM–50% 2� SSC (37 �C, 15 min), 25% HM–75% 2� SSC(37 �C, 15 min), 2� SSC (37 �C, 15 min), 0.2� SSC (37 �C, 2 � 15 min),75% 0.2� SSC–25% PBST (room temperature, 15 min), 50% 0.2� SSC–50% PBST (room temperature, 15 min), 25% 0.2� SSC–75% PBST (roomtemperature, 15 min), PBST (room temperature, 15 min) and PBST/5%non-fat milk (room temperature, 3 h). Finally, the samples were incubatedwith anti-FITC secondary antibody (1:200, sigma) overnight with agita-tion at 4 �C. On the third day, the antibody was removed and saved at4 �C. The samples were then extensively washed by incubation in PBST(room temperature, 4 � 15 min), and staining buffer (100 mM Tris, pH9.5; 50 mM MgCl2, 100 mM NaCl and 0.1% Tween-20) (room tempera-ture, 3 � 15 min). The staining procedure was performed at room temper-ature and monitored with a dissection microscope. The staining reactionwas stopped by removing the staining buffer and washing the samples instop solution. Samples were then stored in stop solution at 4 �C in thedark. For microscopy observation, the samples were de-hydrated with100% MeOH gradually until the samples were incubated in 100% MeOHand stored at 4 �C overnight. Then the samples were treated with clearingagent (Benzyl Benzoate:Benzyl alcohol = 2:1) for several times until theywere cleared. The samples were then mounted on slides with clearing agentand observed under a dissection microscope.

3.8. Microarray

Prior to running the microarrays, RNA integrity was determinedusing Bioanalyzer 2100 analysis. Briefly, total RNA (8 lg) was synthe-sized to cDNA using the Superscript Double-Stranded cDNA synthesiskit (Invitrogen Corp., Carlsbad, CA) and poly T-nucleotide primers thatcontain a sequence recognized by T7 RNA polymerase. The newly syn-thesized cDNA was used as a template to generate biotin-labeled vitrotranscribed (IVT) cRNA using the Bio-Array High Yield RNA transcript

labeling kit (Enzo Diagnostics, Inc., Farmingdale, NY). The cRNA(20 lg) was fragmented to strands of 35–200 bases in length. The frag-ment cRNA was hybridized to an Affymetrix GeneChip Zebrafish Gen-ome Array at 45 �C with rotation for 16 h (Affymetrix GeneChipHybridization Oven 320). The GeneChip arrays were washed and stained(streptavidin phycoerythrin) on an Affymetrix Fluidics Station 400, fol-lowed by scanning.

Acknowledgements

This work was supported by the Department of Biology,The University of Memphis Graduate School and theAmerican Society for Cell Biology Minority Affairs Coun-cil. We thank Drs. Thomas Sutter and Shirlean Goodwinof the Feinstone Center for Genomic Studies for their helpin total RNA isolation and microarray assay.

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