RESEARCH ARTICLE 2955
Development 136, 2955-2964 (2009) doi:10.1242/dev.037739
Zebrafish ale oko, an essential determinant of sensory
neuron survival and the polarity of retinal radial glia,
encodes the p50 subunit of dynactin
Xiaotang Jing and Jarema Malicki*
Although microtubule-dependent motors are known to play many essential functions in eukaryotic cells, their role in the context of
the developing vertebrate embryo is less well understood. Here we show that the zebrafish ale oko (ako) locus encodes the p50
component of the dynactin complex. Loss of ako function results in a degeneration of photoreceptors and mechanosensory hair
cells. Additionally, mutant Müller cells lose apical processes and their perikarya translocate rapidly towards the vitreal surface of the
retina. This is accompanied by the accumulation of the apical determinants Nok and Has/aPKC in their cell bodies. ako is required
cell-autonomously for the maintenance of the apical process but not for cell body positioning in Müller glia. At later stages, the
retinotectal projection also degenerates in ako mutants. These results indicate that the p50 component of the dynactin complex is
essential for the survival of sensory neurons and the maintenance of ganglion cell axons, and functions as a major determinant of
apicobasal polarity in retinal radial glia.
INTRODUCTION
Microtubules play many essential roles in the structure and function
of most, if not all, cells. As a component of the cytoskeleton, they
provide structural support for morphological features of cells, such
as axons and dendrites of neurons or apical specializations of
sensory cells (e.g. Ward et al., 1975; Wen et al., 1982). Perhaps even
more importantly, microtubules provide a surface for the movement
of molecular motors, and so are necessary to generate the
mechanical forces that translocate a variety of cargos inside the cell,
including chromosomes during cell division. Among the two classes
of microtubule-dependent motors, kinesins and dyneins,
cytoplasmic dynein provides the predominant mechanism of minus
end-directed movement. This large multi-subunit protein functions
in conjunction with a second multi-subunit protein, dynactin, which
is believed to increase dynein processivity and to mediate at least
some of its cargo interactions (reviewed by Schroer, 2004).
Dynein/dynactin complexes are involved in the transport of a variety
of intracellular cargos, including chromosomes (Sharp et al., 2000),
mitochondria (Ebneth et al., 1998), pre-Golgi structures (Presley et
al., 1997), phagosomes (Blocker et al., 1997) and lysosomes
(Jordens et al., 2001).
The dynactin complex contains at least 11 different polypeptides
organized into two subunits, referred to as the rod and the projecting
arm (reviewed by Schroer, 2004). The rod consists of eight different
components assembled around an octamer of the Arp1 polypeptide.
The remaining three proteins, p150/Glued, p50/Dynamitin and p24
contribute to the arm subunit (Vaughan and Vallee, 1995; WatermanStorer et al., 1995). The rod and arm moieties are thought to connect
to each other via p50/Dynamitin. When overexpressed, p50 causes
their dissociation and abolishes dynactin activity (see references
below).
Department of Ophthalmology, Harvard Medical School, Boston, MA 02114, USA.
*Author for correspondence ([email protected])
Accepted 30 June 2009
When studied in the context of a whole organism, dynein and
dynactin mutations produce severe embryonic phenotypes. The
strongest mutant alleles of the fly dynein heavy chain gene, Dhc64C,
appear to be cell lethal, and the survival of early fly embryos is most
likely due to the presence of the maternal contribution (Gepner et
al., 1996). A similar phenotype is produced by strong Glued
(dynactin 1) alleles in the fly (Garen et al., 1984). Likewise, mouse
embryos mutant for the dynein heavy chain die very early at the
blastocyst stage (Harada et al., 1998). Weaker mutations in genes
encoding dynein and dynactin components are known to affect
several aspects of neuronal development and physiology. A common
consequence of such mutations in flies, nematodes and mice is the
accumulation of synaptic and/or axonal proteins in neuronal cell
bodies or in axonal swellings (Hafezparast et al., 2003; Koushika et
al., 2004; LaMonte et al., 2002; Martin et al., 1999). Another
common function of genes encoding dynein, dynactin, and
associated proteins is the determination of nuclear position in a
variety of cells, including fungal hyphae, the fly oocyte and
zebrafish photoreceptors (Duncan and Warrior, 2002; Efimov and
Morris, 1998; Shu et al., 2004; Tsujikawa et al., 2007; Whited et al.,
2004). Related to these findings is the observation that interkinetic
nuclear migration is abnormal in the retinal neuroepithelium of a
mutant strain that carries a defect in the zebrafish mikre oko gene
(also known as dctn1a) (Del Bene et al., 2008). Interestingly, this
abnormality is associated with cell fate changes during retinal
neurogenesis. Finally, defects in two components of the dynactin
complex, Arp-1 (also known as Arp87C) and p150, were shown to
destabilize synapses at the neuromuscular junction (Eaton et al.,
2002).
To advance the understanding of dynactin activity in
specialized cells of the nervous system, we embarked on the study
of defects in the zebrafish ale oko (ako) locus, which encodes
p50/Dynamitin, a component of the dynactin complex. Our
analysis reveals that ako plays a role in multiple processes,
including the survival of sensory hair cells, the positioning of the
mitotic spindle in neuroepithelial cells, and the maintenance of
the retinotectal projection. The most striking defects are, however,
DEVELOPMENT
KEY WORDS: ale oko, Dynactin, Photoreceptor, Glia, Eye, Retina, Retinotectal, Hair cell, Axon, Polarity, Spindle
2956 RESEARCH ARTICLE
observed in photoreceptor cells and in retinal radial glia: the
akojj50 mutation produces an exceptionally rapid degeneration of
photoreceptors, and a severe cell-autonomous loss of apicobasal
polarity in Müller glia.
MATERIALS AND METHODS
Fish strains
The akojj50 allele was generated in the Malicki laboratory using a standard
ENU mutagenesis protocol (Solnica-Krezel et al., 1994). The maintenance
and breeding of zebrafish strains were performed using standard procedures
(Malicki et al., 2002). Staging of zebrafish embryonic development was
performed as described previously (Kimmel et al., 1995). All animal
protocols used in this study were reviewed by the MEEI Animal Care
Committee.
Histological and ultrastructural analysis
The preparations of plastic and frozen sections, as well as electron
microscopy, were performed as previously described (Avanesov et al., 2005;
Avanesov and Malicki, 2004).
Development 136 (17)
photoreceptor cells allowed us to recognize these cell classes based on
characteristic morphology. To enhance the detection of transgene
expression, sections were stained with an anti-GFP antibody. This approach
enhances a relatively weak CFP signal in the red channel without
compromising the endogenous glial GFP signal in the green channel.
Tg(brn3c:mGFP) and Tg(pax6-DF4:mGFPs220) transgenes in akojj50 mutant
background lines were used to visualize donor-derived ganglion and
amacrine cells, respectively. During the analysis of the retinotectal
projection, only sections that contained ganglion cell axons extending all the
way from the optic chiasm to the tectum were taken into account.
Live imaging
The Tg(gfap:GFP)mi2001 transgenic line was used for live imaging of Müller
glia using the method described previously (Kay et al., 2004). To monitor
the movements of Müller cells and their processes, z-series of images
through selected cells were acquired every 5-10 minutes for a period of 816 hours. Each series consisted of 30-50 images and spanned 20-40 μm in
the z dimension. Images from each z-series were automatically placed in a
separate folder. ImageJ software was used to generate single plane
projections from each z-series and assemble them into movies.
To map the ako locus, we genotyped F2 embryos from a cross between
heterozygous carriers of the akojj50 allele (AB genetic background) and wildtype WIK strain homozygotes. To clone the full-length dynactin 2 cDNA,
zebrafish coding sequences were obtained from Ensemble and NCBI public
databases and used to design amplification primers. Total RNA was isolated
with Trizol (Invitrogen) reagent at 4 days post-fertilization (dpf). Mutation
site was detected by direct sequencing of PCR products amplified from both
genomic DNA (primers: 5⬘-GTTCAGGGCTCCAGTCTGACG-3⬘, 5⬘GACGCTGCTACACTGGACCA-3⬘) and cDNA (primers: 5⬘-ATCTGGACTCTCTGCTCGGA-3⬘, 5⬘-GTCGCCTTGTGTTTGGCAA-3⬘).
Morpholinos were injected into the yolk of embryos at the one-cell stage
as described previously (Malicki et al., 2002). The following two
morpholinos were used: ako-ATG, 5⬘-GGTTCGCGTACTTCGGGTCGGCCAT-3⬘; and ako-SP, 5⬘-GCTCTAAAGTCTCCTATGGACACAG-3⬘.
For phenocopy experiments, we used 0.2 μg/μl of ATG and 2.0 μg/μl of SP
morpholino. For maternal contribution tests, 2.0 μg/μl of ATG MO were
used. To rescue the ako phenotype, the full-length coding sequence of
dynactin 2 was cloned into the pXT7 vector and transcribed as described
previously (Malicki et al., 2002). Approximately 30 pg of mRNA was
injected into embryos at the one-cell stage. mRNA-treated embryos were
fixed at 3.5 dpf and cut in half to determine their genotype. The caudal
halves were used to extract DNA for PCR amplification of the mutation site
and sequencing, and the rostral halves of embryos were sectioned and
stained to determine retinal architecture.
Immunohistochemistry
Fixation, infiltration and sectioning of embryos, and other steps of the
antibody staining procedures have been described previously (Avanesov et
al., 2005). The following primary antibodies and dilutions were used: mouse
Zpr-1 (1:250, Zebrafish International Resource Center), rabbit anti-carbonic
anhydrase (1:250, gift from Dr P. Linser, The Whitney Laboratory), mouse
Zn8 (1:250, Oregon Monoclonal Bank), mouse anti-α-tubulin (1:1500,
Sigma), mouse anti-acetylated-α-tubulin (1:1000, Sigma), rabbit antiGABA (1:100, Sigma), mouse anti-parvalbumin (1:250, Chemicon), rabbit
anti-neuropeptide Y (1:250, Immunostar), mouse anti-serotonin (1:500,
Sigma), mouse HCS1 (1:250, gift from Dr J. Corwin, University of
Virginia), mouse anti-ZO-1 (1:250, Invitrogen), rabbit-anti-Crb (1:250)
(Omori and Malicki, 2006), rabbit anti-aPKC (1:500, Santa Cruz
Biotechnology), rabbit anti-β-catenin (1:250, gift from Dr Steinbeisser,
University of Heidelberg); and rabbit anti-GFP (1:1500, Clontech).
Mosaic analysis
Blastomere transplantations were performed as previously described
(Avanesov and Malicki, 2004), except that transgene expression was used
to detect donor-derived cells. The Tg(gfap:GFPmi2001; pax6-DF4:mCFPq01)
double transgenic ako line was used to simultaneously visualize all donorderived cells based on CFP expression, and donor-derived Müller glia, based
on GFP expression. CFP expression in donor-derived bipolar and
RESULTS
ako mutations affect early stages of
photoreceptor differentiation
The ale oko (akojj50) allele was recovered in a chemical mutagenesis
screen as a mutation that affects eye size of zebrafish larvae (Fig.
1A⬘,B⬘, compare with the wild type in A,B). Very few elongated
photoreceptors were seen on histological sections of ale oko mutants
at 3 dpf (not shown), and no morphologically normal photoreceptors
were observed by 4 dpf (Fig. 1C⬘,D⬘, compare with C,D). Consistent
with that, immunostaining with the Zpr-1 antibody revealed
photoreceptor loss already at 2.5 dpf (red signal in Fig. 1F⬘, compare
with F). By 3 dpf, all photoreceptors displayed rounded morphology
(not shown), and by 4 dpf ~90% of photoreceptors were absent (Fig.
1G⬘, compare with G; see also Fig. S1 in the supplementary
material). By 7 dpf, the photoreceptor cell layer (PRCL) was absent
(not shown). Mutant fish degenerated and died shortly afterwards.
Ultrastructural analysis revealed that outer segments were missing
in most regions of the ale oko mutant retina. The few outer segments
that did differentiate were frequently directed sidewise or even
basally (Fig. 1E⬘, asterisk, compare with E).
To evaluate whether other retinal cell classes are also affected in
ako mutants, transverse sections collected from the vicinity of the
optic nerve were immunostained with several antibodies, including
anti-carbonic anhydrase antibody for Müller glia (Fig. 1G,G⬘), and
four antibodies for amacrine cells: anti-serotonin (Fig. 1H,H⬘), antiparvalbumin (Fig. 1I,I⬘, green), anti-neuropeptide Y (Fig. 1I,I⬘, red)
and anti-GABA (Fig. 1J,J⬘). In parallel, we also studied the ako
mutant phenotype in two transgenic lines, Tg(brn3c:mGFP) and
Tg(gfap:GFP), which express GFP in ganglion cells (Fig. 1K,K⬘)
and Müller glia (Fig. 1F,F⬘), respectively. The most striking
deficiency revealed by these studies was a severe reduction in the
number of Müller cells (Fig. 1F⬘, compare with F). This decrease is
consistent with the phenotype reported for mutations in the moks309
mutant strain, which harbors a mutation in the dynactin 1 (p150)
gene (Del Bene et al., 2008). We did not observe striking differences
in amacrine or ganglion cell populations at 3 dpf.
ale oko is required for the survival of
mechanosensory hair cells
In addition to photoreceptors, mechanosensory hair cells are also
affected in ako mutants. To visualize hair cells, we stained
embryos with either anti-acetylated tubulin antibody, which
DEVELOPMENT
Cloning, morpholino knockdown and phenotypic rescue
Dynactin in vertebrate development
RESEARCH ARTICLE 2957
highlights their apical surface and the kinocilia (Bang et al., 2001;
Omori and Malicki, 2006), or HCS-1 antibody, a hair cell marker
(Gale et al., 2002). Sharply lower numbers of hair cells were
observed in the posterior macula of ako animals at 5 dpf (Fig. 2A⬘,
compare with A; quantitative data is shown in Fig. 2E). ako mutant
cristae and the anterior macula were, however, relatively normal
(Fig. 2B⬘, compare with B; data not shown). On electron
micrographs, we observed an abundance of cellular debris in the
posterior macula of ako mutants, a likely indication of cell death
(arrowheads in Fig. 2C⬘, compare with C). The hair cell defect was
most pronounced in the lateral line: the majority of hair cells in this
organ degenerated between 4 and 5 dpf (Fig. 2D⬘, compare with
the wild type in D; quantitative data is shown in Fig. 2F). These
results indicate that ako is required for the survival of
mechanosensory cells.
The zebrafish ako locus encodes Dynactin 2
By comparing mapping results with data in public databases, we
found that zebrafish dynactin 2 localizes to the vicinity of the ako
locus in linkage group 6 (Fig. 3A). The cloning of dynactin 2 cDNA
revealed that it contains an open-reading frame encoding 405 amino
acids, and is composed of at least 15 exons. The analysis of the
dynactin 2 sequence from akojj50 animals revealed a 9-bp insertion
in the mutant transcript. This insertion introduces an in-frame stop
codon just upstream of exon 11 (Fig. 3C, compare with B) and leads
to the truncation of 143 C-terminal amino acids. To investigate
whether the insertion was specific to the mutant gene, primers
targeted to the insertion sequence were used for RT-PCR
amplification. As expected, these primers generated an amplification
product in the mutant only, supporting the claim that the insertion
was associated with the ako mutant phenotype (Fig. 3D, lanes 2 and
3). The identity of this amplification product was confirmed by BfaI
digestion (Fig. 3D, lane 4). Importantly, in the genomic sequence we
also uncovered a CTGrCAG transition upstream of exon 11 (red
asterisk in Fig. 3C, compare with B). This sequence change is likely
to generate an ectopic AG splice donor site and is most likely
responsible for the 9-bp insertion in the transcript. The injection of
a combination of ATG- and SP- (splice site)-directed anti-dynactin
2 morpholinos produced a small eye phenotype that resembled that
of akojj50 mutants (Fig. 3F, compare with E; see also Table S1 in the
DEVELOPMENT
Fig. 1. ale oko phenotype in the zebrafish retina. (A-B⬘) External phenotypes of akojj50 mutants (A⬘,B⬘), compared with their wild-type siblings
(A,B). Dorsal (A,A⬘) and lateral (B,B⬘) views are shown at 5 dpf. (C-D⬘) Transverse sections of the ako (C⬘,D⬘) and wild-type (C,D) retinae at 4 dpf.
(E,E⬘) Electron micrographs of wild-type (E) and mutant (E⬘) retinae at 3 dpf. OS, outer segment; IS, inner segment. Asterisks indicate abnormal
mutant outer segments. (F-K⬘) Differentiation of retinal cells classes in ako mutants. Transverse cryosections through retinae of wild type (top) and
akojj50 mutants (bottom). (F-G) Müller glia are visualized via the expression of a GFP transgene (F,F⬘, green, arrows) or by antibody staining against
carbonic anhydrase (G,G⬘, red). Photoreceptors are stained with the Zpr-1 antibody (red in F,F⬘, green in G,G⬘). (H-J⬘) Transverse sections stained
with antibodies to subpopulations of amacrine cells: anti-serotonin (H,H⬘, green), anti-neuropeptide Y (I,I⬘, red, arrows), anti-parvalbumin (I,I⬘,
green) and anti-GABA (J,J⬘). (K,K) Sections through retinae of Tg(brn3c:mGFP) transgenic animals, which express GFP in ganglion cells (green).
Photoreceptors are visualized with the Zpr-1 antibody (red). In G-K⬘, asterisks indicate the optic nerve. L, lens; rpe, retinal pigment epithelium;
prcl, photoreceptor cell layer; inl, inner nuclear layer; ipl, inner plexiform layer; gcl, ganglion cell layer. Scale bars: 1 mm in A-B⬘; 40 μm in C,C⬘,F-K⬘;
30 μm in D,D⬘; 5 μm in E,E⬘.
2958 RESEARCH ARTICLE
Development 136 (17)
supplementary material). Consistent with that, dynactin 2 mRNA
injections rescued both small eye size and photoreceptor phenotypes
(Fig. 3H, compare with G; see also Table S2 in the supplementary
material). The overexpression of dynactin 2 from a Tol2 transposonbased vector using a heat-shock-inducible promoter (Kawakami et
al., 2004) also rescued photoreceptor phenotype (data not shown).
These results indicate that the ako mutant phenotype is due to a
defect in the dynactin 2 gene.
To test whether ako transcript is provided maternally, we
performed a knockdown using an anti-ATG morpholino. This
treatment resulted in a delayed epiboly so that in many embryos yolk
plug closure was not completed even at 12 hours post-fertilization
(hpf). By 24 hpf, the yolk was abnormally elongated, the tail region
was severely stunted, and cell death appeared to be abundant along
the entire embryonic axis. These defects were rescued by ako
mRNA injections (see Table S2 and Fig. S3 in the supplementary
material). These observations suggest that ako function is supplied
maternally in the zebrafish embryo.
ako is not necessary for the initial differentiation
of gross morphological features in GCL and INL
neurons
Several studies revealed that the dynactin complex is involved in
axonal outgrowth, synapse stabilization, and/or axonal transport
(Eaton et al., 2002; Grabham et al., 2007; LaMonte et al., 2002). To
investigate whether this is the case in retinal neurons, we inspected
the morphology of ale oko mutant ganglion and amacrine cells. To
distinguish cell-autonomous and non-autonomous aspects of the ako
phenotype and to circumvent lethality of akojj50 mutants, we
performed blastomere transplantation. To do that, we crossed
the akojj50 mutation into Tg(brn3c:mGFP) and Tg(pax6DF4:mGFPs220) transgenic lines, which express GFP in ganglion
and amacrine cells, respectively (Kay et al., 2004; Xiao et al., 2005).
The use of transgenic strains can potentially allow one to detect
donor-derived clones throughout the lifetime of mosaic animals, and
also provides a better resolution for visualizing neuronal
morphology. Mosaic analysis revealed that at 4 dpf homozygous
akojj50 mutant cells contributed to the ganglion cell layer (GCL) and
formed retino-tectal projections in the wild-type environment.
akojj50 projections did not obviously differ from those produced by
wild-type cells in the wild-type environment at the same stage (Fig.
4A⬘,B⬘, compare with wild-type donor-derived cells in A,B). In the
wild-type host environment, akojj50 mutant cells were also able to
produce extensive dendritic trees at 4 dpf (Fig. 4C⬘,D⬘, compare
with wild-type donor cells in C,D). Similarly, the dendrites of akojj50
amacrine cells contributed to the inner plexiform layer and were
grossly indistinguishable from those of wild-type cells (Fig. 4E⬘,
compare with E). Finally, some akojj50 donor-derived bipolar cells
produced normal apical and basal processes in the wild-type
environment at 4 and 6 dpf (Fig. 4F⬘, compare with F; data not
shown). These results suggest that in contrast to photoreceptors,
retinal interneurons as well as ganglion cells do not require dynactin
2 cell-autonomously for the initial differentiation of their gross
morphological features. However, we cannot exclude the possibility
that subtle changes might exist in the stratification of dendrites, the
distribution of synaptic termini or in the innervation of the optic
tectum by akojj50 cells.
To evaluate whether dynactin 2 is required for the maintenance of
ganglion cell morphology, we repeated mosaic analysis at 10 dpf. In
most cases (12/17), wild-type donor-derived ganglion cells
transplanted into wild-type retinae differentiated densely branched
processes in the optic tectum (Fig. 4G,H). In contrast to wild-type
cells, we have not observed densely branched axonal processes in
any transplants that involved akojj50 mutant ganglion cells at this
stage (0/11; Fig. 4G⬘,H⬘). Instead, mutant projections differentiated
few branches or an irregular branching pattern. In some cases, we
DEVELOPMENT
Fig. 2. ale oko hair cell phenotype. (A-B⬘) Whole-mount staining of ear sensory maculae with anti-acetylated-α-tubulin antibody (green) and
phalloidin (red, actin) to visualize hair cells and their stereocilia, respectively. Posterior (A⬘) and anterior (B⬘) maculae of ako mutants are shown.
(C,C⬘) Electron micrographs of auditory hair cells in the posterior macula of wild-type (C) and ako mutant (C⬘) zebrafish larvae. Cellular debris
(arrowheads) is present in the mutant tissue. (D,D⬘) In the ako mutant lateral line (D⬘), few hair cells survive by 5 dpf compared with the wild type
(D). (E) Quantitation of hair cell numbers in the maculae of wild-type and akojj50 animals at 5 dpf. The average numbers of hair cells in the anterior
(AM) and the posterior (PM) maculae are provided (n=6). For posterior macula, P<0.01 (Student’s t-test). (F) The average number of hair cells per
lateral line neuromast in wild-type and akojj50 animals (n=6). At 4 and 5 dpf, P<0.0001 (Student’s t-test). A,A⬘,D,D⬘ show face views of the apical
surface; in B-C⬘, apical is up. Scale bars: 5 μm in C,C⬘; 20 μm in A-B⬘,D,D⬘.
Fig. 3. The ale oko locus encodes a zebrafish dynactin 2
homolog. (A) Map of the ako genomic region and exon/intron
structure of the ako transcript. The position of mutant sequence in the
akojj50 allele is indicated (red arrowhead). (B,C) Comparison of sequence
trace data from wild-type (B) and mutant (C) animals. The akojj50
mutation activates a cryptic splice site in the genomic sequence. This, in
turn, introduces a 9-bp insertion (red) that encodes a premature stop
codon. (D) RT-PCR amplification of the dynactin 2 transcript using a
primer that recognizes the mutant insertion sequence. Lane 1, DNA
ladder; lane 2, amplification of the mutant transcript produces a single
robust band; lane 3, amplification of the wild-type transcript does not
produce a detectable product; lane 4, BfaI digestion confirms the
identity of the amplification product from the mutant; lanes 5 and 6,
control PCR amplification of actin from wild-type (5) and mutant (6)
animals. The positions of amplification primers in the wild type and
mutant are indicated on the diagram below. (E-H) Phenocopy and
rescue of photoreceptor defects in ako mutant animals. Transverse
cryosections through embryonic retinae were stained with the Zpr-1
antibody to visualize double cones (red). Green signal in E,F originates
from the Tg(brn3c:mGFP) transgene. The injection of ako (H), but not
GFP (G), mRNA rescues photoreceptor morphology in ako mutants. L,
lens. Scale bar: 40 μm.
observed swellings near the termini of akojj50 mutant axons (Fig.
4H⬘, arrows). These results indicate that dynactin 2 is essential for
the maintenance of the retino-tectal projection.
ako functions non-cell-autonomously in
photoreceptor morphogenesis and survival
To determine whether ako functions cell-autonomously in
photoreceptor cells, we generated genetically mosaic animals.
Donor cells were distinguished based on expression of the Tg(pax6DF4:mCFPq01) transgene, and detected by staining with an anti-
RESEARCH ARTICLE 2959
Fig. 4. Cell-autonomous aspects of ako function in neuronal
differentiation. (A-H⬘) Transverse sections through mosaic retinae at
4 (A-F⬘) and 10 (G-H⬘) dpf. Donor cells were derived from the
following fluorescent protein transgenic lines: Tg(brn3c:mGFP) in A-D⬘
and G-H⬘ to visualize ganglion cells; Tg(pax6-DF4:mGFPs220) in E,E⬘ to
visualize amacrine cells; and Tg(pax6-DF4:mCFPq01) in F,F⬘ to reveal
the morphology of bipolar cells. Host tissue is unlabelled. Sections in
A,A⬘ are anterior to those in B,B⬘. The genotypes of donor and host
larvae are indicated above each column. No differences are observed
between wild-type and mutant ganglion, bipolar or amacrine cells at
4 dpf. (G,H) Donor-derived wild-type ganglion cells differentiate
densely branched processes in the optic tectum of wild-type hosts at
10 dpf. (G⬘,H⬘) By contrast, donor-derived akojj50 mutant cells
differentiate few axons, which frequently do not branch, or exhibit an
irregular branching pattern. In some cases, swellings form near the
termini of akojj50 mutant axons (arrows in H⬘). Vertical dashed lines
indicate the midline; arrowheads indicate the optic tecta; L, lens.
Scale bars: 150 μm in A-B⬘,G,G⬘; 40 μm in C-D⬘,H,H⬘; 50 μm in E,E⬘;
30 μm in F,F⬘.
GFP antibody. Both wild-type and mutant-derived donor cells
contributed to all layers of wild-type retinae, including the PRCL at
3.5, 4 and 6 dpf. At 4 dpf, the ratio of donor-derived photoreceptors
to INL cells was 29% for mutant cells in the wild-type environment,
compared with 35% for wild-type cells in the wild-type environment
(see Table S3 in the supplementary material). This survival rate is
dramatically better than that of akojj50 photoreceptors in akojj50
mutant retinae, in which ~90% of photoreceptor cells died by 4 dpf
(see Fig. S1 legend in the supplementary material). In contrast to ako
mutant retinae, ako mutant cells in the wild-type host environment
formed morphologically distinguishable, elongated photoreceptors
even at 6 dpf (see Fig. S2 in the supplementary material). These
results indicate that, similar to mikre oko (Tsujikawa et al., 2007),
the ale oko photoreceptor phenotype contains a non-cellautonomous component.
DEVELOPMENT
Dynactin in vertebrate development
2960 RESEARCH ARTICLE
Development 136 (17)
ale oko is required for the differentiation of
Müller glia
Our initial analysis of the ako retina revealed abnormal morphology
of Müller glia (Fig. 1F⬘, compare with the wild type in F). To
monitor gliogenesis, we crossed the akojj50 mutant allele into the
Tg(gfap:GFP)mi2001 transgenic line, which expresses GFP in Müller
glia (Bernardos and Raymond, 2006). In wild-type larvae, GFPexpressing Müller glia were well differentiated by 60 hpf (Fig. 5A).
Their processes spanned the entire retinal thickness, and perikarya
localized to the INL. By contrast, in the akojj50 retina, the number of
GFP-positive glial cells was reduced by ~40% (19±6, n=5 sections)
compared with the wild type (33±6, n=5), and 40% (39/96, n=5) of
mutant Müller cell perikarya were mislocalized, so that they were
found basal to the INL, compared with ~7% (11/168, n=5) in the
wild type (Fig. 5A; data not shown). In addition, we found that only
18% (17/96, n=5 sections) of these cells had distinguishable apical
processes at 60 hpf, compared with 82% (137/168, n=5) in the wild
type. By 4 dpf, 74% (240/324, n=13) of ako Müller cell perikarya
were basally mispositioned, and apical processes were absent (Fig.
5). In contrast to these observations, brain radial glia appeared to be
grossly normal at least until 8 dpf (not shown). These observations
indicate that dynactin 2 is a key determinant of Müller glia
differentiation.
Müller glia degeneration involves a rapid basal
displacement of their nuclei
To investigate how the dynactin complex is involved in the
positioning of Müller cell perikarya, we performed live imaging
of these cells in the akojj50 mutant strain carrying the
Tg(gfap:GFP)mi2001 transgene. This transgene expresses GFP in
Müller glia making it possible to continuously monitor the
morphology of these cells in living animals. Imaging was
performed for 8-16 hours, starting at 72-76 hpf. While using this
imaging approach, respectively 94% (73/78, n=3) and 95% (74/78,
n=3) of wild-type glial cells featured the apical and the basal
process during the recording period. In all wild-type cells imaged,
apical and basal processes, once differentiated, were continuously
maintained, and we did not observe a displacement of cell
perikarya (see Movie 1 in the supplementary material). By
contrast, in ako mutants only 46% (32/69, n=6) of cells
differentiated the apical process, whereas 84% (58/69, n=6)
featured the basal process. All apical processes of mutant cells
were retracted during the recording period. Interestingly, this was
frequently followed by a rapid basal displacement of perikarya,
which we observed in 38% (12/32) of cells imaged (Fig. 5B; see
also Movies 2 and 3 in the supplementary material). This
displacement frequently occurred in less than 1 hour (Fig. 5B,C),
and in the most extreme cases its maximum velocity exceeded 1
μm/minute. In wild-type cells, the highest displacement velocity
reached ~0.2 μm/minute, and was associated with small saltatory
apicobasal movements of nuclei (Fig. 5D). It is worth pointing it
out that we have not observed any cases of apical displacement of
Müller cell perikarya (0/32), and that all cells that displayed basal
perikaryal displacement maintained their basal process throughout
the course of their translocation. These results indicate that
dynactin 2 plays a key role in the maintenance of the apical
process, and functions in localizing Müller cell bodies to the inner
plexiform layer.
DEVELOPMENT
Fig. 5. The phenotype of Müller glia.
(A,A⬘) Transverse cryosections through the
retinae of wild type (A) and akojj50 mutant
(A⬘) lines at 60 hpf. Müller glia are visualized
by Tg(gfap:GFP) transgene expression. Graphs
to the right show the quantitation of these
phenotypes. ‘Cell body position’ refers to the
position of Müller cell bodies in the inner
nuclear layer. (B) A series of images from a
time-lapse recording of a mutant Müller cell.
Time is indicated above each image in hours
and minutes (h:min). Cell body positions are
indicated with red asterisks. (C) Cell body
position expressed as a ratio of its distance
from the inner limiting membrane (parameter
‘a’ in B) and retinal thickness (parameter ‘b’ in
B). Cell body positions during time-lapse
recording sessions are plotted for three wildtype (blue) and eight mutant (red) cells. All
data were collected from the peripheral retina
from 72-96 hpf. (D) The velocity of perikaryal
displacement calculated for the same set of
cells as in C. L, lens. Scale bars: 40 μm in A,A⬘;
20 μm in B.
Fig. 6. Mosaic analysis of ako Müller glia mutant phenotype.
(A-E⬘) Transverse cryosections through retinae of mosaic zebrafish at
the stages indicated below each image. Blastomere transplantations
were performed using the Tg(gfap:GFPmi2001; pax6-DF4:mCFPq01) donor
double transgenic line either wild type or mutant at the ako locus, as
indicated at the top of each column. A non-transgenic wild-type line
was used as the host. Sections were stained with an anti-GFP antibody
to visualize all donor-derived cells (both CFP and GFP expression, red).
Fluorescence of endogenously expressed GFP marks donor-derived
Müller glia (green signal appears yellow against the red background).
Sections in E,E⬘ are stained with the Zpr-1 antibody to visualize the
photoreceptor cell layer. No defects in the morphology of Zpr-1postitive cells in the vicinity of abnormal glial cells were observed
(asterisk in E⬘). (F) Quantitation of mosaic analysis data. Percentages of
cells with normal cell body position and apical process are provided.
(G) The apical process of the wild-type Müller cell. Arrowheads and
arrows indicate OLM and OPL, respectively. Asterisks indicate Müller cell
perikarya. L, lens. Scale bars: 20 μm in A-E⬘; 10 μm in G.
ako is required cell-autonomously for the
differentiation of apical processes in Müller glia
To investigate whether the Müller glia phenotype results from
defects intrinsic to these cells or is a consequence of abnormal
interactions with surrounding cell classes, we used genetic mosaics.
Blastomere transplantation was performed using the
Tg(gfap:GFPmi2001; pax6-DF4:mCFPq01) double transgenic ako line
as the donor, and a non-transgenic line as the host. Embryos were
stained with an anti-GFP antibody to visualize all donor-derived
cells based on CFP expression (Fig. 6, red signal), and GFP
endogenous transgene expression was used to distinguish donorderived glial cells (Fig. 6, green signal, appears yellow owing to the
overlap with the red signal). Wild-type Müller glia in wild-type hosts
displayed several distinct morphological features: their cell bodies
localized to the INL, and their processes extended both apically and
basally, contacting the outer (OLM) and inner (ILM) limiting
membranes, respectively. Moreover, the apical terminus of each glial
cell was richly branched, and contacted numerous photoreceptors
(Fig. 6G). All of these features were already well formed in wildtype cells at 3.5 dpf (Fig. 6A), and did not display significant
changes at least until 8 dpf (Fig. 6D). Similarly, akojj50 mutant glia
in the wild-type host environment frequently differentiated apical
RESEARCH ARTICLE 2961
processes at 3.5 dpf (24/50, n=8 retinae; Fig. 6A⬘; quantitative data
in Fig. 6F). By 4 dpf, however, the majority of akojj50 Müller glia
that differentiated in the wild-type environment lost apical and basal
processes (39/40, n=6; Fig. 6B⬘, compare with B; quantitated in Fig.
6F). This stunted morphology persisted for several days, until at
least 8 dpf (Fig. 6D⬘).
The loss of the apical process in Müller cells could result from a
photoreceptor defect. This possibility may appear likely as retinal cell
clones tend to contribute to all layers of the retina simultaneously
(Holt et al., 1988; Turner et al., 1990; Wetts and Fraser, 1988). For
this hypothesis to be correct, mutant photoreceptor cells should be
present apical to defective Müller glia at least during early stages of
neurogenesis. Contrary to this prediction, we observed aberrant glial
cells (9/19) in akojj50 clones that did not feature photoreceptors at 4
dpf and later (e.g. Fig. 6D⬘,E⬘). This observation eliminates the
possibility that mutant photoreceptors survive in mosaic retinae but
are unable to interact with Müller glia and thus cause glial defects. In
an alternative scenario, mutant photoreceptors could be present
earlier in development but then die and, consequently, cause glia
defects. This scenario appears unlikely for several reasons. First, one
has to note that a single glial cell (even mutant) contacts many
photoreceptors (see Fig. 6A⬘,G), and thus many photoreceptor cells
would have to die to result in the loss of apical junctions of Müller
glia. Second, in the retinae of several zebrafish photoreceptor
mutants, Müller glia persist largely intact following a complete
photoreceptor loss (Doerre and Malicki, 2002). Third, to detect
photoreceptor defects, we stained mosaic retinae with Zpr-1 antibody,
which recognizes double cones (example shown in Fig. 6E,E⬘). We
have not seen any aberrations suggestive of cell death in the
photoreceptor cell layer. Finally, if photoreceptor cells died rapidly,
then the ratio of photoreceptors to other cell classes would be
drastically changed in akojj50 mutant clones at 3.5-4 dpf as compared
with control wild-type donor-derived clones. This is not the case.
Given these considerations, our data support conclusion that the
Müller glia defect is cell autonomous.
ako mutant Müller glia accumulate determinants
of apicobasal polarity in their cytoplasm
Müller glia are highly polarized, their surface is subdivided into
apical and basolateral domains by a belt of cell junctions, the outer
limiting membrane (OLM) (Rodieck, 1973). We have previously
shown that apical surface determinants, Nagie oko (Nok, also known
as Mpp5a) and Crumbs, localize to the vicinity of the OLM (Omori
and Malicki, 2006; Wei and Malicki, 2002). To investigate the fate
of apical surface proteins in ako mutants, we stained akojj50 retinae
with antibodies to ZO-1 (also known as Tjp1), aPKC and Nok at 3
dpf. As expected, in the wild-type retina, ZO-1, Nok and aPKC
staining was found almost exclusively at the OLM (Fig. 7A,C,E).
Very few ectopically localized staining areas were observed near the
inner surface of the retina (less than one per section for Nok and
aPKC, and four per section for ZO-1; n=5 sections). By contrast, all
three proteins frequently accumulated in ectopic aggregates in
displaced Müller cells of the ako mutant (9, 10 and 16/section for
Nok, aPKC and ZO-1, respectively; n=5; Fig. 7A-F⬙). Respectively,
~80% and ~90% of ectopic Nok and aPKC spots colocalized with
ZO-1 (Fig. 7B-F⬙). These results indicate that ako is required for the
proper localization of apical polarity proteins in Müller glia. As the
centrosome of Müller glia localizes to the apical surface, far away
from the bulk of the cytoplasm (Fig. 7G-H⬘), abnormal localization
of these polypeptides is likely to have resulted from a defect in
minus end-directed microtubule-dependent transport (schematically
illustrated in Fig. 7I).
DEVELOPMENT
Dynactin in vertebrate development
2962 RESEARCH ARTICLE
Development 136 (17)
Fig. 7. Apical polarity determinants are mislocalized in Müller glia. Transverse cryosections through wild-type or ako mutant retinae at 3 dpf.
Sections are stained with antibodies against Nok, ZO-1, aPKC or γ-tubulin as indicated. The expression of the Tg(gfap:GFP) transgene marks Müller
glia (green). (A,B) Nok (red) and ZO-1 (blue) localization in wild-type (A) and ako mutant (B) Müller glia. (B⬘-B⵮) Detail of image shown in B; red,
green and far red (in blue) channels are shown separately. (C-F⬙) Higher magnifications of Nok (red), Has/aPKC (red) and ZO-1 (blue) staining
patterns. Retinae of mutant (D-D⬙,F-F⬙) and wild-type (C-C⬙,E-E⬙) siblings are shown next to each other. Nok (C⬘,D⬘) and aPKC (E⬘,F⬘) proteins
colocalize with ZO-1 (bottom row of images), both in apical processes of wild-type Müller glia and in displaced glia of ako mutant eyes. Each
column presents images obtained from a single section. Red and far red (in blue) channels are shown separately. (G-H⬘) Centrosome localization in
Müller glia. Transverse cryosections through the retina stained with an antibody against γ-tubulin (red) to visualize basal bodies (arrows). (G⬘,H⬘) Red
channel only. (I) Schematic drawing of a neuron (left) and a Müller cell (right) indicating the position of the centrosome (red dot) and the
presumptive polarity of microtubules (blue lines). Asterisks indicate the vitreal (basal) surface of the retina; arrowheads indicate the OLM. Scale bars:
25 μm in A,B; 15 μm in C-F⬙; 10 μm in G-H⬘.
supplementary material). An extreme example of this is shown in
Fig. S4B⬘ in the supplementary material. These observations
indicate that ako function is required for the proper orientation of the
mitotic spindle.
DISCUSSION
We demonstrated that p50, a dynactin component, is involved in
several aspects of nervous system development. Prior to these
studies, knockout of dynein heavy chain in the mouse was shown
to produce early embryonic lethality (Harada et al., 1998). This
phenotype is much more severe than those of the zebrafish
dynactin mutants mok and ako (this work) (Del Bene et al., 2008;
Tsujikawa et al., 2007). A likely factor that accounts for these
differences is the presence of the maternal contribution in the
zebrafish embryo. akojj50 is the second mutation of a dynactin
component to be found in zebrafish. Previously characterized
mutant strains, mokm632 and moks309, carry defects in the p150
(dynactin 1) gene (Del Bene et al., 2008; Doerre and Malicki,
2001; Malicki et al., 1996; Tsujikawa et al., 2007). The akojj50
mutant phenotype is more severe than that of mokm632 in both
photoreceptors and Müller glia (see Fig. S1 in the supplementary
material).
DEVELOPMENT
ale oko function is required for the proper
orientation of the mitotic spindle
Previous studies demonstrated in several different contexts that
dynactin is involved in the positioning of the mitotic spindle (Eshel
et al., 1993; Gonczy et al., 1999; Li et al., 1993; Muhua et al., 1994;
Toyoshima et al., 2007). To investigate whether ako is required for
proper spindle orientation in neural progenitors of the retina,
embryos were genotyped by sequencing at 30 hpf, sectioned, and
stained with anti-α-tubulin antibody to visualize mitotic spindles.
The orientation of mitotic spindle axes were evaluated relative to the
apical surface of the neuroepithelium (see Fig. S4A in the
supplementary material). As reported previously (Das et al., 2003),
in wild-type embryos mitotic spindles were largely parallel to the
apical surface of the neuroepithelium (see Fig. S4B in the
supplementary material). Only 7% of cell divisions (8/121, in 39
embryos) occurred at orientations greater than 30° (see Fig. S4C in
the supplementary material). In ako mutant embryos, however,
mitotic spindles were frequently positioned at wider angles relative
to the apical surface. Twelve percent of akojj50 mitotic spindles
(25/208, in 41 embryos) were oriented at an angle greater than 60°
and 25% (53/208) of mitotic spindles were oriented at an angle
greater than 30° relative to the apical surface (see Fig. S4D in the
Why are sensory cells and Müller glia particularly sensitive to the
loss of dynactin? The answer to this question might lie in the fact
that these cell types feature an elongated and highly polarized
morphology; similar to epithelia, their surfaces are subdivided by
cell junctions into apical and basolateral domains, and their
centrosomes are positioned at the apical cell surface, far away from
biosynthetically active perinuclear regions (Dowling, 1987;
Rodieck, 1973). As a consequence of this, apically directed cargo
travels a long distance and relies on minus-end-directed
microtubule-dependent motors (Troutt and Burnside, 1988). The
importance of a robust apically directed cytoplasmic transport
mechanism is particularly obvious in photoreceptors. In these cells,
many phototransduction cascade components are likely to be
transported towards the outer segment via a microtubule-dependent
mechanism. One has to note, however, that opsin itself may not
require dynactin for its transport (Tai et al., 1999; Tsujikawa et al.,
2007).
A striking feature of cell polarity in Müller glia is the particularly
large distance between their nuclei and apically localized
centrosomes (Fig. 7G,G⬘,I). Each Müller cell extends a long apical
process which branches around several photoreceptors and forms
junctions at their surface (Fig. 6G). The centrosome, presumably the
main microtubule organizing center, localizes to the very apical
termini of Müller glia (Fig. 7G,G⬘,I), and thus the minus ends of
cytoplasmic microtubules most likely also point apically in these
cells. Given the exceptional length and extensive branching of the
apical process in Müller glia, its formation and maintenance is likely
to require a very active minus end-directed microtubule-dependent
trafficking system. Consequently, compared with other cell types,
the reliance of Müller glia on the minus end-directed motor dynein
may be more pronounced, and so defects in this motor might lead to
cell degeneration. The observation that the apical determinants Nok
and Has/aPKC accumulate in the cell bodies of Müller glia supports
this scenario. As transport mechanisms in radial glia are poorly
investigated so far, the analysis of the ako mutant offers one of the
first insights into this area.
Although the mechanism of ako involvement in the maintenance
of the retinotectal projection remains unclear, defects that we
observed in the ako mutant might be medically relevant, as ganglion
cell degeneration, glaucoma, is a frequent cause of human blindness
(Quigley, 1996). It has been, in fact, documented that dynein
components accumulate at the optic nerve head following an
increase of intraocular pressure, a major risk factor for this disease
(Martin et al., 2006). Finally, we note that the mitotic spindle
phenotype in akojj50 mutants suggests that dynactin might be
required for the proper distribution of cell fate determinants during
asymmetric cell divisions and thereby might affect cell fate.
Although the role of mitotic spindle positioning in retinal cell fate
decisions remains unclear (Cayouette and Raff, 2003; Das et al.,
2003; Silva et al., 2002; Zigman et al., 2005), in other developmental
contexts the contribution of mitotic spindle positioning to the
outcome of asymmetric cell divisions is well documented (for a
review, see Gonczy, 2008).
Acknowledgements
Drs Jon Clarke, Ching-Hwa Sun and Paula Alexandre provided helpful
comments on previous versions of this manuscript. We are also grateful to Drs
Jeffery Corwin, Herbert Steinbeisser, Herwig Baier and Pamala Raymond for
antibodies and transgenic animals. Drs Rachel Wong and Chi-Bin Chien
provided helpful advice regarding live imaging of zebrafish embryos. The ako
locus was initially mapped by the mapping facility at the University of
Louisville. These studies were supported by a grant from the Knights Templar
Eye Foundation (to X.J.) and an NIH R01 EY016859 award (to J.M.). Deposited
in PMC for release after 12 months.
RESEARCH ARTICLE 2963
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/136/17/2955/DC1
References
Avanesov, A. and Malicki, J. (2004). Approaches to study neurogenesis in the
zebrafish retina. Methods Cell Biol. 76, 333-384.
Avanesov, A., Dahm, R., Sewell, W. F. and Malicki, J. J. (2005). Mutations that
affect the survival of selected amacrine cell subpopulations define a new class of
genetic defects in the vertebrate retina. Dev. Biol. 285, 138-155.
Bang, P. I., Sewell, W. F. and Malicki, J. J. (2001). Morphology and cell type
heterogeneities of the inner ear epithelia in adult and juvenile zebrafish (Danio
rerio). J. Comp. Neurol. 438, 173-190.
Bernardos, R. L. and Raymond, P. A. (2006). GFAP transgenic zebrafish. Gene
Expr. Patterns 6, 1007-1013.
Blocker, A., Severin, F. F., Burkhardt, J. K., Bingham, J. B., Yu, H., Olivo, J. C.,
Schroer, T. A., Hyman, A. A. and Griffiths, G. (1997). Molecular requirements
for bi-directional movement of phagosomes along microtubules. J. Cell Biol.
137, 113-129.
Cayouette, M. and Raff, M. (2003). The orientation of cell division influences
cell-fate choice in the developing mammalian retina. Development 130, 23292339.
Das, T., Payer, B., Cayouette, M. and Harris, W. A. (2003). In vivo time-lapse
imaging of cell divisions during neurogenesis in the developing zebrafish retina.
Neuron 37, 597-609.
Del Bene, F., Wehman, A. M., Link, B. A. and Baier, H. (2008). Regulation of
neurogenesis by interkinetic nuclear migration through an apical-basal notch
gradient. Cell 134, 1055-1065.
Doerre, G. and Malicki, J. (2001). A mutation of early photoreceptor
development, mikre oko, Reveals cell- cell interactions involved in the survival
and differentiation of zebrafish photoreceptors. J. Neurosci. 21, 6745-6757.
Doerre, G. and Malicki, J. (2002). Genetic analysis of photoreceptor cell
development in the zebrafish retina. Mech. Dev. 110, 125-138.
Dowling, J. (1987). The Retina. Cambridge, MA: Harvard University Press.
Duncan, J. E. and Warrior, R. (2002). The cytoplasmic dynein and kinesin motors
have interdependent roles in patterning the Drosophila oocyte. Curr. Biol. 12,
1982-1991.
Eaton, B. A., Fetter, R. D. and Davis, G. W. (2002). Dynactin is necessary for
synapse stabilization. Neuron 34, 729-741.
Ebneth, A., Godemann, R., Stamer, K., Illenberger, S., Trinczek, B. and
Mandelkow, E. (1998). Overexpression of tau protein inhibits kinesindependent trafficking of vesicles, mitochondria, and endoplasmic reticulum:
implications for Alzheimer’s disease. J. Cell Biol. 143, 777-794.
Efimov, V. P. and Morris, N. R. (1998). A screen for dynein synthetic lethals in
Aspergillus nidulans identifies spindle assembly checkpoint genes and other
genes involved in mitosis. Genetics 149, 101-116.
Eshel, D., Urrestarazu, L. A., Vissers, S., Jauniaux, J. C., van Vliet-Reedijk, J.
C., Planta, R. J. and Gibbons, I. R. (1993). Cytoplasmic dynein is required for
normal nuclear segregation in yeast. Proc. Natl. Acad. Sci. USA 90, 1117211176.
Gale, J. E., Meyers, J. R., Periasamy, A. and Corwin, J. T. (2002). Survival of
bundleless hair cells and subsequent bundle replacement in the bullfrog’s
saccule. J. Neurobiol. 50, 81-92.
Garen, A., Miller, B. R. and Paco-Larson, M. L. (1984). Mutations affecting
functions of the Drosophila gene glued. Genetics 107, 645-655.
Gepner, J., Li, M., Ludmann, S., Kortas, C., Boylan, K., Iyadurai, S. J.,
McGrail, M. and Hays, T. S. (1996). Cytoplasmic dynein function is essential in
Drosophila melanogaster. Genetics 142, 865-878.
Gonczy, P. (2008). Mechanisms of asymmetric cell division: flies and worms pave
the way. Nat. Rev. Mol. Cell Biol. 9, 355-366.
Gonczy, P., Pichler, S., Kirkham, M. and Hyman, A. A. (1999). Cytoplasmic
dynein is required for distinct aspects of MTOC positioning, including
centrosome separation, in the one cell stage Caenorhabditis elegans embryo. J.
Cell Biol. 147, 135-150.
Grabham, P. W., Seale, G. E., Bennecib, M., Goldberg, D. J. and Vallee, R. B.
(2007). Cytoplasmic dynein and LIS1 are required for microtubule advance
during growth cone remodeling and fast axonal outgrowth. J. Neurosci. 27,
5823-5834.
Hafezparast, M., Klocke, R., Ruhrberg, C., Marquardt, A., Ahmad-Annuar,
A., Bowen, S., Lalli, G., Witherden, A. S., Hummerich, H., Nicholson, S. et
al. (2003). Mutations in dynein link motor neuron degeneration to defects in
retrograde transport. Science 300, 808-812.
Harada, A., Takei, Y., Kanai, Y., Tanaka, Y., Nonaka, S. and Hirokawa, N.
(1998). Golgi vesiculation and lysosome dispersion in cells lacking cytoplasmic
dynein. J. Cell Biol. 141, 51-59.
Holt, C., Bertsch, T., Ellis, H. and Harris, W. (1988). Cellular determination in the
Xenopus retina is independent of lineage and birth date. Neuron 1, 15-26.
Jordens, I., Fernandez-Borja, M., Marsman, M., Dusseljee, S., Janssen, L.,
Calafat, J., Janssen, H., Wubbolts, R. and Neefjes, J. (2001). The Rab7
DEVELOPMENT
Dynactin in vertebrate development
effector protein RILP controls lysosomal transport by inducing the recruitment of
dynein-dynactin motors. Curr. Biol. 11, 1680-1685.
Kawakami, K., Takeda, H., Kawakami, N., Kobayashi, M., Matsuda, N. and
Mishina, M. (2004). A transposon-mediated gene trap approach identifies
developmentally regulated genes in zebrafish. Dev. Cell 7, 133-144.
Kay, J. N., Roeser, T., Mumm, J. S., Godinho, L., Mrejeru, A., Wong, R. O. and
Baier, H. (2004). Transient requirement for ganglion cells during assembly of
retinal synaptic layers. Development 131, 1331-1342.
Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B. and Schilling, T. F.
(1995). Stages of embryonic development of the zebrafish. Dev. Dyn. 203, 253310.
Koushika, S. P., Schaefer, A. M., Vincent, R., Willis, J. H., Bowerman, B. and
Nonet, M. L. (2004). Mutations in Caenorhabditis elegans cytoplasmic dynein
components reveal specificity of neuronal retrograde cargo. J. Neurosci. 24,
3907-3916.
LaMonte, B. H., Wallace, K. E., Holloway, B. A., Shelly, S. S., Ascano, J.,
Tokito, M., Van Winkle, T., Howland, D. S. and Holzbaur, E. L. (2002).
Disruption of dynein/dynactin inhibits axonal transport in motor neurons causing
late-onset progressive degeneration. Neuron 34, 715-727.
Li, Y. Y., Yeh, E., Hays, T. and Bloom, K. (1993). Disruption of mitotic spindle
orientation in a yeast dynein mutant. Proc. Natl. Acad. Sci. USA 90, 1009610100.
Malicki, J., Neuhauss, S. C., Schier, A. F., Solnica-Krezel, L., Stemple, D. L.,
Stainier, D. Y., Abdelilah, S., Zwartkruis, F., Rangini, Z. and Driever, W.
(1996). Mutations affecting development of the zebrafish retina. Development
123, 263-273.
Malicki, J. J., Pujic, Z., Thisse, C., Thisse, B. and Wei, X. (2002). Forward and
reverse genetic approaches to the analysis of eye development in zebrafish.
Vision Res. 42, 527-533.
Martin, K. R., Quigley, H. A., Valenta, D., Kielczewski, J. and Pease, M. E.
(2006). Optic nerve dynein motor protein distribution changes with intraocular
pressure elevation in a rat model of glaucoma. Exp. Eye Res. 83, 255-262.
Martin, M., Iyadurai, S. J., Gassman, A., Gindhart, J. G., Jr, Hays, T. S. and
Saxton, W. M. (1999). Cytoplasmic dynein, the dynactin complex, and kinesin
are interdependent and essential for fast axonal transport. Mol. Biol. Cell 10,
3717-3728.
Muhua, L., Karpova, T. S. and Cooper, J. A. (1994). A yeast actin-related protein
homologous to that in vertebrate dynactin complex is important for spindle
orientation and nuclear migration. Cell 78, 669-679.
Omori, Y. and Malicki, J. (2006). oko meduzy and related crumbs genes are
determinants of apical cell features in the vertebrate embryo. Curr. Biol. 16, 945957.
Presley, J. F., Cole, N. B., Schroer, T. A., Hirschberg, K., Zaal, K. J. and
Lippincott-Schwartz, J. (1997). ER-to-Golgi transport visualized in living cells.
Nature 389, 81-85.
Quigley, H. A. (1996). Number of people with glaucoma worldwide. Br. J.
Ophthalmol. 80, 389-393.
Rodieck, R. W. (1973). The Vertebrate Retina: Principles of Structure and Function.
San Francisco, CA: W. H. Freeman.
Schroer, T. A. (2004). Dynactin. Annu. Rev. Cell Dev. Biol. 20, 759-779.
Development 136 (17)
Sharp, D. J., Rogers, G. C. and Scholey, J. M. (2000). Cytoplasmic dynein is
required for poleward chromosome movement during mitosis in Drosophila
embryos. Nat. Cell Biol. 2, 922-930.
Shu, T., Ayala, R., Nguyen, M. D., Xie, Z., Gleeson, J. G. and Tsai, L. H. (2004).
Ndel1 operates in a common pathway with LIS1 and cytoplasmic dynein to
regulate cortical neuronal positioning. Neuron 44, 263-277.
Silva, A. O., Ercole, C. E. and McLoon, S. C. (2002). Plane of cell cleavage and
numb distribution during cell division relative to cell differentiation in the
developing retina. J. Neurosci. 22, 7518-7525.
Solnica-Krezel, L., Schier, A. and Driever, W. (1994). Efficient recovery of ENUinduced mutations from the zebrafish germline. Genetics 136, 1-20.
Tai, A. W., Chuang, J. Z., Bode, C., Wolfrum, U. and Sung, C. H. (1999).
Rhodopsin’s carboxy-terminal cytoplasmic tail acts as a membrane receptor for
cytoplasmic dynein by binding to the dynein light chain Tctex-1. Cell 97, 877887.
Toyoshima, F., Matsumura, S., Morimoto, H., Mitsushima, M. and Nishida, E.
(2007). PtdIns(3,4,5)P3 regulates spindle orientation in adherent cells. Dev. Cell
13, 796-811.
Troutt, L. L. and Burnside, B. (1988). Microtubule polarity and distribution in
teleost photoreceptors. J. Neurosci. 8, 2371-2380.
Tsujikawa, M., Omori, Y., Biyanwila, J. and Malicki, J. (2007). Mechanism of
positioning the cell nucleus in vertebrate photoreceptors. Proc. Natl. Acad. Sci.
USA 104, 14819-14824.
Turner, D., Snyder, E. and Cepko, C. (1990). Lineage-independent determination
of cell type in the embryonic mouse retina. Neuron 4, 833-845.
Vaughan, K. T. and Vallee, R. B. (1995). Cytoplasmic dynein binds dynactin
through a direct interaction between the intermediate chains and p150Glued. J.
Cell Biol. 131, 1507-1516.
Ward, S., Thomson, N., White, J. G. and Brenner, S. (1975). Electron
microscopical reconstruction of the anterior sensory anatomy of the nematode
Caenorhabditis elegans. J. Comp. Neurol. 160, 313-337.
Waterman-Storer, C. M., Karki, S. and Holzbaur, E. L. (1995). The p150Glued
component of the dynactin complex binds to both microtubules and the actinrelated protein centractin (Arp-1). Proc. Natl. Acad. Sci. USA 92, 1634-1638.
Wei, X. and Malicki, J. (2002). nagie oko, encoding a MAGUK-family protein, is
essential for cellular patterning of the retina. Nat. Genet. 31, 150-157.
Wen, G. Y., Soifer, D. and Wisniewski, H. M. (1982). The doublet microtubules
of rods of the rabbit retina. Anat. Embryol. (Berl.) 165, 315-328.
Wetts, R. and Fraser, S. (1988). Multipotent precursors can give rise to all major
cell types of the frog retina. Science 239, 1142-1145.
Whited, J. L., Cassell, A., Brouillette, M. and Garrity, P. A. (2004). Dynactin is
required to maintain nuclear position within postmitotic Drosophila
photoreceptor neurons. Development 131, 4677-4686.
Xiao, T., Roeser, T., Staub, W. and Baier, H. (2005). A GFP-based genetic screen
reveals mutations that disrupt the architecture of the zebrafish retinotectal
projection. Development 132, 2955-2967.
Zigman, M., Cayouette, M., Charalambous, C., Schleiffer, A., Hoeller, O.,
Dunican, D., McCudden, C. R., Firnberg, N., Barres, B. A., Siderovski, D. P.
et al. (2005). Mammalian inscuteable regulates spindle orientation and cell fate
in the developing retina. Neuron 48, 539-545.
DEVELOPMENT
2964 RESEARCH ARTICLE