Phylogenetic Analysis of Gene Structure and Alternative
Splicing in a-Actinins
Monkol Lek,*,1,2 Daniel G. MacArthur,1,2,3 Nan Yang,1,2 and Kathryn N. North1,2
1
Institute for Neuroscience and Muscle Research, The Children’s Hospital at Westmead, Sydney, NSW, Australia
Discipline of Paediatrics and Child Health, Faculty of Medicine, University of Sydney, Sydney, NSW, Australia
3
Wellcome Trust Sanger Institute, Hinxton, United Kingdom
*Corresponding author: E-mail: [email protected].
Associate editor: Dan Graur
2
Abstract
Key words: a-actinin, exon duplication, alternate splicing.
Introduction
The a-actinins are an ancient family of actin-binding proteins that play a key role in the maintenance and regulation
of the cytoskeleton (Blanchard et al. 1989). The a-actinins
have homologues in slime mold (Witke et al. 1986), fungi
(Wu et al. 2001), and metazoans but, surprisingly, are not
present in plants. Among the metazoans, vertebrates
possess four a-actinin genes (ACTN 1–4) postulated to
arise from a single invertebrate ancestral gene (Virel and
Backman 2004). Both of the nonmuscle a-actinins,
a-actinins-1 and -4, are ubiquitously expressed and function as cytoskeletal proteins. a-Actinin-4 has unique functions in kidney tissue (Weins et al. 2007) and has been
implicated in cancer invasion, whereas a-actinin-1 is
highly expressed at focal adhesions and adherens junctions
(Honda et al. 1998). The skeletal muscle a-actinins,
a-actinin-2, and a-actinin-3, are highly expressed in muscle
where they act as major structural components of the
contractile apparatus at the Z-line (Beggs et al. 1992).
The a-actinin family belongs to the larger superfamily of
spectrin proteins that includes spectrin, dystrophin, and
utrophin (Djinovic-Carugo et al. 2002). A general domain topology is conserved within the a-actinin gene family, which
includes an actin-binding domain composed of a CH1 and
CH2 domain, a rod domain composed of four spectrin-like
repeats in metazoans and two EF-hand domains (MacArthur
and North 2004). The actin-binding and EF-hand domains
are highly conserved, whereas the rod domain is variable
in sequence and the number of spectrin-like repeats
(Virel and Backman 2007). In addition to the identified
functional domains, a-actinin contains a PDZ consensus
binding site that mediates interaction with proteins from
the PDZ/LIM family (von Nandelstadh et al. 2008).
There are two known sites of alternative splicing within
a-actinin that serve to modify the sequence contained in
the actin-binding and EF-hand domains, corresponding to
human exons 8 and 19, respectively. In each case, the alternate exons are almost identical, suggesting they were
created by tandem exon duplication followed by relatively
minor sequence divergence. The duplicated exons at each
site are either of the same size, allowing for slight variation
in protein function through amino acid changes, or of different size, allowing for the inactivation of specific exon
functions (Kondrashov and Koonin 2001). In the actinbinding domain, Drosophila is able to create muscle, nonmuscle, and larval muscle tissue isoforms through mutually
exclusive splicing (Roulier et al. 1992). The exon involved in
creating these variants is orthologous to human exon 8. In
humans, alternative splicing in ACTN2 exon 8 creates
a brain-specific isoform (Machuca-Tzili et al. 2006). Despite
the identification of exon 8 alternative splicing, little is
known regarding its functional significance. In contrast,
the alternative splicing of unequally sized variants of exon
19 in vertebrates is known to affect the binding of calcium
at the EF-hand, thus creating calcium-sensitive and
-insensitive isoforms of a-actinins-1 and -4 (Waites et al.
1992), whereas a-actinins-2 and -3 have lost this alternative
splicing and only express the calcium-insensitive isoform.
© The Author 2009. Published by Oxford University Press on behalf of the Society for Molecular Biology and Evolution. All rights reserved. For permissions, please
e-mail: [email protected]
Mol. Biol. Evol. 27(4):773–780. 2010 doi:10.1093/molbev/msp268
Advance Access publication November 6, 2009
773
Research article
The a-actinins are an important family of actin-binding proteins with the ability to cross-link actin filaments when in
dimer form. Members of the a-actinin family share a domain topology composed of highly conserved actin-binding and
EF-hand domains separated by a rod domain composed of spectrin-like repeats. Functional diversity within this family has
arisen through exon duplication and the formation of alternate splice isoforms as well as gene duplications during the
evolution of vertebrates. In addition to the known functional domains, a-actinins also contain a consensus PDZ-binding
site. The completed genome sequence of over 32 invertebrate species has allowed the analysis of gene structure and exon–
gene duplication over a diverse range of phyla. Our analysis shows that relative to early branching metazoans, there has
been considerable intron loss especially in arthropods with few cases of intron gains. The C-terminal PDZ-binding site is
conserved in nearly all invertebrates but is missing in some nematodes and platyhelminths. Alternative splicing in the
actin-binding domain is conserved in chordates, arthropods, and some nematodes and platyhelminths. In contrast,
alternative splicing of the EF-hand domain is only observed in chordates. Finally, given the prevalence of exon duplications
seen in the actin-binding domain, this may act as a significant mechanism in the modification of actin-binding properties.
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Lek et al. · doi:10.1093/molbev/msp268
This calcium sensitivity regulates the ability of a-actinin to
bind to actin by decreasing binding affinity as calcium concentration increases (Noegel et al. 1987). Although the
exon 19 splicing is typically mutually exclusive, both exons
are included in the rat-brain isoform (MacArthur and
North 2004) without causing a frameshift as both exons
are multiples of 3nt in length.
The recent genome sequencing of over 32 invertebrate
species from the phyla Arthropoda, Chordata, Annelida,
Mollusca, Nematoda, Cnidira, Placozoa, and Echinodermata
has allowed us to perform an extensive phylogenetic study
that includes an analysis of gene structure, sequence conservation, and exon/gene duplications. In addition, this
analysis provides the opportunity to determine whether
alternative splicing within the highly conserved actinbinding and EF-hand domains is a feature common to
all metazoan species.
Methods
Identification and Annotation of a-Actinin Genes
The genes for all 32 invertebrates were obtained by TBlastN
(Altschul et al. 1990) searches against their respective databases (table 1). In most cases, the a-actinin gene was fully
contained within a supercontig (or scaffold), and segment
pairs were in close proximity to each other. The gene
spanned over two or three supercontigs for the following
species: Apis mellifera, Nasonia vitripennis, Culex pipiens
quinquefasciatus, and Acyrthosiphon pisum. The exons
and exon boundaries were identified from the TBlastN output as high-scoring segment pairs or gaps within these segment pairs. In addition, exons were checked for correct
ordering and strand. Adjacent exons with high pairwise
protein identity were classified as tandem exon duplications. All hits were then manually analyzed for splice acceptor and donor sites to ensure the correct exon–intron
boundaries. Finally, multiple sequence alignment using
MAFFT was performed to ensure there were no gaps.
Multiple Sequence Alignments
The program ClustalW2 (Thompson et al. 1994) with default settings was used to perform multiple sequence alignment to use as input for PHYML and exon 8 alignments.
Building Trees
The phylogenetic trees based on protein sequences were
generated using the maximum likelihood method employed
by PHYML (Guindon and Gascuel 2003) using a Jones,
Taylor and Thornton model with an estimated proportion
of invariable sites and bootstrapping (1,000 replicates).
Results
Identification and Annotation of a-Actinin Genes
The a-actinin genes were identified using TBlastN against
the corresponding genome sequence using the Drosophila
melanogaster (NP_477484) or Caenorhabditis elegans
(NP_506127) a-actinin as the query sequence. The metazoan species analyzed included 12 Arthropods, 2 Annelids,
3 Platyhelminths, 7 Nematodes, 3 Chordates, 1 Echino774
Table 1. The a-Actinin Sequences Were Obtained Using TBlastN
Searches Across the Genomes of the Various Species Obtained
from Their Respective Sources.
Species
Species Abbreviation
Choanoflagellatea
Monosiga brevicollis
Mbre
Cnidaria
Nematostella vectensis
Nvec
Placozoa
Trichoplax adhaerens
Tadh
Annelida
Capitella sp I
Ccap
Helobdella robusta
Hrob
Mollusca
Lottia gigantea
Lgig
Aplysia californica
Acal
Platyhelminthes
Schmidtea mediterranea
Smed
Schistosoma mansoni
Sman
Echinococcus multilocularis
Emul
Nematoda
Pristionchus pacificus
Ppac
Heterorhabditis bacteriophora
Hbac
Trichinella spiralis
Tspi
Haemonchus contortus
Hcon
Strongyloides ratti
Srat
Brugia malayi
Bmal
Caenorhabditis elegans
Cele
Arthropoda
Daphnia pulex
Dpul
Pediculus humanus corporis
Phum
Bombyx mori
Bmor
Tribolium castaneum
Tcas
Nasonia vitripennis
Nvit
Acyrthosiphon pisum
Apis
Apis mellifera
Amel
Drosophila melanogaster
Dmel
Anopheles gambiae
Agam
Aedes aegypti
Aaeg
Culex pipiens quinquefasciatus
Cqui
Rhodnius prolixus
Rpro
Echinodermata
Strongylocentrotus purpuratus
Spur
Chordata
Branchiostoma floridae
Bflo
Ciona savignyi
Csav
Saccoglossus kowalevskii
Skow
Source
DOE JGI
DOE JGI
DOE JGI
DOE JGI
DOE JGI
DOE JGI
Broad
WUGSC
Sanger
Sanger
WUGSC
WUGSC
WUGSC
Sanger
Sanger
TIGR
UCSC
DOE JGI
NCBI WGS
SilkDB
NCBI WGS
NCBI WGS
NCBI WGS
NCBI WGS
UCSC
NCBI WGS
NCBI WGS
NCBI WGS
WUGSC
UCSC
DOE JGI
Broad
Baylor
NOTE.—DOE JGI 5 Department of Energy Joint Genome Institute, Broad 5 Broad
Institute, WUGSC 5 Washington University Genome Sequencing Centre, UCSC 5
University of California Santa Cruz Genome Browser, SilkDB 5 Silkworm Database,
Sanger 5 Sanger Institute, Baylor 5 Baylor College of Medicine Human Genome
Sequencing Centre, and TIGR 5 J. Craig Venter Institute.
derm, 2 Molluscs, 1 Cndiarian, and 1 Placozoan (table 1).
The choanoflagellate Monosiga brevicollis was used as an
outgroup for subsequent phylogenetic analysis. The resulting TBlastN output was then manually annotated using
consensus intron–exon splice junction donor and acceptor
sequences to correctly identify the exon boundaries. The
highly variable exon 1 made the correct identification of
its boundaries ambiguous, and this exon was thus excluded
from subsequent analyses. Only partial sequences for
the following species could be identified; A. mellifera, N.
vitripennis, C. p. quinquefasciatus, Schistosoma mansoni,
Echinococcus multilocularis, Pristionchus pacificus, Haemonchus
contortus, Aplysia californica, and Ciona savignyi.
Phylogenetic Analysis of a-actinins · doi:10.1093/molbev/msp268
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FIG. 1. Maximum likelihood tree generated from complete a-actinin protein sequences (excluding exon 1). Numbers at the branch points
represent the bootstrap values from 1,000 replicates.
Phylogenetic Analysis of a-Actinin Genes
The phylogenetic tree generated included all invertebrates
in which full sequences (excluding exon 1) could be obtained, and variant 8a was used where applicable. The tree
generated reflects in general the phylogenetic relationship
between the species (fig. 1). Low bootstrap values (,400)
at several branch sites can be explained by branching
events that occurred close together relative to the overall
divergence time, and therefore, these branching events
cannot be estimated with high confidence. Overall,
protein-sequence identity and similarity were high displaying at least 63% pairwise similarity. The four spectrin-like
repeats between the actin-binding domain and EF-hand
domains account for the majority of the protein-sequence
variation and long branch lengths observed.
Conservation of a-Actinin Introns
Analysis of intron positions within the a-actinin gene
revealed that the Choanoflagellate M. brevicollis was
intron-poor, whereas the early branching metazoans
Nematostella vectensis and Trichoplax adhaerens are intronrich (fig. 2). Considerable variability in intron loss has dominated in the arthropods, whereas other invertebrate phyla
havemaintainedthemajorityoftheintronspresentintheearly
branching metazoans. Overall, there appear to be few intron
gains with the silkworm, Bombyx mori, harboring the most.
Conservation of Alternatively Spliced Exons
The known splice variants in a-actinin can be split in two
groups, those generated by mutually exclusive alternate
splicing of exons orthologous to the human a-actinin exon
8 or exon 19. Variants were labeled based on their position
relative to surrounding exons; variant ‘‘a’’ is closer to the
preceding exon, whereas variant ‘‘b’’ is closer to the following exon. Exons 8a and 8b are conserved in all arthropods
but are not conserved in all nematodes and platyhelminths.
These exons differ in two conserved sites (fig. 3a), position
19, which is a cysteine in 8a and serine in 8b, and positions
28 or 29, which creates a charge change between 8a and 8b.
Interestingly Ciona savignyi has three variants created by
duplicating exon 8 twice instead of just once. In addition,
the 8a and 8b variants are conserved in vertebrate ACTN4,
and a pattern of species-specific loss was observed for
ACTN2 and ACTN1 (fig. 5). The switch between cysteine
and serine has been conserved in a-actinin-4; however,
compared with arthropods, the exon order in ACTN4
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FIG. 2. A scale diagram of the intron positions mapped on the a-actinin protein sequence relative to the Dmel sequence. The diagram spans
residues 39–844 with the first and last exons omitted. The green triangles represent conserved introns, and red triangles represent intron gains
based on parsimony. The human a-actinin-1 was abbreviated to Hsap. The first and last triangles on Hsap represent first and last introns,
respectively.
appears to be switched. This suggests that either the ACTN4
exon-8 duplication was independent of the arthropod duplication or a small chromosome translocation involving
exon 8 had occurred during vertebrate evolution. In contrast
to arthropods, a-actinin-2 switches between positively and
negatively charged residues at the C-terminal end (fig. 3c).
The 8c variant identified in D. melanogaster could not be
identified in any arthropod species apart from the genomes
of species within the Drosophila genus (data not shown),
whereas the variants 19a and 19b identified in vertebrates
are restricted to chordates. In addition to the known variants, the analysis identified exon duplications of exon 4 in
birds; exon 5 in Strongylocentrotus purpuratus; exon 6 in
molluscs, annelids, and birds; and exon 7 in Branchiostoma
floridae (fig. 5). All these exon duplicates have conserved
splice acceptor and donor sites and therefore can potentially act as splice variants.
Identification of Three a-Actinin Genes in
Helobdella robusta
All invertebrates annotated in this study have only one
a-actinin gene except for the annelid H. robusta, which
had three a-actinins. A pairwise comparison was performed
at the DNA-sequence level to rule out the possibility that
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the additional paralogues are in fact errors in genome assembly. At the protein level, the three genes are approximately 75% similar, a difference comparable with the
vertebrate a-actinin orthologues. Interestingly, an inspection of the exon 8 region of H. robusta, actinin genes revealed residue differences typically seen in variants 8a
and 8b common to arthropods and nematodes (fig. 3b).
These results suggest that H. robusta has taken the route
of gene duplication rather than alternate splicing to increase sequence diversity in its a-actinin gene. In addition,
the observation that each gene duplicate can only express
one of the exon 8 variants fits well with the subfunctionalization model. The subfunctionalization model involves
the degeneration of different functions in each of the duplicates such that both duplicates are required to complement
the functions of the preduplicate gene (Force et al. 1999). In
H. robusta, degeneration of functions mediated by exon
8a or 8b was observed in a-actinin duplicates. However,
Capitella sp I, another annelid, has only one a-actinin gene
and no variants created by alternative splicing of exon 8,
suggesting that H. robusta may be an exception among annelids. Besides the three actinin genes observed in H. robusta,
we were able to identify possible tandem gene duplications
in N. vectensis, Daphnia pulex, and B. mori. However, on
closer inspection, the gene duplicates from N. vectensis
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Phylogenetic Analysis of a-actinins · doi:10.1093/molbev/msp268
FIG. 4. Maximum likelihood tree generated from the largest
overlapping region (last 156 residues) of partial a-actinin protein
sequences from nematodes and platyhelminthes. The arrows
indicate where the PDZ-binding site was lost from the C-terminal
of a-actinin. Numbers at the branch points represent the bootstrap
values from 1,000 replicates.
Loss of PDZ-Domain Consensus Site in
Nematodes and Flatworms
FIG. 3. Multiple sequence alignment of variants a and b of exon 8.
(A) Alignment of Arthropod exon 8 showing conserved differences
at positions 19 and 28–29. (B) Alignment of non-Arthropod
invertebrate exon 8. (C) Alignment of human exon 8 variants and
a-actinin paralogues (ACTN1–4).
An analysis of the C-terminal of actinin revealed a highly
conserved class I PDZ-binding site (S/T-X-w-COOH), where
X is any amino acid and w is hydrophobic (Stiffler et al.
2007). The consensus sequence is not present in either
yeast or slime mold a-actinin, first emerging prior to the
origin of metazoans (based on its presence in M. brevicollis)
and subsequently conserved throughout the metazoan lineage. The nematodes, however, have lost the last three
amino acids and remain an exception to this conservation.
The phylogenetic tree (fig. 4) suggests the loss of this consensus site occurred after the divergence from arthropods.
Furthermore, the nematode sequences available suggest
this loss is confined to the class Secernentea. Similarly,
the loss in platyhelminths also appears to be confined
to a class, suggesting the loss of this consensus site occurred
independently twice in evolution.
Discussion
and D. pulex are almost identical at the DNA-sequence
level, a finding that could be explained either by genome
assembly errors or very recent gene duplication. In contrast,
the B. mori gene duplicate has considerable divergence at
the DNA-sequence level and therefore is more likely to be
due to a gene-duplication event followed by degeneration.
In support of this finding, B. mori is known to have more
genes compared with Drosophila (Xia et al. 2004).
The a-actinin gene of the closest living relative to metazoans, M. brevicollis is intron-poor, whereas the early metazoan, N. vectensis is intron-rich. This suggests that either an
ancient eukaryotic ancestor was intron-rich and there was
massive intron loss in M. brevicollis or there was a massive
intron invasion during early metazoan evolution. Given
that the a-actinin genes of M. brevicollis and Dictyostelium
discoideum are intron-poor, the most parsimonious
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FIG. 5. Tandem exon duplication across animal a-actinin genes.
Duplication within the actin-binding domain (exons 1–8) is present
in all animals, whereas duplication of exon 19 within the EF-hand
domain is restricted to chordates. Duplicated exons were
considered degenerated if splicing acceptor and donor sites were
lost and/or contained a stop codon.
scenario is the latter. The majority of the introns in vertebrate a-actinins have been conserved from N. vectensis,
whereas massive intron loss is observed in some arthropods. In general, N. vectensis genes showed high conservation of introns with vertebrates, but there was considerable
intron loss in Drosophila (Putnam et al. 2007). There has
been little intron gain within the a-actinin gene with
the largest number observed in the arthropod B. mori. This
may be expected as compared with D. melanogaster genes,
B. mori genes are larger with more exons, possibly due to an
increase in transposable elements (Xia et al. 2004).
The interaction between a-actinin and the PDZ/LIM
proteins Actinin-associated LIM protein (ALP) and Z-band
alternatively spliced PDZ-motif protein (ZASP) is well characterized in mouse (Pashmforoush et al. 2001; Zhou et al.
2001), C. elegans (Han and Beckerle 2009), and D. melanogaster (Jani and Schock 2007). This interaction is mediated by two interaction sites, between the ZM motif and
the a-actinin spectrin repeats and between the PDZ domain and the last three residues of a-actinin (Klaavuniemi
et al. 2004; Klaavuniemi and Ylanne 2006). Both interaction
sites in isolation can colocalize to a-actinin; however, only
mutations in the PDZ domain disrupt localization to
a-actinin (Zhou et al. 2001). Ablation of ZASP appears to
affect structural integrity and maintenance at the muscle
Z-lines (Zhou et al. 2001; Jani and Schock 2007). Surprisingly,
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yeast two hybrid (Li et al. 2004) and colocalization (Han and
Beckerle 2009) result from the nematode C. elegans suggest
that the interaction between ALP and a-actinin is still maintained despite the loss of the PDZ-binding site. Therefore,
either the ZM motif is maintaining this interaction and/or
the PDZ domain is interacting with the spectrin-like repeats
that was originally reported (Xia et al. 1997).
The a-actinin variants 8a and 8b are conserved among all
arthropods and some nematodes and platyhelminths. In addition, these variants are also conserved in a subset of vertebrate a-actinin paralogues (fig. 5). The identification of
variant 8b in ACTN4 is a novel finding and may have therapeutic applications, because mutations in the 8a variant
exon cause a rare kidney disease (Kaplan et al. 2000). Therefore, strategies designed to promote replacement of the 8a
exon with the 8b variant may act to ameliorate the severity
of this disease, assuming that the 8a exon does not encode
essential kidney-related functions. In addition, the design of
the K255E Actn4 knock-in mouse involved the insertion of
a neomycin cassette 400 bp after exon 8a (Yao et al. 2004).
This is unlikely to disrupt exon 8b splicing as it lies approximately 1.5 kb after exon 8a, but it nonetheless highlights the
importance of identifying possible functional regions in introns before disrupting them to create mouse models. The
human ACTN2 8a variant is a brain-specific isoform; however, it is not conserved in all species and in particular mouse
and rat (fig. 5). This suggests that studies of a-actinin-2 in
cells isolated from rat hippocampus cells (Wyszynski et al.
1997) may not be accurate models for human biology.
The mutually exclusive alternative splice forms of exon 8
(8a and 8b) modify the C-terminal sequence of the CH2
domain and may function as a way of altering actin-binding
affinity and regulation. There are several observations that
support this hypothesis. First, the CH1 domain is essential
for actin binding, whereas the CH2 domain is thought to
act as a modifier for actin binding (Gimona et al. 2002).
Second, mutations in exon 8 cause a structural kidney disease, focal segmental glomerulosclerosis (FSGS) (Kaplan
et al. 2000), suggesting exon 8 is important for normal function. In addition, the Actn4 K255E mutation within exon 8
associated with FSGS results in an increased affinity for actin and loss of calcium-regulated actin binding (Weins et al.
2007). Last, in D. melanogaster, variants 8a and 8b create
a muscle and nonmuscle isoform (Roulier et al. 1992). In
vertebrate nonmuscle a-actinins, variants 19a and 19b create calcium-sensitive and nonsensitive isoforms, whereas
muscle a-actinins only encode the calcium-insensitive variant (Waites et al. 1992). Furthermore, putative alternate
splicing of exon 6 in molluscs and annelids and exon 7
in B. floridae also acts to modify the sequence of the
CH2 domain. Thus, the alternate splicing within the
CH2 domain may allow actinin to vary its actin-binding
properties. Human but not mouse brain expresses an alternate ACTN2 exon 8. Based on the observations noted
above, this alternate isoform may have functional significance for brain development and function in humans.
The 8c variant identified in D. melanogaster larval muscle tissue (Roulier et al. 1992) could not be identified in any
Phylogenetic Analysis of a-actinins · doi:10.1093/molbev/msp268
other arthropod species and was only identified in genomes of species within the Drosophila genus. The 8c variant is actually an extension of exon 8a and not another
duplication of exon 8. In addition, the correct splicing of
exon 8 in D. melanogaster is dependent on muscleblind
(Machuca-Tzili et al. 2006). Therefore, it is possible that
the 8c variant is an incorrectly spliced exon 8a that has coincidently conserved the reading frame and may be considered as just noise in the splicing machinery (Melamud and
Moult 2009).
In summary, there are two known regions of alternate
splicing (exon 8 and exon 19) in a-actinin genes that have
both been created by exon-duplication events. This is evident by comparing the high protein-sequence similarity
and conservation of intron phase between the two exons.
In addition, we have identified more regions of exon duplication that have intact splice acceptor and donor sites and
are therefore likely to create additional splice variants. Variants 6a and 6b are conserved in molluscs and annelids and
in the draft genomes of chicken and zebra finch (fig. 5).
Interestingly, all exon duplications discovered among invertebrates are confined to the actin-binding domain,
and none were detected in the four spectrin-like repeats.
This suggests that exon duplications may act primarily as
a mechanism to alter actin-binding properties.
The mechanisms of exon duplication and gene duplication, which result in alternative splicing and expanded
gene families, respectively, allow for variation in gene
function without compromising existing functions. Our
analysis has shown that the invertebrate a-actinins have
typically achieved this variation in function through exon
duplication within a single a-actinin gene, allowing for
tissue-specific alternative splicing. In contrast, vertebrate
a-actinins have four gene duplicates and alternate splice
isoforms, thus permitting richer variation in gene function
and expression.
Acknowledgments
We would like to thank the various sources: US Department of Energy Joint Genome Institute, The Genome Center at Washington University School of Medicine in St
Louis, The Wellcome Trust Sanger Institute, The Broad Institute, The Human Genome Sequencing Center at Baylor
College of Medicine, and the J. Craig Venter Institute (outlined in table 1) for making unpublished sequence data
available for our analysis.
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