Published online before print
February 12, 2003, 10.1101/gr.490303
Vol 13, Issue 3, 369-381, March 2003
LETTER
Segmental Duplications in Euchromatic Regions of Human Chromosome 5: A Source of Evolutionary Instability and Transcriptional Innovation
Anouk Courseaux1,5,
Florence Richard2,3,
Josiane Grosgeorge4,
Christine Ortola1,
Agnes Viale1,6,
Claude Turc-Carel4,
Bernard Dutrillaux2,
Patrick Gaudray4 and
Jean-Louis Nahon1,7
1Institut de Pharmacologie Moléculaire et Cellulaire
Unité Mixte de Recherche-Centre National de la
Recherche Scientifique, 06560 Valbonne, France; 2Unité
Mixte de Recherche-Centre National de la Recherche Scientifique,75231
Paris cedex 05, France; 3Université
Versailles-Saint-Quentin, 78035 Versailles, France;4
Unité Mixte de Recherche-Centre National de la
Recherche Scientifique, 6549, Faculté de Médecine,
06107 Nice cedex 2, France
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ABSTRACT
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Recent analyses of the structure of pericentromeric and subtelomeric
regions have revealed that these particular regions of human
chromosomes are often composed of blocks of duplicated genomic segments
that have been associated with rapid evolutionary turnover among the
genomes of closely related primates. In the present study, we show that
euchromatic regions of human chromosome 5—5p14, 5p13, 5q13,
5q15–5q21—also display such an accumulation of segmental
duplications. The structure, organization and evolution of those
primate-specific sequences were studied in detail by combining in
silico and comparative FISH analyses on human, chimpanzee, gorilla,
orangutang, macaca, and capuchin chromosomes. Our results lend support
to a two-step model of transposition duplication in the euchromatic
regions, with a founder insertional event at the time of divergence
between Platyrrhini and Catarrhini (25–35 million
years ago) and an apparent burst of inter- and intrachromosomal
duplications in the Hominidae lineage. Furthermore,
phylogenetic analysis suggests that the chronology and, likely,
molecular mechanisms, differ regarding the region of primary
insertion—euchromatic versus pericentromeric regions. Lastly, we show
that as their counterparts located near the heterochromatic region, the
euchromatic segmental duplications have consistently reshaped their
region of insertion during primate evolution, creating putative mosaic
genes, and they are obvious candidates for causing ectopic
rearrangements that have contributed to evolutionary/genomic
instability.
[Supplemental material is available online at
www.genome.org. The following individuals kindly provided reagents,
samples, or unpublished information as indicated in the paper: D.
Le Paslier, A. McKenzie, J. Melki, C. Sargent, J. Scharf and
S. Selig.]
There is compelling evidence that gene duplication and transposition
events have played essential roles during vertebrate
evolution (Brosius 1999a ,b; Patthy 1999; Long 2001; Friedman and Hughes
2001a; McLysaght et al. 2002). However, the importance of
such evolutionary mechanisms is not restricted to ancient times. The
initial sequencing and analysis of the human genome, in combination
with previously published reports, have revealed that our genome is
constituted of a remarkably complex pattern of both ancient and recent
duplications (Lander et al. 2001; Bailey et al. 2002a). It is now
estimated that  5% (probably more) of our genetic material is
composed of duplicated genomic segments that have emerged during the
past 35 million years of primate evolution. These blocks (termed
segmental duplications) range in size from a few kilobases to hundreds
of kilobases, and share a high degree of sequence identity (>90%). In
contrast to whole-genome polyploidization events, segmental
duplications have originated from the duplicative transpositions of
small portions of chromosomal material, often containing intron-exon
structure of known genes, and tend to be localized in pericentromeric
and subtelomeric regions (for review, see Eichler 2001; Samonte and
Eichler 2002). Preliminary analyses indicate that these particular
regions of the genome have experienced extraordinary rates of
evolutionary turnover, which result in considerable structural change
and rapid gene innovation in the genomes of man and great apes (for
review, see Nahon 2002; Samonte and Eichler 2002).
In earliest studies (Viale et al. 1998, 2000; Courseaux and Nahon
2001), we have seen that recently duplicated segments of the human
chromosome 5 (HSA5) were harboring two novel chimeric genes
PMCHL1 and PMCHL2, whose mRNA or protein have been
exapted (Gould and Vrba 1982) into a new functional role during the
last stages of primate evolution. These genes were shown to have been
created by a complex mechanism of exon shuffling through
retrotransposition of an antisense MCH-messenger RNA (AROM mRNA)
coupled to de novo creation of splice sites, followed by a last event
of duplication involving a large chromosomal region (Courseaux and
Nahon 2001). In the course of the characterization of the
PMCHL genes, we established that PMCHL2 was located
on chromosome band 5q13, proximal to the predisposition locus of the
spinal muscular atrophy (SMA)—an autosomal recessively inherited
neurodegenerative disorder with variable clinical severity (Munsat et
al. 1990; Viale et al. 2000). Interestingly, this region of the human
genome was shown to harbor a class of transcribed
pseudogenes/gene-derived sequences that are similar in their
organization to the PMCHL genes (Sargent et al. 1994;
Theodosiou et al. 1994; Selig et al. 1995). These sequences are not
typical processed pseudogenes, but rather contain introns and exons
with related copies on both the q and p arms of human chromosome 5, and
show at least 90% nucleic acid sequence identity to functional genes
located elsewhere in the genome. A preliminary PCR analysis of the DNA
of different primate species revealed that those sequences have emerged
during the last 35 million years (A. Viale and C. Ortola, unpubl.).
Such a clustering of recent transcribed pseudogenes in duplicated
genomic regions is reminiscent of the characteristics of
primate-specific segmental duplications (Bailey et al. 2002b; Crosier
et al. 2002) and raise questions about when these sequences have
arisen, how they were maintained, and the role that they may have
played in chromosome and genome evolution.
To address these questions, we first focus on three types of
gene-derived sequences evidenced by different laboratories during their
attempts to identify potential candidate genes for SMA disease as
follows: (1) Psdex4 and c41-cad, two sequences derived from the
CDH12 gene, a neuronal cadherin gene residing on 5p14.3. They
were shown to be almost identical to CDH12 exons 4 and 6 and
surrounding intronic sequences, respectively (Selig et al. 1995). (2)
Glu5–10, a sequence that exhibits high-sequence similarity
to exons 5, 9, 10, and adjacent intronic sequences of the
 -glucuronidase gene (GUSB), which is located on chromosome
band 7q11.21 (Sargent et al. 1994; Theodosiou et al. 1994). By
combining FISH and in silico analyses, we reveal the series of
transposition and inter- and intrachromosomal duplication events that
have led to the creation and spreading of the GUSB and
CDH12-derived sequences. Many of the recently duplicated
segments demonstrated to date, were shown to be associated with
recurrent chromosomal structural rearrangements and generation of gene
diversity (Samonte and Eichler 2002). We thus investigate the potential
of human chromosome 5 euchromatic segmental duplications (1) to be
involved in the chromosomal/evolutionary instability associated with
the predisposition locus of the SMA disease (Lewin 1995), and (2) to
evolve novel transcripts by examining the duplicated regions for
evidence that a transposition and/or duplication event has generated a
new transcript.
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RESULTS
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Clustering of Gene-Derived Sequences in Euchromatic Regions of HSA5
To first determine the precise chromosomal localization of all the
paralogous copies of the gene segments, we used a combination of mono-
and dual-color FISH experiments on human metaphase chromosomes. This
analysis revealed that the CDH12- and GUSB-derived
sequences were present in several copies on different loci of the human
chromosome 5, and that they were colocalized with PMCHL1 and
PMCHL2 on both 5p14 and 5q13 (Fig.
1A,B). Interestingly, the hybridization
signals obtained with c41-cad and Glu 5–10 on chromosome 5 were shown
to vary between individuals (data not shown). These observations
suggest that the copy number of the duplicated fragments on chromosome
5 is highly polymorphic, as proposed previously (Sargent et al. 1994;
Theodosiou et al. 1994).
Emergence and Spreading of Gene-Derived Sequences During Primate Evolution
To provide further insights into the evolutionary mechanisms that
have generated such a genomic diversity and gene-segment clustering, we
extended our FISH analysis to other primates. The metaphase chromosomes
of six species were compared (Homo sapiens [HSA], Pan
troglodytes [PTR], Gorilla gorilla [GGO], Pongo
pygmaeus [PPY], Macaca sylvana [MSY], and Cebus
capucinus [CCA]). The FISH hybridization signals suggested that
the mechanism responsible for the emergence and spreading of the
gene-variant sequences comprised at least two distinct steps (Fig.
2). First, transposition events from the
ancestral chromosomal regions equivalent to HSA12q23 and HSA7q21.11
have led to the insertion of PMCHL1 and Glu 5–10, respectively, on
the ancestral chromosomal segment equivalent to the short arm of HSA5,
25–35 million years ago (Mya) before the divergence of
Catarrhini. This was followed by a burst of duplication
events, which operated in the Hominidae lineage within the
last 5–10 million years, and which led to their distribution/expansion
onto various loci of the ancestral chromosome equivalent to HSA5. The
history of the spreading of the CDH12-derived sequences is
quite different, as the founder event was more likely to be operated
only at the time of divergence of humans and African great apes, about
10 Mya. Although we could not firmly exclude the presence of copies of
Psdex4 and c41-cad on the equivalent of HSA5p in PPY, MSY, and CCA, we
hypothesized that the hybridization signals observed were due to
cross-hybridization with the homologous functional CDH12 gene
that is present on the band equivalent to HSA5p14 (Selig et al. 1997).
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Figure 2. Summary of the in situ hybridization results of the gene-derived
sequences/SMA genes on human and other primate chromosomes equivalent
to human chromosome 5 (HSA5). The scale indicates the generally
accepted times of divergence of Anthropoids from the human lineage
(Hacia 2001). On the basis of the phylogenetic classification of
Goodman (1999), we placed Pongo in the family
Pongidae and grouped Gorilla, Pan, and
Homo in the family Hominidae. The same color code as
in Figure 1 was used. The number of dots indicates the average signal
intensity observed at a given location with a particular clone. Red
dots on Psdex4 and c41-cad hybridization patterns indicate
cross-hybridization with the genuine CDH12 gene. The names and
chromosomal localizations of the original genes, whose gene segments
derived from are indicated at bottom. The yellow
bar indicates hybrization signals obtained with two genes (SMN
and NAIP) specific to the SMA locus. The position of the
homologs to human SMA locus is indicated as a chromosomal index of the
region equivalent to HSA 5q13. Brackets indicate the homology between
HSA and other primate chromosomes. For Cebus capucinus (CCA),
CCA1 is formed by the equivalent of the whole HSA5, plus a small
segment of HSA7 (Richard et al. 1996). For Macaca sylvana
(MSY) and Pongo pygmaeus (PPY), the equivalent to HSA5 (MSY5
and PPY4, respectively) have a banding pattern very similar to HSA5
(Dutrillaux 1979). For Gorilla gorilla (GGO), GGO4 and GGO19
are the products of a reciprocal translocation between ancestral
chromosomes equivalent to HSA5 and HSA17, respectively (Dutrillaux et
al. 1973). Evolutionary breakpoints occurred in regions equivalent to
HSA5q13.3 and HSA17p12 (Stankiewicz et al. 2001). Brackets indicate the
homology between GGO and HSA chromosomes. For Pan troglodytes
(PTR), PTR4 differs from HSA5 by a pericentromeric inversion,
inv(5)(p14.3;q13.3) (Yunis and Prakash 1982; Marzella et al. 1997).
Grey lines on PTR chromosomes indicate the pericentromeric inversion
breakpoints.
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To further address the question of the original emergence of the
CDH12-derived sequences, we performed an in silico analysis of
the genomic structure of the human CDH12 gene. A BLAST search
against the HTGS and nr databases of GenBank allowed us to identify
several clones in HTGS phase encompassing the CDH12 locus. The
analysis of the draft sequence revealed that the neuronal cadherin gene
spanned a larger region than suspected previously (>1 Mb) (Selig et
al. 1997). Interestingly, both PMCHL1 and a
GUSB-derived sequence were detected inside
CDH12-intronic sequences in the vicinity of corresponding
exons 4 and 6, respectively. PMCHL and CDH12-exon
4-derived sequences linkage was established by both (1) PCR
amplification from BAC clones bearing either the PMCHL1 locus
on 5p14 (CIT-HSP283L20) or the PMCHL2 locus on 5q13
(CIT-HSP811M22, CIT-HSP 484D2), which we analyzed previously (Courseaux
and Nahon 2001), and (2) sequence comparison between clones
AC091956/AC092358 and AC108106 in HTGS phase-specific 5p and 5q loci,
respectively. This analysis led us to conclude that both
PMCHL2 and Psdex4 have arisen, 5–10 Mya (Fig. 2) through an
intra-chromosomal duplication event that involved a large chromosomal
region equivalent to HSA5p14, encompassing more than 100 kilobases.
Similarly, the link between a GUSB-derived sequence and the
exon 6 of CDH12 strongly suggested that c41-cad have arisen
through the duplication of a large genomic region equivalent to
HSA5p14. The additional copies of Psdex4 and c41 detected, particularly
on the equivalent of HSA5p in PTR and GGO, were probably generated by
further rearrangements after the divergence between African great apes
and human lineages.
To refine the evolutionary scenario responsible for the appearance and
spreading of Glu 5–10, structural analysis of GUSB-derived
paralogous sequences was performed by an in silico analysis of the
human genome draft sequence. This study revealed the existence of
several classes of unprocessed pseudogenes that share a high degree
(90%–95%) of identity with different parts of the GUSB
genomic locus (Fig. 3A). The genomic
distribution of those sequences was in agreement with our previous FISH
analysis. The Glu 5–10 probe strongly hybridized on chromosome 5
(5p14, 5p13, 5q13, 5q15) (Figs. 2 and 3B). Additional hybridizing loci
were found at 6p11 and 6p21, and further weak signals were also
detected on 22q11 and 7q11 (Fig. 3B). The chronology of emergence and
spreading of the GUSB-derived sequences was evidenced from the
chromosomal distribution of the related sequences found in the
different primate species. As illustrated in Figure 3B, the
hybridization signals on primate chromosomes suggested that after
Platyrrhini divergence—about 35 Mya—the ancestral
GUSB-genomic locus served as a donor hot spot for
transposition to the ancestral chromosomal regions homologous to HSA5p
and to pericentromeric regions of HSA7 and HSA22. According to the
present genomic organization of the GUSB-paralogous sequences
in human, we propose that initial transposition event(s) may have
involved large portion(s) of the -glucuronidase gene, parts of which
subsequently became deleted. Once the initial GUSB autosomal
copy had been transposed into a pericentromeric region, it was rapidly
used as a seed for further transposition/duplication in nonhuman
primates. On the ancestral region homologous to HSA5p, the initial
GUSB copy has taken a different evolutionary route. The first
event of insertion was followed by a period of apparent paucity in
terms of duplication of 20 million years. In contrast, the multiple
copies on the equivalents of HSA5 and HSA6 in the Hominidae
genomes (Fig. 3B) seem to be the result of a burst of intra- and
inter-chromosomal duplication events that occurred after the separation
from the ancestral Pongidae, 14 Mya.
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Figure 3. (A) Schematic representation of the genomic structure of the
-glucuronidase gene (GUSB) and GUSB-derived
paralogous sequences. The genomic structure of the GUSB gene
was established according to sequence comparison and alignment between
the -glucuronidase mRNA complete sequence (M15182) and genomic
sequence of the BAC clone RP11-252P18 (AC073261). Dark gray boxes
indicate the exons, and thick light-gray lines indicate intervening
intronic sequences. For each GUSB-derived sequence found in
the draft sequence of the human genome, the chromosomal localization
and the GeneBank ID of the clone(s) they were derived from are
indicated. (*) Clones in HTGS phase in GenBank—according to the June
2002 freeze of the genome draft sequence. Horizontal black bold lines
illustrate chromosomal separations, horizontal black plain lines,
subchromosomal separations, and horizontal black broken lines,
gene-copy separations. The bracket indicates that the same clone was
sequenced twice. (B) Summary of the in situ hybridization
results obtained with a Glu 5–10 cosmid probe on human (HSA) and other
primate chromosomes. A molecular timescale for primate evolution is
indicated. The number of dots indicates the average signal intensity
observed at a given location, resulting from the number of loci and the
hybridization efficiency. The brackets illustrate the homologies
between Macaca or Cebus and human chromosomes. In the
presumed ancestral karyotype of placental mammals, the HSA7 homolog was
composed of two parts. These two components fused before the separation
of Cercopithecoidea and Hominidae (25 Mya), and
then pericentric and paracentric inversions occurred in the
Hominidae lineage. Thus, HSA7 is a fairly recent chromosome
shared by HSA and PTR only. In Cebus capucinus, the smallest
part homolog to HSA7 is on CCA1 associated with the homolog to the
whole HSA5 (Richard et al. 1996). In Macaca sylvana, MSY2 is
formed by the equivalents of the whole HSA7 and HSA21, and MSY9 by the
equivalents of HSA20 and HSA22 (Muleris et al. 1984).
HSA22/PTR23/GGO23/PPY23 on one hand, and HSA6/PTR5/GGO5 on the other
hand, differ mostly only by heterochromatic variations (Yunis and
Prakash 1982). The differences in the chronology of expansion/spreading
of the GUSB paralogous sequences is depicted by a horizontal broken
line, top, primo-insertion in pericentromeric regions on
ancestral HSA7 and HSA22, bottom, primo-insertion in the
ancestral HSA5p euchromatic region.
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Spreading of the Gene-Derived Sequences Through a Process of Segmental Genomic Duplication
As proposed previously (Courseaux and Nahon 2001) and in this study,
the last events of duplication likely involved large genomic segments.
Sequence homology was thus expected to extend to flanking sequences
that lie between the genes-derived sequences. To confirm this
hypothesis, we expanded our comparative in silico analysis of the
genomic structure to the flanking regions of GUSB pseudogenes
residing on HSA5 and HSA6. This allowed us to identify several BAC/PAC
clones that share strong sequence similarity (>90%) over large
regions. A schematic representation of the homology extent between the
different loci is given in Figure 4.
Interestingly, the series of database searches used to characterize
these segmental duplications has led to the identification of three
other paralogous sequence variants (PSVs), MW3, P211, and FLJ. MW3
shows a high degree of identity to the THE-1 (transposable human
element) retrotransposon gene family (Francis et al. 1995). Further
FISH analysis on human and primate chromosomes suggested that MW3 was
distributed onto Hominidae chromosomes equivalent to HSA5
(represented graphically in supplemental information) and HSA6 at the
same time and place as Glu 5–10. FLJ and P211 were named on the basis
of their strong similarity to cDNAs FLJ23032fis and DKFZp434P211
(85%–92%), respectively. Both appeared highly amplified throughout
the human genome. Other FLJ-related sequences (80%–95% nucleic acid
identity) have been found in clones specific to the HSA2q14–HSA2q21,
HSA13q32, and Xq21 genomic regions, and numerous P211-related sequences
were also identified on HSA22q11 and HSA20p11 (Table
1). The respective genomic organization and
chromosomal localization of FLJ and P211-related sequences suggest that
those sequences have originated primarily on the ancestral chromosome
homologous to HSA5 through the transposition of segmental parts of
genes located elsewhere in the genome, as PMCHL and Glu 5–10
have done. Sequences between and outside of the PSVs are composed of a
mixture of highly repetitive elements (50%, mainly LINE1, Alu
repeats, and LTR elements) and putative transcripts—sequences that
display strong sequence similarity (>95%) with ESTs and Unigene
clusters as illustrated in Figure 4. The molecular structure of the
paralogous regions proved to be amazingly complex, with components
being present, duplicated, or absent among different representative
copies. When located adjacent to one another, the duplicated elements
are arranged tandemly. The molecular basis of the variation in the
extent of paralogy is unclear. Although it is possible that these
differences reflect the size variation of the genomic segments that
were further distributed, they may also be the result of secondary
events that rearranged these large blocks after duplication. The net
effect of these duplication events is the generation of large (30 to
>100 kb) blocks of paralogy among multiple nonhomologous chromosomal
loci, each of which is a mosaic of smaller duplicated segments.
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Figure 4. Paralogy map and sequence content of the HSA5/HSA6 duplicon. The
genomic organization of the duplicated region was deduced from the
AL157879 and AL133255 clones that were chosen as reference seed
sequence. The thick gray lines illustrate the homology extent between
paralogous loci. The chromosomal localization and the GeneBank ID of
the clones are indicated. The P211, FLJ, and GUSB gene-derived
sequences are boxed in dark gray. For exon-intron organization of the
GUSB-derived sequences, see Figure 3A. The THE-1 transposable
element MW3 is boxed in light gray. X, X1, and X2 are sequences
paralogous to exons of the chimeric genes described in Figure 6. Gray
arrows indicate the localization of sequences highly homologous
(>95%) to ESTs and human UNIGENE clusters as follows: Hs.186379,
Hs.312136, Hs.224604, Hs.145839, Hs.274528, Hs.297663, Hs.50454,
Hs.202243, Hs.183256, Hs.132586, AW963166, BE780832, Hs.317160,
Hs.326016, Hs.121081, Hs.294040, Hs.166361, Hs.321499, Hs.324135,
Hs.7569, Hs.131950, and Hs.186180. The brackets indicate that the same
clone was sequenced twice. (*) Clones in HTGS phase in
GenBank—according to the June, 2002 freeze of the genome draft
sequence.
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Organization of Segmental Duplications at the SMA Locus on 5q13
To study the impact of segmental duplications on the genomic
organization of the SMA region (Lewin 1995), we first analyzed the
distribution of the PSVs through a PCR analysis on a series of
overlapping YAC clones spanning the region of predispostion to the
disease (Fig. 5A). To further specify the
genomic structure of this locus, a series of BLAST similarity searches
against the nr and HTGS databases of GenBank were performed to reveal
all of the clones sequenced as part of the Human Genome Project that
contain one or more genes specific to the SMA region (SMN,
SERF1A, NAIP, or BFT2p44) (Lefebvre et al.
1995; Burglen et al. 1997; Carter et al. 1997; Scharf et al. 1998). The
searches identified 15 clones issued from 6 different genomic libraries
that were selected for subsequent sequence comparisons to provide the
best alignment (Fig. 5B). Such an analysis showed that what was
considered primarily to be an inverted duplication of some several
hundreds of kilobases (Lewin 1995), was far more complex than expected
previously. The telomeric part of the 5q duplicon that contains the
functional copies of the SMA locus genes (BTF2p44t,
NAIP, SMN1, SERF1At) was shown to display a
compact gene organization of 200 kb in length. In contrast, the
centromeric duplicon appeared to be composed of a complex patchwork of
recently duplicated genomic segments (Fig. 5B). A combination of PCR
and FISH analysis with PAC clones described previously (Roy et al.
1995) was used to confirm the chromosomal distribution and the
cytogenetic band positions of the duplicated segments (data not shown).
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Figure 5. Genomic organization of the SMA locus on 5q13. (A) Consensus
YAC contig spanning the SMA region. Genes of the locus (SMN,
NAIP, and BFTp44) are symbolized on the black line
representing the extent of the clones, and the PSVs underneath. Legend
at right. Thick gray and white lines symbolize YAC chimerism
and instability, respectively. (B) Genomic organization of the
SMA locus established through sequence analysis of PAC/BAC clones
(a–p). Despite the fact that critical pieces of information about this
chromosomal region are still missing or unclear, we succeeded in
reconstructing the sequence organization of six nonoverlapping genomic
areas encompassing the disease locus (I. to VI.). Tel. and Cen. refer,
respectively, to the telomeric and centromeric parts of the duplication
as described in the literature. Sequences specific to this duplicon are
highlighted in orange. Thick gray lines represent duplicons whose
original locus is on 5p14; (1) thick light-gray lines represent areas
paralogous to the 5p14 region, and (2) thick dark-gray lines represent
paralogous sequences distributed among several loci on human
chromosomes 5 and 6 (see Fig. 4). Arrows indicate the extent and the
5'–3' orientation of the genes. The NAIP represent deleted forms of
the NAIP gene. SMN1-6, NAIP11-17, and OCLN5-9 are deleted
forms of the SMN, NAIP, and OCLN genes,
respectively. (C161) CATT1-G1/C161 dinucleotide repeat marker (Burghes
et al. 1994). (X, X1, X2, X3, C161) Sequences paralogous to exons of
the chimeric genes depicted in Figure 6. Gene symbols and PSVs are
denoted according to the symbol used in Figures 4 and 5A. (a)
AC016554/CTC-340H12; (b) AC005031/GSP13996; (c) U80017/125D9; (d)
AC044797/RP11-195E2; (e) AC004999/D215P15; (f) AC022119/CTC-566F17; (g)
AC010272/CTC-492P2; (h) AC010237/CTC-348J20; (i) AC010217/CTC-202F24;
(j) AC093202/CTC-249C14; (k) AC044800/RP11-34J8; (l)
AC011244/RP11-497H16; (m) AC012369/RP11-551B22; (n)
AC110009/RP11-508M8; (o) AC010355/CTD-2027K22; (p) AC015470/RP11-2H18.
(*) Clones in HTGS phase in GenBank—according to the June, 2002 freeze
of the genome draft sequence.
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Segmental Duplications of HSA5: A Source of Gene Diversity?
To determine whether the recently duplicated regions of HSA5 could
contain functional genes specific to the primate lineage, we have
looked for transcripts that were likely to have originated recently
during primate evolution as the PMCHL genes did (Courseaux and
Nahon 2001). By systematic searches in databases and careful
examination of the literature, we discovered nine chimeric-processed
transcripts whose genomic organization is suggestive of a recent origin
(Fig. 6). Their creation could not predate
the duplication events that have led to the juxtaposition of exonic
sequences of which they are constituted. The first class of transcripts
(SMA3, SMA4, SMA5, NIH_MGC_19) contains exons homologous to exons 5, 9,
and 10 of the GUSB gene associated with exonic sequences,
which were shown to be present only in the duplicons specific to HSA5
and HSA6 (Fig. 4). These mosaic transcripts could lead to the fusion of
new protein modules as illustrated in Figure 6 (ORF, a–j). Another
transcript, 5CMP1, was shown to display a mosaic structure. It is
composed of (1) four exons homologous to the GUSB gene (exons
5, 9, 10, and 11), (2) an exon X3 present in the duplicon paralogous to
the HSA5p14 (see Fig. 5B), and (3) two exons, N7 and C161, homologous
to the functional NAIP gene. N7 is paralogous to exon 7 of
NAIP, and C161 was named on the basis of its strong sequence
identity to the CATT-G1/C161 multiple subloci polymorphic marker, whose
one copy lies into the intron between exons 7 and 8 of the
NAIP gene. The mapping of CATT1 suggests that the
5CMP1 gene may be transcribed from one subloci localized on
HSA5q13.1 (Burghes et al. 1994). Regarding the last class of
transcripts (CE3, CD19, CD23, CC14.1), only partial information was
available. No sequence has been deposited in GenBank, and consequently,
the sequence of the 3' part was obtained from the literature (Roy et
al. 1995). Although fragmentary, these data were sufficient to
establish that the four transcripts resulted from the fusion of
NAIP and OCLN-derived exons associated with an exon
X4, whose genomic origin remains undetermined.
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Figure 6. Schematic representation of chimeric transcripts and potentially
functional ORFs. Exonic sequences are boxed and were deduced from
comparative analyses between the DNA sequences of the transcripts and
genomic clones. Sequences highly similar (86%–100% identity) to
other known gene-exonic sequences are colored as follows: green,
GUSB derived; blue, OCLN derived; orange,
NAIP derived. These exons are numbered according to the exonic
sequences they are derived from, that is, N7, sequence paralogous to
NAIP exon 7. GUSB-derived chimeric transcripts are
drawn to scale. Regarding NAIP-OCLN-derived transcripts, only
partial information was available. The 3' part of these cDNAs was
assigned previously as NAIP exon 17 (Roy et al. 1995).
However, further in silico analysis revealed that the 3' ends actually
consisted of four exons homologous to OCLN exons 5–9, and
were associated with the exonic sequence X4, whose genomic origin
remains undetermined. (Rp) Repeated sequence as follows: (Rp.1)
LTR/pTR5; (Rp.2) LINE/HAL1-SINE/Alu Sx; (Rp.3) MER3-SINE/Alu Y; (Rp.4)
LTR/pTR5-LINE/L1. (C161) Exonic sequence containing the CATT1-G1/C161
dinucleotide repeat marker. Gray brackets numbered a–l indicate the
extent of potentially functional ORFs initiated from an ATG codon in a
reasonable initiation context (Kozak 1984). For genomic localizations
of sequences paralogous to exons C161 and X3, see Figure 5B, and for
exons X, X1, and X2, see Figures 4 and 5B.
|
|
|
DISCUSSION
|
|---|
The recent data concerning the organization of segmental
duplications in primate genomes revealed that these unusual genomic
regions represent an extraordinary source of information about several
biological processes (Eichler 2001; Samonte and Eichler 2002). They
constitute a rich palaeontological record, holding crucial clues about
evolutionary events and forces. As passive markers, they shed light on
chromosome structure and dynamics; as active agents, they have reshaped
the genome by creating entirely new genes and causing ectopic
rearrangements that may further have led to inherited pathology.
On the basis of previous analysis (Christian et al. 1999; Jackson et
al. 1999; Shaikh et al. 2000; Courseaux and Nahon 2001; Horvath et al.
2001) and the present findings, a plausible scenario for the emergence
of paralogous segments in the human lineage can be drawn. The
multiplicity of segmental gene copies in the human genome might be
explained by an intense activity of gene duplication—through
retrotransposition for PMCHL1, but yet unknown mechanisms for
the other gene models—that occurred in the ancestral genome at the
time of divergence between Catarrhini and
Plathyrrhini—about 25–35 Mya for PMCHL1 and Glu
5–10, predating Catarrhini divergence for the other gene models. Some
ancestral gene loci appeared to have acted as hot-spot donors of
sequences (e.g., the ancestral MCH/AROM and GUSB
loci). Whereas other regions of the genome appeared to have served as a
safe haven for transposons/duplicons that have escaped inactivation or
elimination—for example, ancestral HSA5p14 or pericentromeric or
subtelomeric regions of chromosomes (Horvath et al. 2000; Bailey et al.
2001). The further expansion/spreading of the segmental gene copies
occurred then by subsequent inter- and
intra-chromosomal duplication events involving large chromosomal
regions, whose chronology and mechanism should be different regarding
the region of primary insertion. We have shown that, on the ancestral
region homologous to HSA5p, a period of 20 million years, has
apparently separated the first event of insertion and the following
massive duplication events (Fig. 3B). Conversely, once transposed into
a pericentromeric region, DNA segments were shown to have been rapidly
used as a seed for further transposition/duplication. The quantitative
differences in the distribution of the GUSB-derived sequences,
as we observed among representative members of the Hominidae
(Fig. 3B), are consistent with the rapid evolutionary turnover that has
been found in the pericentromeric region of human and other primate
chromosomes (Eichler et al. 1999). Although it is possible that these
regions simply have a great tolerance for large insertions, the
possibility of specific mechanisms for recent insertion of segmental
genomic duplication has been suggested on the basis of unusual GC-rich
DNA (Eichler et al. 1999) and satellite sequences (Regnier et al. 1997;
Guy et al. 2000) flanking the insertions. The authors postulated that
these elements, in addition to serving as transposition integration
signals, may also represent focal points for the transfer of genomic
material among pericentromeric regions. The quantitative and
qualitative differences in the distribution of these duplicons shown
among representative members of human and apes, suggests that the
dynamic spread of these sequences may be a still-ongoing evolutionary
process in the Hominoid genomes.
In the last few years, intrachromosomal duplications have been
implicated in many recurrent chromosomal rearrangements associated with
microdeletion and microduplication syndromes (Lupski 1998). It is
noteworthy that many of the regions associated with intrachromosomal
rearrangements and disease have also been active for segmental
duplication events, suggesting that the processes of duplication and
genomic instability are closely intertwined and contribute to disease
etiology and genomic structural variations (for review, see Ji et al.
2000; Emanuel and Shaikh 2001). Here, we address this question in the
context of the etiology of the SMA pathology by analyzing the impact of
segmental duplications on the genomic organization of the
predisposition locus. This region of the human genome has proven to
have an highly unstable and intricate genomic structure (Lewin 1995)
and to show a marked heterogeneity in the normal population (Burghes
1997; Campbell et al. 1997). Differences in the structure of the SMA
region among human individuals are well documented. Markers and genes
that lie within this locus are represented multiple times but also may
vary in copy number even in a highly inbred population (Burghes 1997;
Campbell et al. 1997). Sequence homology between the recently
duplicated DNA segments, which we show here (Fig. 5), does provide a
chance for misalignment during meiosis, leading to unequal exchange and
chromosome rearrangements that result in the multiplication of all or
parts of genes/PSVs observed within this region. This unusual form of
polymorphism may also explain why this region is purported to be
particularly recalcitrant to YAC subcloning and the lack of correlation
between the different physical maps that have been published previously
(Lewin 1995). Human genomic libraries consist of clones from at least
two different haplotypes, making a consistent map of such a region from
mixed haplotype libraries present even more of a challenge. Finally, it
is obvious that the complex, highly duplicated nature of these regions
is not amenable to high-throughput assembly methods—that is,
restriction fingerprinting methods for determining clone
overlap—(Bailey et al. 2001). As a consequence, the current assembly
of the SMA locus was revealed to be erroneous and not able to
recapitulate the organization published in the literature. The most
common error was the merging of nearly identical sequence-duplicated
segments into a single contig. To circumvent all of the problems
inherent in physical mapping, we have used the sensitivity of sequence
data to further specify the potential implication of PSVs in mediating
the high degree of instability observed within the SMA region. Such an
analysis showed that the insertion of PSVs associated with the burst of
duplication events that occurred in the Hominidae lineage has
fully reshaped the SMA locus during primate evolution. Although a
direct connection is yet to be established, it is particularly tempting
to speculate that there is a link between the dynamic duplicative
nature of genomic duplicons and the rearrangements mediating the SMA
disease (e.g., deletions and gene conversion events). The definite
proof will come from the isolation and characterization of breakpoints
from multiple patients.
Despite its possible deleterious effects, gene duplication remains one
of the primary forces of evolutionary change. An evolutionary benefit
of such genomic plasticity may be to juxtapose different cassettes from
diverse genes to create a reservoir of gene/protein modules with a
potentially new function. In the present study, we revealed that the
evolutionary rearrangements that have participated in the origin of the
two chimeric functional genes PMCHL1 and PMCHL2
during primate evolution (Courseaux and Nahon 2001) were also involved
in the emergence and dispersal of various gene segments containing
duplicated regions throughout several loci of human chromosomes 5 and
6. It was therefore crucial to determine whether such an abundant
recently duplicated material could contain other functional genes
specific to the primate lineage. By in silico screening, we identified
numerous chimeric transcripts. A first class (SMA3, SMA4, SMA5,
NIH_MGC_19) consisted of GUSB-derived exons associated with
different combinations of exonic sequences that were shown to be
present only in the duplicon described in Figure 4 and to display no
similarity to any other expressed sequence of the GenBank databases.
This observation rendered very unlikely the hypothesis that those new
sequences might be part or duplicates of previously existing genes.
Rather, those exons should have originated from unique genomic
sequences that fortuitously evolved as standard intron-exon structures,
as we found previously for the PMCHL1 gene (Courseaux and
Nahon 2001). This led us to propose that after its insertion, about
35 Mya, the transposed GUSB gene segment has recruited through
subsequent mutations events, intronic and exonic components into
transcription unit(s). Furthermore, sequence analysis revealed that
these mRNAs did not result from alternative splicing events, but rather
suggested that they were the product of genes located on different
paralogous segments. Several ORFs were deduced from the sequences of
the SMA3, SMA4, SMA5, and NIH_MGC_19 transcripts (bracketed in Fig.
6). The analogy here, with the PMCHL genes, was
particularly striking, as these ORFs overlapped exons originating from
distinct paralogous loci and led to putative new fused proteins
domains. Another class of transcripts was found to result from the
juxtaposition of NAIP and OCLN-derived exons (Fig.
6). In the human genome, as many as six copies of truncated and/or
internally deleted NAIP loci have been detected previously
(Rajcan-Separovic et al. 1996). Our in silico analysis of the SMA locus
allowed us to analyze the genomic environment of three of them (Fig.
5B). Interestingly, the three loci were localized at the boundaries
between different duplicons. In particular, one of them, the internally
deleted form NAIP1–615–17 (Fig. 5B),
was adjacent to a 5' truncated version of the OCLN gene
(OCLN5-9). This strongly suggested that CE3, CD19, CD23, and
CC14.1 are the transcriptional products of at least one chimeric gene
created 5–10 Mya, as the result of the rearrangements at the SMA
locus. These have led to the clustering of new emerging deleted forms
of the NAIP and OCLN genes, thus supporting the
concept of exon shuffling (Gilbert 1978).
Recently, several fusion transcripts involving endogenous genes and
exonic portions of segmental duplications have been described in the
human genome. Although their biological significance is unknown, it is
notable that the transcription of all of these recent transcripts is
found mainly in germ-line, fetal, or cancerous tissues (Fig. 6; Viale
et al. 2000; Courseaux and Nahon 2001; Bailey et al. 2002b). The vast
majority of such evolutionary experiments are probably failures at a
functional level, and thus require further experimental confirmation.
However, according to the sheer abundance of transcripts that have been
created or modified through transposition and/or duplication events
during the last 35 million years—an estimate of 500–1100 (Bailey et
al. 2002b)—we can expect functional genes to arise from the hodgepodge
of segmental duplications. The identification within recently
duplicated segments of members of the PMCHL and
morpheus gene families that were shown to exhibit,
respectively, indications of exaption and of positive selection among
Hominoids, strongly support this hypothesis (Courseaux and Nahon 2001;
Johnson et al. 2001). Taken together, these results reveal one of the
most unexpected recurrent roles of chromosomal duplication during
primate evolution and support the idea that processes of duplicon
clustering joined to positional polymorphism could be a nursery for
generating gene diversity and a source of phenotypic variation among
humans as proposed previously by B. Trask, E. Eichler, and coworkers
(Linardopoulou et al. 2001; Samonte and Eichler 2002). The generation
of human-specific genes may also be related to the major quantitative
changes in gene/protein expression in the human brain by comparison
with other mammalian species, including chimpanzees, as reported
recently by S. Pääbo and coworkers (Enard et al. 2002). It is
tempting to speculate that newly created genes in Hominoids may
contribute as master genes in the control of the species- and
tissues-specific enhancement of gene activity (Nahon 2003).
Finally, whether the process of segmental duplication as described here
is a true attribute of primate genome evolution, remains to be
determined. The contribution of large genomic duplications to generate
new gene families is firmly established in nonvertebrate species, such
as Caenorhabditis elegans and yeast (Friedman and Huges
2001a). However, human and its closest relatives are the only ones to
demonstrate duplication of unrelated gene cassettes within nonrandom
regions of chromosomes. Primary analyses of the human genome draft
sequence revealed that such a high proportion of large duplications
containing mosaic genes clearly distinguishes the human genome from
other genomes sequenced to date (Lander et al. 2001). A firm answer
will probably come from the complete sequences of other vertebrates,
such as the mouse and rat. In a more general context, there is a highly
controversial debate regarding the overt contribution (or not) of
ancient whole-genome duplication (WGD) to shape vertebrate genomes
(Friedman and Huges 2001b; McLysaght et al. 2002).
Recently, a global search in the draft human-genome sequence for groups
of parologs (parologons) and a molecular clock analysis of orthologs in
fly and nematodes provided some support for the WGD hypothesis
(McLysaght et al. 2002). However, if massive transposition of large
genomic duplicons, as exemplified here during Hominidae
evolution, was an active process during early vertebrate evolution,
this might also explain recurrent duplications in the genomes of
vertebrates. Lastly, it is worth noting that transposable elements and
retrotransposons have played a major role in shaping the vertebrate
genomes (Brosius 1999b; Kidwell and Lisch 2001). Recent studies have
revealed that apart from an exceptional burst of activity of Alu
repeats peaking around 40 Mya, a fairly steady decline in transposon
activity has occurred in the Hominid lineage over the past 35–50
million years, whereas no similar decline was detected in the mouse
genome (Lander et al. 2001). Thus, it would be interesting to
investigate whether the occurence of segmental duplication and
transposon activity may be inter-related at the molecular level in
vertebrates, particularly in the Hominids.
|
METHODS
|
|---|
FISH Analysis
On human chromosomes, FISH was performed (as described previously)
(Courseaux and Nahon 2001) on metaphase chromosomes from peripheral
blood lymphocytes of seven normal unrelated individuals. For
polymorphism estimation, at least 100 metaphases per individual and per
probe were analyzed. Fluorescent images were captured using
high-resolution cooled charge-coupled device (CCD) camera C4880
(Hamamatsu). Image acquisition, processing, and analysis were performed
using the Vysis software package (Quips SmartCapture FISH).
Nonhuman metaphase spreads were obtained from cell cultures of
fibroblasts conserved in liquid nitrogen in the tissue bank of the
Institut Curie (Paris, France). Probes were labeled with biotin-11-dUTP
by nick translation (kit Mix enzyme NTL, Roche Molecular Biochemicals).
Fluorescence in situ hybridization was performed essentially as
described previously (Viale et al. 1998). Observations were performed
under an epifluorescent microscope (Microphot-FXA, Nikon) and images
were captured using a cooled CCD camera (Photometrics), and a capture
software (Quips-Smart, Vysis).
A complete list of the clones used as FISH probe and the combination of
probes used in dual-color FISH experiments are detailed in Supplemental
Table 1.
In Silico Modeling
The in silico analysis and detection of segmental duplications were
conducted through BLAST searches of GenBank against many databases in
the Web site of the National Center for Biotechnology Information of
the National Institutes of Health (www.ncbi.nlm.nih.gov/). Multiple
alignments of the DNA sequences were done using the flat query-anchored
with identities alignment view option of the BLAST program. The extent
of the homology of paralogous regions, analysis of HTGS, extension, and
completion of contigs were further performed by using the NIX tool
(www.hgmp.mrc.ac.uk/), which combines many DNA analysis programs
(GRAIL, Fex, Hexon, MZEF, Genemark, Genefinder, Fgene, BLAST [against
many databases], Polyah, RepeatMasker, and tRNAscan).
The genomic organization of the SMA locus was established by a method
in which public sequences (GenBank databases) were analyzed at the
clone level. SMA genes and PSVs were first used as anchor markers for
sequence comparison to show overlapping genomic areas encompassing the
disease locus (100% sequence identity). The extent of the overlaps was
further refined through adjacent sequence comparison.
The translation of DNA sequences to protein sequences was conducted on
the Web site of National Cancer BI of the National Institutes of Health
(www.NCBI.nlm.nih.gov/). Chromosomal localizations were established
according to both ENSEMBL (http://www.ensembl.org/) and the Mapview
resource (http://www.ncbi.nlm.nih.gov/) and/or sequence comparisons.
YACs Contig Construction
The degree of overlap between YACs is based on sequence content and
data of the literature (Roy et al. 1995). Sequence content of the YAC
clones were established by PCR experiments. Primers and PCR conditions
are described in Supplemental Table 2.
|
WEB SITE REFERENCES
|
|---|
http://www.ensembl.org/; Ensembl, a joint project
between EMBL-EBI and the Sanger Institute that produces automatic
annotation on eukaryotic genomes.
http://www.ncbi.nlm.nih.gov/; National Center for Biotechnology
Information (NCBI).
www.hgmp.mrc.ac.uk/; UK Human Genome Mapping Project Resource Centre
(HGMP-RC).
|
Acknowledgements
|
|---|
We thank M.C. Potier (ESPCI, CNRS UMR7637, Paris, France) and P.
Vernier (Institut A. Fessard, CNRS UPR 2197, Gif-sur-Yvette, France)
for critical reading of the manuscript. We thank the following
individuals for providing reagents or samples: D. Le
Paslier (UMR CNRS 8030, Evry, France), A. McKenzie (Molecular Genetics
Laboratory Children's Hospital, Ottawa, Canada), J. Melki (INSERM
EMI-9913, Universite d'Evry, France.), C. Sargent (University of
Cambridge, UK), J. Scharf (Howard Hughes Medical Institute, Boston,
USA) and S. Selig (Department of Nephrology, Rambam Medical Center,
Haifa, Israel). A.V. was supported by the ADER-PACA (CAR9312/2679;
1994–1996) and the Association Française contre les Myopathies (AFM)
(1997). A.C. was a recipient of post-doctoral fellowships from AFM
(1997) and from Association pour la Recherche contre le Cancer (ARC)
(1998–2000). We thank F. Cuzin (U470 INSERM, Nice) for the financial
support (Hoechst Marion Roussel) attributed to A.C. (2001). Supported
by grants from the AFM (ASI 1996–1998) and the Centre National de la
Recherche Scientifique (CNRS) (Programme OHLL, 2002).
The publication costs of this article were defrayed in part by payment
of page charges. This article must therefore be hereby marked
"advertisement" in accordance with 18 USC section 1734 solely to
indicate this fact.
|
Footnotes
|
|---|
5 Institut National de la Santé et
de la Recherche Médicale, U470, Centre de Biochime Parc Valrose
06108 Nice cedex 2, France; 
6 Genomic Core Laboratory,
Memorial Sloan Kettering Cancer Center, New York, New York 10021, USA.
Present addresses:
7 Corresponding author.
E-MAIL nahonjl{at}ipmc.cnrs.fr; FAX 33-493-957-708.
Article and publication are at
http://www.genome.org/cgi/doi/10.1101/gr.490303. Article published online before print in February
2003.
|
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ABSTRACT
RESULTS
-98-1 probe). The
hybridization signals observed on 5p with the c41-cad probe are due to
cross-hybridization with the genuine CDH12 gene.