Vol 13, Issue 5, 821-830, May 2003
LETTER
Genome Size Evolution in Pufferfish: A Comparative Analysis of Diodontid and Tetraodontid Pufferfish Genomes
Daniel E. Neafsey1,3 and
Stephen R. Palumbi2
1Department of Organismic and Evolutionary Biology,
Harvard University, Cambridge, Massachusetts 02138, USA;2
Hopkins Marine Lab, Stanford University,
Pacific Grove, California 93950, USA
 |
ABSTRACT
|
|---|
Smooth pufferfish of the family Tetraodontidae have the smallest
vertebrate genomes yet measured. They have a haploid genome size of
 400 million bp (Mb), which is almost eight times smaller than the
human genome. Given that spiny pufferfish from the sister family
Diodontidae and a fish from the outgroup Molidae have genomes twice as
large as smooth puffers, it appears that the genome size of smooth
puffers has contracted in the last 50–70 million years since their
divergence from the spiny puffers. Here we use renaturation kinetics to
compare the repetitive nature of the smooth and spiny puffer genomes.
We also estimate the rates of small (<400 bp) insertions and deletions
in smooth and spiny puffers using defunct non-LTR retrotransposons. We
find a significantly greater abundance of a transposon-like repetitive
DNA class in spiny puffers relative to smooth puffers, in addition to
nearly identical indel rates. We comment on the role that large
insertions may play in the evolution of genome size in these two
groups.
[The sequence data from this study have been submitted to
GenBank under accession nos. AY212336–AY212504.]
Early surveys of nuclear DNA content in eukaryotes
were baffling. Genome size differences were enormous, with protozoans
varying 5800-fold, arthropods 250-fold, fish 350-fold, algae 5000-fold,
and angiosperms 1000-fold (for review, see Gregory 2001 ), but there
appeared to be very little correlation between genome size and
organismic complexity (the "C-value paradox"). Ultracentrifugation
and DNA renaturation kinetics ultimately demonstrated that much of the
variation in genome size was not due to differences in the number of
genes, but rather in the amount of previously unknown noncoding DNA
(Britten and Kohne 1968). Although some of this DNA encodes
regulatory information (rather than proteins), much of it was found to
consist of presumably nonfunctional simple repeats of varying lengths
and random sequence (John and Miklos 1988).
Understanding the significance of this noncoding DNA requires
elucidation of the processes responsible for its maintenance.
Comparative study of genome size evolution within taxonomic groups is a
powerful tool in discovery of these regulatory processes. In this
context, the pufferfish are particularly well suited for studies of the
process of genome size evolution. Smooth pufferfish (family
Tetraodontidae) have the smallest vertebrate genomes measured to date,
with a haploid genome size of 400 million bp (Mb; Hinegardner and
Rosen 1972). Pufferfish have approximately the same complement of genes
as other vertebrates (Brenner et al. 1993), thus this curiously small
size has presumably resulted from a loss of repetitive or other
nonfunctional, noncoding DNA. Spiny pufferfish of the sister family
Diodontidae have genomes that are roughly twice as large, 800 Mb
(Hinegardner and Rosen 1972; Brainerd et al. 2001). Mola mola(Molidae), a member of the closest outgroup to these two families
of pufferfish, also has a genome size of 800 Mb (Brainerd et al.
2001). Therefore, the difference in genome size between tetraodontid
and diodontid puffers presumably has resulted from a reduction of
genome size in smooth tetraodontid puffers relative to their spiny
diodontid cousins during the 50–70 million years since their
divergence (Tyler 1980; Brainerd et al. 2001; Tyler and Santini 2002).
Table 1 lists genome size estimates for
multiple species within these groups. Finally, because the genomes of
two smooth pufferfish (Fugu rubripes and Tetraodon
nigroviridis) have been nearly completely sequenced, they offer a
tremendous opportunity to study genome size evolution by providing an
extremely detailed glimpse of the coding and noncoding sequence of
these organisms (Roest Crollius et al. 2000; Aparicio et al. 2002).
Here we use DNA renaturation kinetics to quantitatively describe and
compare the abundance of repetitive DNA in the differently sized
genomes of spiny and smooth pufferfish. Spiny pufferfish are
genetically uncharacterized, and the construction of Cot curves using
renaturation kinetics (Britten and Kohne 1968) has allowed us to build
the first portrait of the abundance of different classes of repetitive
DNAs relative to single-copy sequence in this unstudied genome.
Analyses of renaturation kinetics are based on the principle that when
a solution of sheared denatured DNA is kept at a temperature that
permits renaturation, the rate-limiting step of the reaction is the
collision of complementary fragments. Because the probability of
collision of complementary fragments is proportional to the square of
the concentration of those fragments in solution, highly repetitive DNA
sequences renature first, followed by repetitive sequences of lower
copy number, and then finally the single-copy sequences corresponding
to genes and other unique sequences. Measurement of the amount of
double-stranded DNA in a renaturing solution over time permits the
construction of a Cot (concentration x time) curve, which is used to
quantify different repetitive classes.
To complement our Cot-based description of the pattern of genome size
change in these fish, we test the hypothesis that a change in the
processes of insertion and deletion is responsible for genome size
differences in puffers. We estimate the genome-wide incidence of
small-scale insertions and deletions using nonfunctional retroelements
after the method of Petrov et al. (1996). Many non-LTR retroelement
transposition events result in a 5' truncation of the newly inserted
element, rendering it incapable of further transposition. Such elements
are said to be "dead on arrival" (DOA). Previous analyses indicate
that mutations affecting these DOA elements after truncation are
effectively neutral not only in terms of their effect on the elements'
coding capacity, but also in terms of their effects on genome size
(Petrov and Hartl 1998) and the propensity of the local sequence to
participate in ectopic recombination (Blumenstiel et al. 2002). They
are therefore ideal for estimation of the neutral mutation rate.
Phylogenetic analysis of multiple retrotransposon sequences is used to
separate mutations that occurred during the active, nonneutral phase of
the elements' lives from those mutations occurring during their
nonautonomous DOA phase. The former type of mutation is expected to be
found more than once in a densely sampled data set and therefore map to
the internal branches of an element tree. Unselected insertions,
deletions, and point substitutions are expected to map to the terminal
branches of a phylogenetic tree, which permits estimates of DNA loss or
gain relative to the nucleotide substitution rate (Petrov et al. 1996;
Petrov and Hartl 1997).
A similar rate of DNA loss between these two families of puffers,
combined with an underrepresentation of middle-repetitive DNA in
smooth puffers, lead us to invoke a reduced rate of large insertions as
a probable cause of the genome size reduction in smooth puffers. We
speculate on the role that transposable elements may have played in
this genome size reduction, and evaluate the importance of large
insertions versus small deletions as determinants of genome size.
|
RESULTS
|
|---|
Renaturation Kinetics
Cot curves for Fugu rubripes, a smooth puffer, and
Diodon hystrix, a spiny puffer, are presented in Figure
1. The curves were analyzed with the
least-squares regression program of Pearson et al. (1977).
Two-component curves were fitted to the data of each fish and were used
to compute the descriptive statistics for the repetitive and
single-copy fractions of the two genomes (Table
2). The most obvious difference between the
two curves is in the percentage of double-stranded DNA present
initially in the renaturation reaction. For F. rubripes and
D. hystrix, 19% and 7%, respectively, of the genomic DNA is
already present in double-stranded form at the earliest ECot
value (ECot represents the product of a correction factor for
the salt concentration of the buffer solution (E), the molar
concentration of nucleotides (Co), and the time of
renaturation in seconds (t; Britten et al. 1974).
View larger version (15K):
[in this window]
[in a new window]
|
Figure 1. Cot curves for a tetraodontid (smooth) puffer, Fugu rubripes,
and a diodontid (spiny) puffer, Diodon hystrix. Filled circles
indicate F. rubripes data points, and open circles represent
D. hystrix data points. Open arrows mark the Cot1/2
values of the D. hystrix repetitive (left) and
single-copy (right) classes, at which half of those components
are estimated to have renatured. The filled arrow indicates the
Cot1/2 value of the single-copy component of the F.
rubripes genome. The Cot1/2 value of the F.
rubripes repetitive component is estimated to be –3.3, and does
not appear on this graph.
|
|
The nature of DNA that has already renatured by the earliest assay
point in a Cot curve has been interpreted in various manners by
different authors. This DNA is sometimes assumed to be the highly
repetitive component of the genome, which renatures too quickly for
observation (Krajewski 1989; Verneau et al. 1991). Other authors assert
this component to be "foldback DNA," made up of a mixture of
repetitive and nonrepetitive fragments that share the property of being
able to form stem–loop structures upon themselves very quickly
(Britten et al. 1974; Jack and Hardman 1980; Pizon et al.
1984). To distinguish between these two possibilities, we constructed a
partial curve for Escherichia coli with our protocol (data not
shown). This curve yielded starting values of renatured DNA around 7%.
Because E. coli is known to not contain satellite DNA or other
highly repetitive sequences (Britten and Kohne 1968), it is not
possible in this case that the already renatured DNA is a purely
repetitive class. Therefore, in this analysis we do not consider the
already renatured DNA in the puffer curves to be a purely highly
repetitive class. The double-stranded DNA present at the earliest
ECot values in the pufferfish curves is likely a mixture of
fragments from different repetitive classes that are either
self-complementary, resistant to the initial denaturation step, or
otherwise refractory to digestion with S1 nuclease.
The total repetitive component of the smooth F. rubripes
genome based on renaturation analysis is 3%. This value is lower than
but comparable to partial library-based analyses of the repetitive
content of sequenced smooth pufferfish genomes, which place the
repetitive component at <10% of the genome (Elgar et al.
1999; Roest Crollius et al. 2000). In contrast, the
repetitive fraction of the spiny D. hystrix genome is 22%,
indicating a greater abundance of satellite DNA or transposable
elements in the spiny pufferfish family. Computation of the repetitive
frequency of this fraction from kinetic rate constants (see Methods)
indicates that middle repetitive rather than highly repetitive DNA
constitutes most of this fraction, because the average repeat occurs
150 times in the Diodon genome. The repetitive frequency of
the F. rubripes repetitive fraction is higher
(9 x 105 copies/genome), and may indicate that most of
this fraction is composed of satellite DNA.
Single-copy sequences were estimated to comprise 79% and 71% of the
F. rubripes and D. hystrix genomes, respectively
(Table 2). The repetitive frequency of these components was assumed to
be 1 copy/genome. The complexity of these components, defined as the
estimated length of the longest nonrepeating sequence in the class,
was, as expected, at least two orders of magnitude greater than the
complexity of the repetitive components in both curves.
Deletion/Insertion Profile
We used genomic DNA from four species of smooth pufferfish and four
species of spiny pufferfish to estimate insertion and deletion rates
(see Methods for species and sample sources). We PCR-amplified a
conserved region of the reverse transcriptase from non-LTR
retrotransposons. Between 15 and 25 sequences 300–600 bp in length
were generated from cloned PCR-amplified mixtures from one of each
species of fish. The final smooth pufferfish alignment contained 73
sequences of the previously described Maui non-LTR
retrotransposon (Poulter et al. 1999) and was 706 bp in length. The
final spiny pufferfish alignment contained 96 sequences of an
undescribed non-LTR retrotransposon and was 485 bp in length. This
undescribed element is more similar to Maui than any other
non-LTR element identified in F. rubripes according to BLAST
analysis (Altschul et al. 1997), but exhibits only 25%–30% amino
acid identity, and therefore cannot be assumed to be orthologous. It
was not possible to amplify any non-LTR element sequences from spiny
puffer genomic DNA using Maui-specific primers, and degenerate
primers failed to amplify any spiny elements with greater homology to
Maui.
Terminal branch mutations occurring in the sequences were determined
through phylogenetic analysis of aligned sequences for each group of
puffers. Maximum parsimony and neighbor-joining analyses yielded very
similar topologies and branch lengths for each data set, with most
differences in topology occurring in deep internal branches. The single
most parsimonious tree for the spiny puffer alignment and a 50%
majority rule consensus of the six most parsimonious trees for the
smooth puffer alignment were used for calculation of the mutation rate.
Mutations that mapped to terminal branches of each tree were considered
to have occurred during the neutral phase of an element's evolution,
and were used in subsequent analyses of mutation profiles.
Virtually all insertions and deletions mapped to the terminal branches
of the smooth and spiny puffer trees, but a small number of shared
deletions were observed in both data sets. These deletions were in each
case shared between two elements amplified from the same species, and
therefore likely represent alleles of a common locus rather than
separate insertions from a functional element containing a deletion.
Sequences did not group strictly according to species in the trees,
indicating that multiple active element lineages were possibly present
in the ancestors of each family. The average number of terminal branch
substitutions per base pair in the smooth and spiny alignments was
0.016 and 0.021, respectively. Assuming a time-uniform nucleotide
substitution rate on the order of 10–9 per base pair per
generation and a 1.5-yr generation time indicates that on average the
sequences in these alignments have been neutrally evolving for 25 to 35
million years. We are confident, then, that our mutation rate estimates
reflect the period of time during which the change in genome size may
have taken place in the tetraodontid lineage, rather than a time period
preceding the divergence of the two pufferfish families.
The distribution of insertion and deletion sizes was similar between
the two groups of fish (Fig. 2). The number
of indel events in each sequence is expected to be positively
correlated with the number of point substitutions after a period of
neutral evolution. This is the case for both the smooth and the spiny
pufferfish data sets, as seen in Figure 3
(Spearman's rs = 0.501, p < 0.0001
for spiny; rs = 0.367, p < 0.002 for
smooth).
View larger version (13K):
[in this window]
[in a new window]
|
Figure 2. Frequency histogram of deletion sizes for tetraodontid (smooth) and
diodontid (spiny) puffers. Represented are 31 deletions from the
tetraodontid alignment and 48 deletions from the diodontid alignment.
|
|
Neutral evolution in the terminal branches of the smooth and spiny
puffer trees is further supported by analysis of the distribution of
point substitutions among codon positions. Terminal branch
substitutions are approximately equally distributed among the three
codon positions ( 2 = 2.98, p < 0.22 for
spiny; 2 = 5.61, p < 0.061 for smooth).
Third-position substitutions are highly overrepresented on the internal
branches, however, where purifying selection favors synonymous
mutations (2 = 16.13, p < 0.001 for spiny;
2 = 29.42, p < 0.001 for smooth).
Estimates of deletion and insertion profiles for these two pufferfish
groups are very similar (Table 3). Deletion
and insertion rates scaled according to terminal nucleotide
substitutions (tns) were estimated with a maximum likelihood technique
that assumes these mutations are Poisson processes (Blumenstiel et al.
2002). The 95% confidence intervals were estimated by performing
bootstrap resampling of the data to re-estimate the parameter
(Blumenstiel et al. 2002). Tests for significance of rate differences
were carried out by comparing 1000 bootstrap estimates of the
parameters and determining the proportion of comparisons in which one
parameter estimate was larger than the other estimate (Blumenstiel et
al. 2002).
Neither the scaled deletion rate nor the scaled insertion rate is
significantly different between smooth and spiny puffers (deletions:
p < 0.095; insertions: p < 0.569). In spiny
puffers the deletion rate is significantly larger than the insertion
rate (0.06 deletions/tns vs. 0.03 insertions/tns;
p < 0.001). In smooth puffers the insertion and deletion
rates are not significantly different (0.04 deletions/tns vs. 0.03
insertions/tns; p < 0.142).
Average deletion and insertion sizes were also computed for both
groups, and no significant difference between smooth and spiny puffers
was found for either parameter (Wilcoxon test, deletions:
p < 0.122; insertions: p < 0.94). The average
deletion size is significantly larger than the average insertion size
for both groups, however (Wilcoxon test, smooth: p < 0.001;
spiny: p < 0.004). The average deletion size in smooth and
spiny puffers, respectively, is 19.8 and 19.1 bp. The average insertion
size in smooth and spiny puffers, respectively, is 2.7 and 2.6 bp.
Given that the smooth and spiny pufferfish alignments differ in length,
the possibility exists that the shorter spiny pufferfish alignment
might bias the average insertion and deletion size estimates downward
as a result of the alignment's smaller detection window. We sequenced
30% more spiny than smooth elements (96 vs.73) to provide some
protection against this bias. Furthermore, we note that there were no
deletions in the smooth puffer alignment large enough to escape capture
in the smaller spiny puffer window (Fig.
4). Truncation of the observed deletion
spectra above 40 bp yields average deletion size estimates of 8.96 bp
in smooth puffers and 7.96 bp in spiny puffers, which are not
significantly different (Wilcoxon test, p < 0.2122). We
conclude that the small-scale insertion/deletion profiles for these two
families of pufferfish are remarkably similar.
|
DISCUSSION
|
|---|
Spiny and smooth pufferfish have different genome size, and show
variation in the proportions of DNA in different repetitive classes.
Cot curve analysis has permitted us to efficiently characterize the
repetitive nature of the undescribed genome of the spiny pufferfish
D. hystrix, and compare it to the thoroughly sequenced genome
of the smooth pufferfish F. rubripes. Our complementary
estimation of the small-scale deletion/insertion profile in these
puffer families has yielded new information on a mutational process
thought to affect genome size (Petrov 2002). When the different
repetitive composition of these genomes is considered together with the
similar small-scale deletion and insertion rates of these two families
of pufferfish, a decline in the rate of large-scale insertions is
implicated as a probable cause of the genome size reduction in the
tetraodontid (smooth) puffer lineage.
Renaturation Data
Analysis of the Cot curves produced for smooth and spiny pufferfish
reveal that the genomes of fish in these two families differ not only
in size, but also repetitive content (Fig. 1, Table 2).
We detected a significant amount of double-stranded "foldback" DNA
present at the start of the renaturation reactions for both F.
rubripes and D. hystrix. Although the foldback components
of these pufferfish curves must be considered impure in terms of their
repetitive composition (see Results), it is nevertheless likely that
they contain the bulk of the highly repetitive satellite DNA found in
these genomes. The centromeric heterochromatin of the smooth puffers
F. rubripes and T. nigroviridis has been found to be
largely composed of 118-bp tandem repeats (Brenner et al. 1993; Roest
Crollius et al. 2000; Fischer et al. 2000). These tandem
repeats possess some degree of self-complementarity, and analysis with
the Mfold software program (SantaLucia 1998; SantaLucia et al.
1999) indicates they are capable of forming stem–loop
structures at the temperatures and salt concentrations used in this
renaturation experiment. Assuming there are similar tandem repeats in
D. hystrix, these satellite sequences may contribute heavily
to the foldback components observed in the curves. Although the
proportion of foldback DNA differs between the two puffer genomes, the
absolute amount in base pairs is very similar owing to the difference
in genome size (56 Mb in spiny vs. 76 Mb in smooth).
The quantity of highly repetitive and middle repetitive DNA in the
F. rubripes genome was too low to resolve the repetitive
component into these two fractions with our experimental protocol. As
the average repetitive frequency of this component is on the order of
105, however, this indicates that satellite DNA is the
primary component of this fraction. Pizon et al. (1984) constructed a
Cot curve for another species of smooth puffer, Arothron
diadematus, and recorded values of 5% and 7%, respectively, for
the proportions of highly repetitive and middle repetitive DNA classes
in the genome of this fish. The smaller amount of repetitive DNA we
detected in F. rubripes may be explained in part by the fact
that F. rubripes has a slightly smaller genome than A.
diadematus, with a haploid genome size of 0.42 pg of DNA/cell vs.
0.45 pg of DNA/cell for A. diadematus (Pizon et al. 1984;
Brenner et al. 1993). The discrepancy may also be caused in part by the
different protocols used to obtain each estimate. Some of the sequences
belonging to our foldback component, for example, may be represented in
the repetitive components of A. diadematus. Both estimates
place the smooth puffer genome in stark contrast to the repetitive
nature of other well-characterized vertebrate genomes in which, for
example, 45% of the human genome is composed of transposable
elements (International Human Genome Sequencing Consortium 2001).
Our renaturation data for D. hystrix indicate that 22% of
its genome is composed of repetitive DNA, which is almost seven times
the proportion of repetitive DNA we detected in F. rubripes.
As with F. rubripes, highly repetitive and middle repetitive
DNA classes were not resolvable. However, the repetitive frequency of
the repetitive component of the genome is 150, indicating an
abundance of middle repetitive sequences such as transposable elements
relative to highly repetitive sequences such as satellite DNA. As with
F. rubripes, this highly repetitive satellite DNA is likely
represented in the foldback component of the D. hystrix Cot
curve.
The single-copy components of the F. rubripes and D.
hystrix genomes do not differ very much in proportion, but do vary
in terms of absolute size when the difference in genome size is
considered. Our renaturation data indicate that single-copy sequences
comprise 316 Mb (79%) of the 400 Mb F. rubripes genome,
and 568 Mb (71%) of the 800 Mb D. hystrix genome. The
contraction of this component in smooth puffers has probably come
mainly at the expense of noncoding intergenic spaces and introns. Even
the smooth puffer genome is not at the minimum viable size for
vertebrates, however. It is estimated that only 108 Mb of the F.
rubripes genome is genic sequence (Aparicio et al. 2002), which
leaves nearly 200 Mb of single-copy sequence to which no function has
been assigned.
Deletion/Insertion Profile
All previous measurements of small insertions and deletions in other
organisms have revealed an overall bias toward DNA loss (Graur et al.
1989; Petrov and Hartl 1998; Petrov et al. 2000; Robertson 2000;
Bensasson et al. 2001). This bias is observed whether the sequence data
are derived from nuclear pseudogenes, nuclear mitochondrial
pseudogenes, or defunct non-LTR elements as in the case of our data
(for review, see Petrov 2002). This bias derives from a higher rate of
deletions than insertions, as well as a larger average deletion size
than insertion size. Pufferfish of the families Tetraodontidae and
Diodontidae exhibit a similar bias toward DNA loss, generated by
deletions that are larger and more frequent than insertions.
Higher rates of DNA loss through small ( 400 bp) deletions have been
observed to be negatively correlated with genome size in arthropods
(Petrov et al. 2000; Bensasson et al. 2001). Laupala crickets
and Podisma grasshoppers, which have genomes roughly 10 times
and 100 times, respectively, the size of Drosophila
melanogaster, have respective rates of DNA loss 13 times and 75
times slower than the rate of loss in Drosophila (for review,
see Petrov 2002).
We did not detect a significant difference in the insertion or deletion
profiles between the Tetraodontidae and Diodontidae families, despite a
twofold difference in genome size. This similarity in mutational
profiles should not necessarily be surprising, however, as Petrov and
Hartl (1998) failed to detect such a difference between two
Drosophila species with a similar difference in genome size.
Either the small-scale deletion bias is not an important determinant of
genome size variation among arthropods or vertebrates with only twofold
differences in genome size, or perhaps the present method is
insufficient to detect important variations in this parameter.
Dasilva et al. (2002) recently reported an independent estimation of
the small-scale deletion/insertion profile in the smooth puffer T.
nigroviridis from pseudogene data. Their data indicate a smaller
deletion bias than do our observations from DOA retrotransposons (0.56
vs. 0.71 bp lost per nucleotide substitution; Table 3), but are
concordant with our results in that they found a deletion bias greater
than that detected in mammals (Graur et al. 1989). From analysis of 66
copies of the Trapeze pseudogene, Dasilva et al. (2002) found
an average deletion size of 7.7 bp and an average insertion size of 1.5
bp. These values are approximately half the size of the averages we
computed from mutations observed in DOA retrotransposon sequences
(Table 3). One cause for the discrepancy may be selection acting on
mutations in the T. nigroviridis pseudogenes (Dasilva et al.
2002). Dasilva et al. (2002) found a higher incidence of substitutions,
insertions, and deletions in the exons than in the introns of the
pseudogene sequences, and they suggest that there may be selection
pressure to inhibit the pseudogenes' capacity to form functional mRNA
or proteins. In any case, the discrepancy between the pseudogene
estimate and our estimate of deletion bias in smooth puffers does not
affect conclusions deriving from the relative values of rates we
measured in smooth and spiny puffers.
Mechanisms of Genome Size Evolution in Pufferfish
Our complementary analyses of repetitive pattern and mutational
process in pufferfish genomes have permitted us to build a dynamic
understanding of genome evolution in this taxonomic group. To extend
these observations it is important to establish whether these
pufferfish genomes are in flux or equilibrium (Petrov 2002). If
tetraodontid and diodontid pufferfish genomes are presently at
equilibrium with regard to size, then a deletion bias at the length
scale observable in this experiment (<400 bp) must be counteracted by
an insertion bias at a larger scale (else the genomes would be quickly
reduced to their minimum viable size). It is not clear whether the
tetraodontid genomes are still contracting, but it seems likely that
the diodontid genomes are at an equilibrium size, given that the four
measurements of genome size for fish in this family are highly similar
to each other as well as to the genome size of the outgroup taxon
M. mola (Table 1). Given this equilibrium, we may therefore
infer that there is an unobserved input of DNA at larger length scales
into the diodontid genome to compensate for the observed small-scale
deletion bias.
The difference in genome size between the puffer families implies that
an ancestral equilibrium was disturbed in the smooth tetraodontid
lineage following its divergence from the spiny diodontids. A lack of
variation between the pufferfish families in the insertion/deletion
profile at the length scale observable in this experiment indicates
that the disturbance in equilibrium resulted from a change in the
insertion/deletion profile in the tetraodontid lineage at a larger
scale. In other words, the rate or size of large deletions relative to
large insertions may have increased, or the rate or size of large
insertions relative to large deletions may have decreased. Petrov
(2002) has speculated on the nature of large-scale insertions and
deletions, and points out that there are fundamental limits to the size
of deletions that are tolerated by the genome. Because all deletions
have two breakpoints, a larger deletion creates a greater chance that
one or both of the breakpoints will occur in a region that is under
natural selection, or that the deletion wholly removes a functional
sequence. Such deletions will probably be selectively deleterious, and
because they will be eliminated from the population they will have no
effect on genome size. Insertions have no such constraints. If an
insertion occurs between two individual nucleotides in a stretch of
noncoding DNA, it should make no difference whether the insertion is 4
nucleotides in length or 4000. The insertion will likely be selectively
neutral in either case, and will be fixed or lost from the population
with the same probability as other neutral mutations. A reduction in
the rate of large insertions in the tetraodontid lineage, rather than
an increase in large deletions, may therefore explain that family's
genome size contraction.
It is possible to calculate whether a difference in large insertions
alone would be sufficient to create the difference in genome size
observed between diodontid and tetraodontid puffers. If one assumes
that no large insertions or deletions >400 bp are occurring, the
amount of DNA lost (L) from small-scale deletion bias over a
given time period can be calculated:
where insertion and deletion rates are scaled according to nucleotide
substitutions, µ is the nucleotide substitution rate, G is
the haploid genome size in base pairs, and D is the number of
generations since divergence. Assuming a conservatively low nucleotide
substitution rate of 10–9 per base pair per generation, a
diploid genome size of 800 Mb, and 40 million generations since the
divergence of the families (assuming a 1.5-yr generation time), one can
calculate that the amount of DNA lost is estimated to be on the order
of 350 Mb. This is nearly sufficient to account for the difference in
size between the smooth and spiny puffer genomes. As a result, the
contraction of smooth puffer genome size could be caused by a severe
reduction of large insertions in this lineage compared with the spiny
puffers. Indeed, our renaturation data indicate that some portion of
these missing large insertions may result from reduced transposable
element activity, as middle repetitive sequences are rare in F.
rubripes but comprise roughly a fifth of the D. hystrix
genome.
Small Deletions Versus Large Insertions as Determinants of Genome Size
The small size of introns and intergenic spaces in the F.
rubripes genome has long been thought to have been generated by a
higher rate of DNA loss in this fish compared with humans (Brenner et
al. 1993; Aparicio et al. 2002). This hypothesis is supported by our
study. Although Graur et al. (1989) measured a small-scale deletion
rate in primates and rodents very similar to what we measured in
pufferfish, they detected a much smaller average deletion size. The
average deletion size in mammals is only 3.2 bp, which is nearly six
times smaller than the 19-bp average deletion size seen in puffers. The
insertional profile (size and rate) is similar in mammals and
pufferfish. Thus, the small-scale deletion bias seems to explain at
least part of the nearly eightfold difference in genome size between
smooth tetraodontid puffers and humans.
Indeed, the small-scale deletion biases we measured in smooth and spiny
pufferfish accord well with an observed inverse correlation between
this parameter and genome size in a wide variety of organisms (Petrov
2002), and support a hypothesis of a causal relationship between these
factors. For example, the rate of small-scale DNA loss (base pairs per
1 substitution) in smooth and spiny puffers is slower than the rate in
Drosophila melanogaster (Petrov and Hartl 1998) and
Caenorhabditis elegans (Robertson 2000), each of which have
smaller genomes than smooth and spiny pufferfish. Similarly,
Laupala crickets and Podisma grasshoppers have larger
genomes than pufferfish, and exhibit a slower rate of small-scale DNA
loss (Petrov et al. 2000; Bensasson et al. 2001).
Although the small-scale deletion bias may offer explanatory power with
regard to many-fold differences in genome size between these distant
animal taxa, it is not capable of explaining the twofold difference in
genome size between diodontid (spiny) and tetraodontid (smooth)
pufferfish. A substantial amount of DNA has been lost in smooth
pufferfish relative to their spiny cousins since their divergence
50–70 million years ago, without any significant change in the rate of
small-scale DNA loss. Indeed, the rate of deletion in the larger genome
of spiny puffers is actually slightly larger than in smooth puffers,
although not significantly so (Table 3). Because the average-sized
spiny puffer genome appears to be in equilibrium, one might argue that
it is primarily a recent contraction of genome size in smooth puffers
that is responsible for their curiously small genome size. To evolve
the smallest vertebrate genome yet measured, it was necessary that
tetraodontid puffers not only experience a high rate of DNA loss, but
also a low rate of large DNA insertions.
Small deletions have been given much attention as a possible
determinant of eukaryotic genome size. However, given that the
minuscule size of the smooth puffer genome may have been achieved
through a change in the rate of large insertions, it may be argued that
this side of the mutational spectrum should receive more attention.
Indeed, the observation that few eukaryotic genomes are even close to
their presumed minimum viable size implies that large-scale insertions
must constantly buoy them up against a seemingly ubiquitous small-scale
deletion bias. Systematic exploration of both increases and decreases
in transposable element activity over time at various taxonomic scales
should do much to clarify the role these pervasive characters have
played in eukaryotic genome evolution.
|
METHODS
|
|---|
Renaturation Analysis
Somatic tissue samples from F. rubripes and D.
hystrix were collected in Japan and Puerto Rico, respectively.
Genomic DNA was obtained from somatic tissue of F. rubripes
and D. hystrix using a phenol/chloroform extraction after the
method of Sambrook et al. (1989). Samples were treated with RNase and
then verified through a UV/vis spectrophotometer to have an
OD260/280 ratio of at least 1.80 and an OD260/230
ratio of at least 2.30. High-molecular-weight DNA was diluted with
water to a concentration of <500 µg/mL and then sonicated for four
periods of 30 sec to fragment it. DNA was checked on 1.5% agarose gels
to ensure that no fragments larger than 1 kb were present and that the
average fragment size was 500 bp.
Sheared DNA was precipitated and then resuspended in a solution of 2x
SSC to a concentration of 0.25 µg/µL. Purity was rechecked on a
spectrophotometer. Resuspended DNA was aliquoted in 10-µL volumes to
200-µL PCR tubes for denaturation and renaturation in a
thermalcycler. Aliquots were denatured through boiling for 9 min. The
temperature in the thermalcycler was then dropped to 55°C to permit
renaturation. Aliquots were removed from the thermalcycler at
predetermined time points in groups of three replicates and immediately
frozen in liquid nitrogen to prevent further renaturation. Frozen
aliquots were then thawed at room temperature after a 10-µL volume of
1.5 U/µL S1 nuclease solution was added to each tube. Samples were
incubated at 37°C for 80 min to permit nuclease digestion of
single-stranded sequences. Following incubation, nuclease activity was
halted by adding 20 µL of stop buffer (1 M Tris at pH 9.0, 0.1 M
EDTA). The remaining double-stranded DNA in solution was labeled with
Hoechst 33258 bisbenzamide dye and quantified with a fluorescence
spectrophotometer (with excitation at 356 nm and emission at 456 nm).
Cot curves were constructed by plotting the proportion of DNA remaining
in solution after digestion against the log of the ECot value
of each sample. ECot values were determined by multiplying a
correction factor for nonstandard salt concentrations (E) with
the molar concentration of nucleotides (Co) and time permitted
for renaturation in seconds (Britten et al. 1974). Data were analyzed
with the least-squares reassociation kinetics program of William
Pearson (Pearson et al. 1977), which fits a least-squares curve to the
data and then calculates a rate constant and complexity value for each
component. Cot1/2 values were determined by taking the log of
the inverse of the rate constants provided by the program for each
component (data not shown). The repetitive frequency of the repetitive
components was estimated by assuming the repetitive frequency of the
single-copy component was 1 copy/genome, and then dividing the rate
constant of each repetitive component by the rate constant of the
single-copy component for the same genome.
A partial Cot curve for E. coli strain TOP10 (Invitrogen) was
also constructed using the above techniques.
Insertion and Deletion Rate Estimates
Genomic DNA was extracted from somatic tissue samples of four
species of tetraodontid (smooth) puffers and four species of diodontid
(spiny) puffers using NucleoSpin columns (Clontech). The smooth puffers
studied were F. rubripes, T. nigroviridis,
Canthigaster valentini, and Canthigaster jactatus.
The spiny puffers used were D. hystrix, Diodon
holacanthus, Diodon eydouxii, and Chilomycterus
schoepfi. Tissue samples from F. rubripes and D.
eydouxii were collected in Japan, samples from T. nigroviridisand D. holacanthus were obtained from fish in the aquarium
trade, and the remaining samples were obtained from the University of
Kansas Museum of Natural History Tissue Collection. The reverse
transcriptase region of non-LTR retrotransposons was initially
amplified from D. holacanthus and F. rubripes using
the degenerate primers DVO144 and DVO145 of Wright et al. (1996)
according to the PCR protocol that accompanies their description. The
resultant amplifications were cleaned with Qiaquick columns (QIAGEN)
and cloned into XL1-Blue cells using a TOPO TA cloning kit
(Invitrogen). Random clones were screened with vector primers, and
inserts of all sizes were sequenced to prevent bias in
insertion/deletion rate estimates. Multiple sequences from each
transformation were generated on an ABI 3100 sequencer. All inserts
were sequenced in the forward and reverse directions. Ambiguous base
calls were discounted from further analysis or resequenced. BLAST
analysis (Altschul et al. 1997) confirmed that the sequences from
D. holacanthus were an undescribed non-LTR retrotransposon,
and indicated that the majority of sequences from F. rubripes
were the previously described non-LTR retrotransposon Maui
(Poulter et al. 1999). Sequences were aligned with CLUSTAL X (Thompson
et al. 1997), with all parameters set to default except for the gap
opening penalty (set to 20.0) and the DNA transition rate (set to 0).
Alignments were then adjusted by hand in BioEdit (Hall 1999) to
minimize the number of insertions and deletions. Adjustments were
performed to minimize the number of terminal substitutions when such
adjustments did not add to the number of insertions or deletions.
Ambiguous indel alignments were discounted from further analysis.
Variations in the length of monomeric microsatellite repeats in both
alignments were ignored. Both alignments contained a minority of
sequences that appeared to be from different element families. These
sequences were discarded to prevent bias in mutation rate estimation.
Conserved regions from alignable sequences were used to design
family-specific primers, as the degenerate DVO144/145 primers only
weakly amplified reverse transcriptase sequences from these groups.
MAUIF1 (ACCAGATGTGCTGACTGTGG) and MAUIR1 (TTGAGGAACTCCATGGCTAAC) were
used to amplify Maui non-LTR reverse transcriptase sequences from
smooth puffer samples, and DIODF1 (GTGGACAACAATAGCGCCAC) and DIODR1
(CCTTACAGATGAAATTACGGAGC) were used to amplify the undescribed element
from the spiny puffer samples. All PCR reactions were performed under
standard conditions with an annealing temperature of 56°C. Products
amplified with these family-specific primers were cleaned, cloned,
sequenced, and aligned as above. Sequences are deposited in GenBank
under the following accession numbers: AY212336–AY212504.
Reverse transcriptase alignments for smooth and spiny puffers were used
to build maximum parsimony and neighbor-joining phylogenies in PAUP*
4.0b10 (Swofford 2002). Heuristic searches with TBR branch-swapping
were performed, and gaps were ignored. Branch length estimates from
parsimony and distance trees were compared and verified to be
significantly correlated (Spearman's
rs < 0.001). Nucleotide substitutions,
insertions, and deletions that mapped to the terminal branches of the
most parsimonious tree for spiny puffers and a 50% majority rule
consensus of six most parsimonious smooth puffer trees were used in
subsequent analyses of mutation rate. These terminal,
autapomorphic mutations are assumed to have arisen during
the nonautonomous, neutral phase of the retroelements' evolution, and
are assumed to represent the true, neutral, genome-wide mutation rate
(Petrov et al. 1996). Maximum likelihood estimates of the ratio of
insertions to nucleotide substitutions and the ratio of deletions to
nucleotide substitutions were made using a method described in
Blumenstiel et al. (2002), which accounts for biases resulting from
differences in the observed versus the presumed original length of the
reverse transcriptase sequences. This method assumes that nucleotide
substitution and indel formation are Poisson processes. Bootstrap
resampling of the original data was used to generate 1000 replicates of
each parameter estimate, from which 95% confidence intervals for the
rate estimates were derived (Blumenstiel et al. 2002).
|
Acknowledgements
|
|---|
We thank Justin Blumenstiel and William Pearson for help with our
data analysis. We thank Toshiaki Itami, Christian Landry, Steve
Vollmer, and the Museum of Natural History at the University of Kansas
for tissue samples. Comments and criticism from Elena
Lozovsky, Cristian Castillo-Davis, and two anonymous reviewers improved
this paper. This work was supported by grants from the NSF to S.R.P.
D.E.N. was supported in part by an NSF Pre-Doctoral Fellowship.
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
|
|---|
3 Corresponding author. 
E-MAIL neafsey{at}oeb.harvard.edu; FAX (617) 496-5854.
Article and publication are at
http://www.genome.org/cgi/doi/10.1101/gr.841703.
|
REFERENCES
|
|---|
Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. 1997. Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucleic Acids Res. 25: 3389-3402.[Abstract/Free Full Text]
Aparicio, S., Chapman, J., Stupka, E., Putnam, N., Chia, J.M., Dehal, P., Christoffels, A., Rash, S., Hoon, S., Smit, A.F., et al. 2002. Whole-genome shotgun assembly and analysis of the genome of Fugu rubripes. Science 297: 1301-1310.[Abstract/Free Full Text]
Bensasson, D., Petrov, D.A., Zhang, D.X., Hartl, D.L., and Hewitt, G.M. 2001. Genomic gigantism: DNA loss is slow in mountain grasshoppers. Mol. Biol. Evol. 18: 246-253.[Abstract/Free Full Text]
Blumenstiel, J.P., Hartl, D.L., and Lozovsky, E.R. 2002. Patterns of insertion and deletion in contrasting chromatin domains. Mol. Biol. Evol. 19: 2211-2225.[Abstract/Free Full Text]
Brainerd, E.L., Slutz, S.S., Hall, E.K., and Phillis, R.W. 2001. Patterns of genome size evolution in tetraodontiform fishes. Evol. Int. J. Org. Evol. 55: 2363-2368.
Brenner, S., Elgar, G., Sandford, R., Macrae, A., Venkatesh, B., and Aparicio, S. 1993. Characterization of the pufferfish (Fugu) genome as a compact model vertebrate genome. Nature 366: 265-268.[CrossRef][Medline]
Britten, R.J. and Kohne, D.E. 1968. Repeated sequences in DNA. Hundreds of thousands of copies of DNA sequences have been incorporated into the genomes of higher organisms. Science 161: 529-540.[Free Full Text]
Britten, R.J., Graham, D.E., and Neufeld, B.R. 1974. Analysis of repeating DNA sequences by reassociation. Methods Enzymol. 29: 363-418.[Medline]
Dasilva, C., Hadji, H., Ozouf-Costaz, C., Nicaud, S., Jaillon, O., Weissenbach, J., and Roest Crollius, H. 2002. Remarkable compartmentalization of transposable elements and pseudogenes in the heterochromatin of the Tetraodon nigroviridis genome. Proc. Natl. Acad. Sci. 99: 13636-13641.[Abstract/Free Full Text]
Elgar, G., Clark, M.S., Meek, S., Smith, S., Warner, S., Edwards, Y.J., Bouchireb, N., Cottage, A., Yeo, G.S., Umrania, Y., et al. 1999. Generation and analysis of 25 Mb of genomic DNA from the pufferfish Fugu rubripes by sequence scanning. Genome Res. 9: 960-971.[Abstract/Free Full Text]
Fischer, C., Ozouf-Costaz, C., Roest Crollius, H., Dasilva, C., Jaillon, O., Bouneau, L., Bonillo, C., Weissenbach, J., and Bernot, A. 2000. Karyotype and chromosome location of characteristic tandem repeats in the pufferfish Tetraodon nigroviridis. Cytogenet Cell Genet. 88: 50-55.[CrossRef][Medline]
Graur, D., Shuali, Y., and Li, W.H. 1989. Deletions in processed pseudogenes accumulate faster in rodents than in humans. J. Mol. Evol. 28: 279-285.[CrossRef][Medline]
Gregory, T.R. 2001. Coincidence, coevolution, or causation? DNA content, cell size, and the C-value enigma. Biol. Rev. Camb. Philos. Soc. 76: 65-101.[Medline]
Hall, T.A. 1999. BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucl. Acid Symp. Ser. 41: 95-98.
Hinegardner, R. and Rosen, D.E. 1972. Cellular DNA content and the evolution of teleostean fishes. Am. Nat. 106: 621-644.[CrossRef]
International Human Genome Sequencing Consortium 2001. Initial sequencing and analysis of the human genome. Nature 409: 860-921.[CrossRef][Medline]
Jack, P.L. and Hardman, N. 1980. Sequence organization in nuclear deoxyribonucleic acid from Physarum polycephalum. Physical properties of foldback sequences. Biochem. J. 187: 105-113.[Medline]
John, B. and Miklos, G.L.C., 1988. The eukaryotic genome in development and evolution, pp. 1–416. Allen and Unwin, London, UK.
Krajewski, C. 1989. Comparative DNA reassociation kinetics of cranes. Biochem. Genet. 27: 131-136.[Medline]
Ojima, Y. and Yamamoto, K. 1990. Cellular DNA contents of fishes determined by flow cytometry. La Kromosomo II 57: 1871-1888.
Pearson, W.R., Davidson, E.H., and Britten, R.J. 1977. A program for least squares analysis of reassociation and hybridization data. Nucleic Acids Res. 4: 1727-1737.[Abstract/Free Full Text]
Petrov, D.A. 2002. Mutational equilibrium model of genome size evolution. Theoret. Pop. Biol. 61: 531-544.[CrossRef][Medline]
Petrov, D.A. and Hartl, D.L. 1997. Trash DNA is what gets thrown away: High rate of DNA loss in Drosophila. Gene 205: 279-289.[CrossRef][Medline]
___, 1998. High rate of DNA loss in the Drosophila melanogaster and Drosophila virilis species groups. Mol. Biol. Evol. 15: 293-302.[Abstract]
Petrov, D.A., Lozovskaya, E.R., and Hartl, D.L. 1996. High intrinsic rate of DNA loss in Drosophila. Nature 384: 346-349.[CrossRef][Medline]
Petrov, D.A., Sangster, T.A., Johnston, J.S., Hartl, D.L., and Shaw, K.L. 2000. Evidence for DNA loss as a determinant of genome size. Science 287: 1060-1062.[Abstract/Free Full Text]
Pizon, V., Cuny, G., and Bernardi, G. 1984. Nucleotide sequence organization in the very small genome of a tetraodontid fish, Arothron diadematus. Eur. J. Biochem. 140: 25-30.[Medline]
Poulter, R., Butler, M., and Ormandy, J. 1999. A LINE element from the pufferfish (fugu) Fugu rubripes which shows similarity to the CR1 family of non-LTR retrotransposons. Gene 227: 169-179.[CrossRef][Medline]
Robertson, H.M. 2000. The large srh family of chemoreceptor genes in Caenorhabditis nematodes reveals processes of genome evolution involving large duplications and deletions and intron gains and losses. Genome Res. 10: 192-203.[Abstract/Free Full Text]
Roest Crollius, H., Jaillon, O., Dasilva, C., Ozouf-Costaz, C., Fizames, C., Fischer, C., Bouneau, L., Billault, A., Quetier, F., Saurin, W., et al. 2000. Characterization and repeat analysis of the compact genome of the freshwater pufferfish Tetraodon nigroviridis. Genome Res. 10: 939-949.[Abstract/Free Full Text]
Sambrook, J., Fritsch, E.F., and Maniatis, T., 1989. Molecular cloning: A laboratory manual Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
SantaLucia, J., Jr. 1998. A unified view of polymer, dumbbell, and oligonucleotide DNA nearest-neighbor thermodynamics. Proc. Natl. Acad. Sci. 95: 1460-1465.[Abstract/Free Full Text]
Swofford, D.L., 2002. PAUP*: Phylogenetic Analysis Using Parsimony (*and Other Methods), Version 4, Sinaur Associates, Sunderland, MA.
Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., and Higgins, D.G. 1997. The CLUSTAL_X windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 25: 4876-4882.[Abstract/Free Full Text]
Tyler, J.C. 1980. Osteology, phylogeny, and higher classification of the fishes of the order Plectognathi (Tetraodontiformes). NOAA Tech. Rep. NMFS Circ. 434: 1-422.
Tyler, J.C. and Santini, F. 2002. Review and reconstructions of the tetraodontiform fishes from the Eocene of Monte Bolca, Italy, with comments on related Tertiary taxa. Studi e ricerche sui Giacimenti Terziari di Bolca. Mus. Civico di Storia Nat. di Verona 9: 117-119.
Verneau, O., Renaud, F., and Catzeflis, F.M. 1991. DNA reassociation kinetics and genome complexity of a fish (Psetta maxima: Teleostei) and its gut parasite (Bothriocephalus gregarius: Cestoda). Comp. Biochem. Physiol. B 99: 883-886.[CrossRef][Medline]
Vinogradov, A.E. 1998. Genome size and GC-percent in vertebrates as determined by flow cytometry: The triangular relationship. Cytometry 31: 100-109.[CrossRef][Medline]
Wright, D.A., Ke, N., Smalle, J., Hauge, B.M., Goodman, H.M., and Voytas, D.F. 1996. Multiple non-LTR retrotransposons in the genome of Arabidopsis thaliana. Genetics 142: 569-578.[Abstract]
Received September 27, 2002;
accepted in revised format February 25, 2003.
13:821-830 © by 2003 Cold Spring Harbor Laboratory Press ISSN 1088-9051/03 $5.00
 CiteULike  Connotea  Del.icio.us  Digg  Reddit  Technorati What's this?
This article has been cited by other articles:
|
|
|
|
 
M. C. Jost, D. M. Hillis, Y. Lu, J. W. Kyle, H. A. Fozzard, and H. H. Zakon
Toxin-Resistant Sodium Channels: Parallel Adaptive Evolution across a Complete Gene Family
Mol. Biol. Evol.,
June 1, 2008;
25(6):
1016 - 1024.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
C. M. Bergman and D. Bensasson
Recent LTR retrotransposon insertion contrasts with waves of non-LTR insertion since speciation in Drosophila melanogaster
PNAS,
July 3, 2007;
104(27):
11340 - 11345.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
M. Brandstrom and H. Ellegren
The Genomic Landscape of Short Insertion and Deletion Polymorphisms in the Chicken (Gallus gallus) Genome: A High Frequency of Deletions in Tandem Duplicates
Genetics,
July 1, 2007;
176(3):
1691 - 1701.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
J.-N. Volff, H. Lehrach, R. Reinhardt, and D. Chourrout
Retroelement Dynamics and a Novel Type of Chordate Retrovirus-like Element in the Miniature Genome of the Tunicate Oikopleura dioica
Mol. Biol. Evol.,
November 1, 2004;
21(11):
2022 - 2033.
[Abstract]
[Full Text]
| |
![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
Top
ABSTRACT
RESULTS
