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© 2002 Oxford University Press
Nucleic Acids Research, 2002, Vol. 30, No. 12 e52
Improved detection of small deletions in complex pools
of DNA
Mark Edgley, Anil D’Souza1, Gary Moulder1, Sheldon McKay, Bin Shen, Erin Gilchrist,
Donald Moerman and Robert Barstead1,*
Biotechnology Laboratory and Department of Zoology, University of British Columbia, Vancouver, BC V6T 1Z4,
Canada and 1Department of Molecular and Cell Biology, Oklahoma Medical Research Foundation, 825 N.E. 13th
Street, Oklahoma City, OK 73104, USA
Received December 14, 2001; Revised and Accepted April 16, 2002
ABSTRACT
About 40% of the genes in the nematode Caenorhabditis
elegans have homologs in humans. Based on the
history of this model system, it is clear that the
application of genetic methods to the study of this set
of genes would provide important clues to their function in humans. To facilitate such genetic studies, we
are engaged in a project to derive deletion alleles in
every gene in this set. Our standard methods make use
of nested PCR to hunt for animals in mutagenized
populations that carry deletions at a given locus. The
deletion bearing animals exist initially in mixed populations where the majority of the animals are wild type at
the target. Therefore, the production of the PCR fragment representing the deletion allele competes with the
production of the wild type fragment. The size of the
deletion fragment relative to wild type determines
whether it can compete to a level where it can be
detected above the background. Using our standard
conditions, we have found that when the deletion is
<600 bp, the deletion fragment does not compete
effectively with the production of the wild type fragment in PCR. Therefore, although our standard
methods work well to detect mutants with deletions
>600 bp, they do not work well to detect mutants with
smaller deletions. Here we report a new strategy to
detect small deletion alleles in complex DNA pools. Our
new strategy is a modification of our standard PCR
based screens. In the first round of the nested PCR, we
include a third PCR primer between the two external
primers. The presence of this third primer leads to the
production of three fragments from wild type DNA. We
configure the system so that two of these three fragments cannot serve as a template in the second round
of the nested PCR. The addition of this third primer,
therefore, handicaps the amplification from wild type
template. On the other hand, the amplification of
mutant fragments where the binding site for the third
primer is deleted is unabated. Overall, we see at least a
500-fold increase in the sensitivity for small deletion
fragments using our new method. Using this new
method, we report the recovery of new deletion alleles
within 12 C.elegans genes.
INTRODUCTION
Until recently, most genes in the nematode Caenorhabditis
elegans were known through their mutant phenotypes. Now,
however, more C.elegans genes are known through sequencing
than through genetics (1); of the approximately 19 000 genes in
C.elegans, only about 1300 have mutant alleles. As genetics is
one of the most important tools in the arsenal of C.elegans
biologists, we and others have devised methods to derive mutations in C.elegans genes known only through their sequence
(2–6). Our present methods are conceptually simple. We treat
worms at the L4 larval stage with a mutagen. The progeny of
the mutagenized worms are subdivided into populations that
are allowed to reproduce. We then extract the DNA from ∼30%
of each population. The extracted DNA samples are pooled and
subjected to PCR with nested primer sets (Fig. 1). Candidate
populations are identified by the presence of a PCR product
that is smaller than the size predicted by the genomic DNA
sequence. Each candidate population is subdivided and
subjected to similar growth and PCR analysis. This process of
sib selection continues until we recover a single individual
with the deletion. Using this protocol, typically we can recover
such an individual in three steps of growth and sib selection.
In principle, our ability to detect a deletion should be a
function of the resolution of the gels used for electrophoresis.
In practice, however, gel resolution does not determine the size
of the deletions that we identify; typically, we do not detect
deletions that are less than ∼600 bp. This limitation is a consequence of our effort to reduce, as far as possible, the cost and
time required to identify a candidate deletion. To accommodate
the relatively low frequency at which deletions are induced by
chemical mutagens (7), we pool the DNA template samples to
minimize the numbers of PCRs that are needed to detect a deletion. Using our present DNA pooling strategy, for every copy
of the deletion DNA we have approximately 12 000 copies of
wild type DNA. This places the mutant template at a substantial
disadvantage in PCR. Deletion fragments that are close to the
size of wild type DNA apparently are not able to overcome this
initial disadvantage.
*To whom correspondence should be addressed. Tel: +1 405 271 1766; Fax: +1 405 271 3153; Email: [email protected]
e52 Nucleic Acids Research, 2002, Vol. 30, No. 12
Figure 1. Protocol schematic. Our current protocols use nested PCR to identify
smaller than normal PCR fragments derived from a mutagenized population of
worms. Typically, we use two external (A and C) and two internal PCR primers
(D and E). As shown, the poison primer method adds a primer (B) in the first
round of the nested PCR.
However, recent work has shown that a significant proportion
of deletions induced by trimethylpsoralen (TMP) treatment
followed by UV irradiation are <600 bp, falling in the range of
50–600 bp (E.Gilchrist, M.Edgley, G.Mullen, B.Shen,
T.Rogalski, T.Szczygielski and D.Moerman, manuscript in
preparation) (8). Based on these data, it is likely that we do not
detect a significant number of TMP-induced deletion mutants
using our standard screening strategy. The challenge then
became to develop methods that would allow us to detect small
deletions. This paper reports such a method.
MATERIALS AND METHODS
Reconstruction experiments
Our initial tests were done with a known 133 bp deletion in the
C.elegans gene dim-1(gk54) (GenBank locus U39667). We
used the following oligonucleotide primers in nested PCR as
shown in Figure 1: first round, 5′-CTCAGTCGATCACAGTACA-3′, 5′-TCCACCAACAAGCTTTTGCC-3′; second round,
5′-ACACTTCCCACAACAACCAG-3′, 5′-CGGTAAGCTTCAGGTTGAAG-3′; internal poison, (A) 5′-CGAACAAGGGAAGCGACAGC-3′, (B) 5′-GTTGGCACTGAAGCGTCCAG-3′.
Each primer was used at a final concentration of 1 µM. We
did PCR using template DNA purified from wild type or
dim-1 mutant strains. Each of the necessary deoxynucleotide
triphosphates was at a concentration of 250 µM. The PCR
cycling parameters were as follows: 94°C, 30 s; 60°C, 30 s;
72°C, 90 s. We did 35 total cycles. PCR fragments were
analyzed on 1% agarose gels.
Generation and screening of mutant libraries
To induce deletion mutations, young adult hermaphrodites
were treated with trimethylpsoralen and UV light as described
(7) (E.Gilchrist, M.Edgley, G.Mullen, B.Shen, T.Rogalski,
T.Szczygielski and D.Moerman, manuscript in preparation). F1
progeny of mutagenized animals were cultured in 1152 groups
of 50 worms each. After one generation, DNA was prepared
from each population by proteinase K lysis. PCR was used to
identify populations with animals carrying deletions using the
cycling parameters described below. Populations carrying a
deletion were repeatedly subdivided until homozygotes
carrying the deletion were obtained. Deletion endpoints were
determined by sequencing PCR products that spanned the
deleted region.
PCR conditions
External-round screening PCRs were 10 µl and contained 2 µl
library DNA, 4 pM each external primer and poison primer,
200 µM each dNTP, 1× standard PCR buffer containing
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1.5 mM MgCl2 and 0.2 U Taq polymerase (Roche). Internalround PCRs were identical in all respects except for input DNA
(0.2 µl from external round) and primers (4 pM each internal
primer). Replication of the external round into the internal
round was done with a Robbins Hydra 96 pipetting station.
Thermal cycling was done on MJ Research PTC-200 DNA Engines
or Biometra Uno II thermal cyclers using the following
conditions: 94°C/30s, 35 cycles of 94°C/30 s – 61°C/30 s
(external round) or 55°C/30 s (internal round) – 72°C/60 s,
followed by cooling to 4°C. Loading buffer (10 µl; 15% Ficoll
with bromophenol blue and xylene cyanol) was added to each
reaction, and 1 µl of this mix was loaded into each lane of 2%
agarose gels. Gels were stained for 30 min with fresh 1:10 000
SYBR® Green (BMA) and imaged with a Molecular Dynamics
Fluorimager. The molecular weight marker is BioRad 100 bp
PCR Molecular Ruler (Cat. 170-8206). The wild type amplification product is 1151 bp and the deletion product is 810 bp.
RESULTS
Our initial experiments were done with a known small deletion
in the C.elegans gene dim-1 (T.Rogalski and D.Moerman,
unpublished data). We first tested different pooling strategies,
hoping to reduce the competition between the deletion and
wild type fragments. We found that a 133 bp deletion could be
detected only if the initial ratio of wild type to deletion
template did not exceed 10:1 (Fig. 2). Together with the
forward frequency for PCR detectable deletions generated by
TMP (probably <1 × 10 –5 for the average gene) this indicates
that an unreasonably large number of PCRs would be required
to detect a small deletion. It was not feasible, therefore, simply
to reduce the pooling depth to improve our ability to detect
small deletions.
Reconstruction experiments with known deletion alleles
To improve the sensitivity of the PCR assay for deletion fragments that are close to the wild type size, we developed the
strategy shown in Figure 1. This strategy is a modification of
our original protocol, in which we used a two-step nested PCR
series to detect deletions between the primer sets. In the modified version, a third functional PCR primer that falls between
the two external primers is included in the first round of nested
PCR. Amplification from the wild type template leads to the
production of two fragments, one full length and the other
relatively short. In practice, the shorter fragment is produced
much more efficiently than the longer. Amplification from a
mutant template, in which the site for the third internal primer
is deleted, leads to the production of a single mutant fragment
from the external primers. In the second round of PCR, we use
two primers placed just inside the external first round primers.
The shorter wild type band from the first round cannot serve as
a template for the second round PCR because it does not
include one of the second round primer binding sites. The
longer wild type fragment can serve as a template, but because
its production was limited by competition in the first round, its
production in the second round is limited correspondingly. The
internal functional primer is called a ‘poison’ because it interferes
with production of the full-length wild type fragment. The effectively lower level of wild type product gives the deletion fragment
an advantage. Our data show that we can detect a 133 bp
deletion at wild type to mutant ratios of 5000:1 (Fig. 2).
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Nucleic Acids Research, 2002, Vol. 30, No. 12 e52
Figure 3. Poison primer strategy works to identify new deletion alleles. Gel
image of nested PCR amplification products illustrating the effect of a poison
primer in the external amplification round on deletion detection. Twelve
samples of pooled library DNA, one of which contained a 341-bp deletion in
the gene H32C10.3 at a complexity of one deletion chromosome in 240, were
subjected in duplicate to PCR amplification with and without a poison primer
that lies within the deleted region. Starting after the marker lane (M), the two
leftmost lanes contain products of reactions with the poison primer, and the
two rightmost lanes contain products of reactions without the poison. The wild
type and deletion fragment positions are as indicated.
Figure 2. Poison primers improve sensitivity for small deletions. We tested the
poison primer method on a known deletion mutant. We did nested PCR using
two different internal poison primers, designated competing oligos A and B
that were known to fall within the deleted sequence. Different amounts of wild
type and deletion mutant DNAs were mixed at the indicated ratios. With no
poison primer, wild type gives a 701 bp PCR fragment. The deletion mutant
gives a 568 bp fragment. Without the poison primer, we lose the ability to
detect the deletion at wild type:mutant ratios between 10:1 and 100:1. With the
poison primer, we can detect the deletion fragment at ratios of 5000:1.
Screens for new alleles
We tested our new strategy in screens for new deletion
mutants. We cultured the F1 progeny of mutagenized parents in
sets of 10 or 50 worms. We harvested a portion of each F2
population and prepared DNA for PCR. Although, as
described above, we were able to detect a small 133 bp deletion
in pools where the ratio of wild type to mutant DNA reached
5000:1, we did not exceed a ratio of 1200:1 in screens for
new unknown deletions. Figure 3 shows an example of a
341 bp deletion identified using the poison primer method
and a pooling ratio of 240:1. The detection of this deletion
required the presence of the poison primer, as the deletion
band was clearly visible when it was in the reaction mix but was
not detected otherwise (Fig. 3). This example clearly illustrates
the enhanced detection that can be obtained using this strategy.
We identified 12 deletion strains from a UV/TMP mutagenized library of mutants using the poison primer method.
The deletions ranged from 78 to 850 bp. Examples of these are
shown in Figure 4A. Figure 4B shows the position of a small
deletion that we recovered in a gene that resides entirely within
the intron of another gene. DNA sequencing showed that 11 of
the 12 deletions encompass the poison primer binding sites.
The poison primers were required to detect 10 of these 11
deletions in the initial mutant library screens (data not shown).
The exceptional case was a deletion of 850 bp in a 1335-bp
wild type interval.
DISCUSSION
Genome sequence data provide a platform for standard
mutational genetics, a powerful strategy to examine gene
function. To exploit genetic methods, however, one must identify mutations at target loci. Using our original protocols, we
could detect deletions >600 bp in an interval of 3000 bp. Using
the poison primer protocol, we have detected deletions as small
as 78 bp. We also tested whether we could improve the method
through the simultaneous use of two closely spaced poison
primers annealing to opposite strands of the target. Although
our data set is small, two poisons appeared to be somewhat
better at reducing the level of the competing wild type PCR
product (data not shown).
Although the poison primer method allows for the detection
of small deletion alleles in complex DNA pools, it changes the
target for deletion relative to our typical strategy. With the
poison primer method the deletion must eliminate the internal
poison primer site. In a 3000 bp interval, therefore, with a
poison annealing to a single site one can detect only ∼7% of the
total number of 100 bp deletions, 14% of the 200 bp deletions,
etc. The poison primer method, however, gives greater control
over the position of the deletion within the target, thereby
allowing for the recovery of more precise ‘designer’ deletions.
Cases where this method might be useful include the deletion
of a single exon in alternatively spliced genes, the deletion of
parts of promoters, or the deletion of genes that fall within
introns of other genes. The data in Figure 4B validate the
poison primer method for such uses.
Although the exact endpoints of the deletions are not subject
to our control, we generally target the 5′ end of the coding
sequence to enhance the odds that the recovered deletion alleles
will eliminate all functions of the gene. We have developed a
public interactive web site that can be used to identify primer
sequences that meet our design criteria (http://ko. cigenomics.bc.ca/
oligos.shtml).
We have used the nematode C.elegans to develop a new
method to detect deletion mutations in known genes. Our new
method enhances the reverse genetic tools available for
C.elegans and other genetic model systems and, further, adds to
the growing list of genetic tools developed to screen for
mutations in humans in appropriate clinical or research
populations.
e52 Nucleic Acids Research, 2002, Vol. 30, No. 12
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Figure 4. Detection of deletions with the poison primer technique. The gene structure for each of the genes listed was predicted from genomic DNA sequence.
Grey boxes and thin lines represent exons and introns, respectively. The positions of the deletions and their sizes are shown below each gene. Arrows represent the
poison primers designed to interfere with the amplification of a full-length wild type PCR product. Particular regions of the gene can be targeted based on the
placement of such primers. (A) Representative deletion mutations. Deletion breakpoints were determined by DNA sequencing of the internal PCR products amplified
from DNA isolated from single worms. The deletions shown are 342 NT, 769 NT, 483 NT and 369 NT in the coding regions of H32C10.3, C01G8.2, T19E10.1a
and Y56A3A.20, respectively. Note that the deleted regions contain all or part of the poison primer-binding site. (B) Precise excision of a nested gene. R13H4.1 is
a large gene whose coding sequence is on the minus strand of clone R13H4 of chromosome II of C.elegans. The gene R13H4.2, one of two genes nested within
introns of the larger gene on the opposite strand, was targeted with two oppositely oriented poison primers. The deletion mutation detected with these primers
excised the entire 3′ end of R13H4.2, including the target exon, but did not extend outside of the intron of the larger gene, leaving its coding sequence intact.
ACKNOWLEDGEMENTS
We thank Marco Marra and Steven Jones of the Genome
Sequence Center in Vancouver BC for their advice and
enthusiasm for this project, and for the use of their fluorimager
to scan gels. This work was supported by a National Institutes
of Health grant R01 HG01843 to R.B., and a grant from the
Canadian Institute for Health Research to D.G.M. Additional
support was provided by the Biotechnology Laboratory and
U.B.C. to the C.elegans Reverse Genetics Core Facility.
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