From the Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, Massachusetts 02115
Received for publication, October 16, 2002, and in revised form, December 19, 2002
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ABSTRACT |
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Gene 2.5 of bacteriophage T7 encodes a
single-stranded DNA-binding protein that is essential for viral
survival. Its crystal structure reveals a conserved
oligosaccharide/oligonucleotide binding fold predicted to interact with
single-stranded DNA. However, there is no experimental evidence to
support this hypothesis. Recently, we reported a genetic screen for
lethal mutations in gene 2.5 that we are using to identify functional
domains of the gene 2.5 protein. This screen uncovered a number of
mutations that led to amino acid substitutions in the proposed DNA
binding domain. Three variant proteins, gp2.5-Y158C, gp2.5-K152E, and gp2.5-Y111C/Y158C, exhibit a decrease in binding affinity for oligonucleotides. A fourth, gp2.5-K109I, exhibits an altered mode of
binding single-stranded DNA. A carboxyl-terminal truncation of gene 2.5 protein, gp2.5- Single-stranded DNA
(ssDNA)1-binding proteins
lack sequence specificity and bind ssDNA with a higher affinity than
they bind double-stranded DNA or RNA (1). Primarily, ssDNA-binding
proteins function to bind any exposed regions of ssDNA in cells,
forming a protective coat around the reactive bases and thus
restricting the formation of secondary structures. However, their role
is not restricted to extending and protecting DNA in that they also physically and functionally interact with other replication proteins. Bacteriophage T7 encodes its own ssDNA-binding protein, the product of
gene 2.5. Gene 2.5 protein is essential for phage survival (2) and
plays multiple roles in DNA replication, recombination, and repair
(2-12). Gene 2.5 protein interacts directly with both the T7 DNA
polymerase (9) and the gene 4 helicase/primase (7), stimulating the
activity of each protein. Presumably these interactions explain why
coordination of leading and lagging strand synthesis in
vitro is dependent upon gene 2.5 protein (13). Furthermore, gene
2.5 protein facilitates homologous DNA base
pairing,2 a process that is
important during viral recombination (10, 11, 14) and in the repair of
double-stranded breaks in the T7 chromosome (12).
Despite functional similarity with other ssDNA-binding proteins, namely
the Escherichia coli SSB protein and the bacteriophage T4
gene 32 protein, T7 gene 2.5 protein has no sequence homology with
these proteins (15, 16). Furthermore, these proteins cannot substitute
for gene 2.5 protein in vivo (2, 17). The mode of binding of
gene 2.5 protein to ssDNA differs from that of E. coli SSB
and T4 gene 32 protein. Using a fluorescence based study gene 2.5 protein was found to have a binding constant for ssDNA binding of
1.2 × 106 M In the absence of DNA, gene 2.5 protein aggregates to form a stable
homodimer in solution (8). Dimer formation is postulated to be
dependent upon the interactions of its highly acidic carboxyl terminus
(21). Its association with the other replication proteins is also
facilitated by its carboxyl-terminal amino acids (13, 21). A similar
role has also been shown for the acidic carboxyl-terminal tail found in
the E. coli SSB protein (22) and the bacteriophage T4 gene
32 protein (23, 24).
The recently solved crystal structure of a carboxyl-terminal truncation
of gene 2.5 protein to a resolution of 1.9 Å (16) revealed a core that
consists of a conserved oligosaccharide/oligonucleotide binding fold
(OB fold) (Fig. 1), a structure common to other ssDNA-binding proteins
(25). As the name suggests this fold is found in proteins that bind
either ssDNA such as E. coli SSB protein (26, 27), human
mitochondrial SSB protein (28, 29), all three subunits of human
replication protein A (30-32), and staphlocococcal nuclease (33) or
oligosaccharides as found in E. coli heat liable enterotoxin (34). The OB fold is comprised of a five-stranded anti-parallel In the current study we have sought experimental evidence to support
our hypothesis that the DNA binding domain lies within the OB fold of
gene 2.5 protein. A previously reported random mutagenesis screen of
gene 2.5 (35) uncovered a number of lethal mutations that lead to
alterations in amino acids that were predicted by the crystal structure
to interact with ssDNA. Here we show that both the highly conserved
aromatic residue Tyr158, one component of the trinucleotide
binding motif, and the positively charged Lys152, flanking
this motif, are required for the interaction with ssDNA. In addition we
provide evidence for the involvement of Lys109 in the
protein-DNA interaction. Finally, we show that the carboxyl-terminal tail deleted protein, gp2.5- Materials
Bacterial Strains, Bacteriophage, and Plasmids--
E.
coli strains BL21(DE3) and HMS 174(DE3), which contain a DNA and Oligonucleotides--
M13 mGP1-2 is a 9950-nucleotide
derivative of vector M13 mp8 containing an insert of phage T7 DNA (37)
and was provided by S. Tabor (Harvard Medical School). BCMP 206, a
25-mer oligonucleotide with the sequence
5'-TAACGCCAGGGTTTTCCCAGTCACG-3', was synthesized by the Biopolymer
Facility at Harvard Medical School. The 70-mer and 38-mer
oligonucleotides used in the electrophoretic mobility shift assay for
assessing ssDNA binding had the sequences
5'-GACCATATCCTCCACCCTCCCCAATATTGACCATCAACCCTTCAC CTCACTTCACTCCACTATACCACTC-3' and 5'-CCTTTAAGTTCAAATGCTGCGC
GTCTTTCCAAGACAAG-3', respectively. These oligonucleotides were both
synthesized and gel purified by Oligos Etc. Inc. For cloning
purposes the following oligonucleotides were purchased from Oligos
Etc. Inc. T7 2.5 BamHI 5'-CGTAGGATCCACTTAGAAGTCTCCGTC-3' and T7 2.5 NdeI
5'-CGTAGGATCCATATGGCTAAGAAGATTTTCACCTC-3'. The oligonucleotides, pET17b
upstream, 5'-CTTTAAGAAGGAGATATACATATG-3' and pET17b downstream,
5'-GCTTCCTTTCGGGCTTTG-3', used to sequence the Y111C/Y158C construct
were obtained from Integrated DNA Technologies. All radioactive
nucleotides were purchased from Amersham Biosciences.
Proteins, Enzymes, and Chemicals--
All restriction enzymes as
well as T4 polynucleotide kinase, T4 DNA ligase, and calf intestinal
phosphatase were purchased from New England Biolabs. S. Tabor provided
E. coli SSB protein and T7 DNA polymerase- Methods
Mutagenesis of T7 Gene 2.5--
pET17(b) plasmids expressing
mutated gene 2.5, which lead to the alterations gp2.5-Y158C,
gp2.5-K109I, and gp2.5-K152E, were isolated from a genetic screen for
lethal mutants of gene 2.5 as previously described (35). To express the
variants gp2.5-Y111C/Y158C and gp2.5-Y111C separate vectors were
constructed using Stratagene's QuikChangeTM site-directed
mutagenesis kit. Initially both pET17(b) expressing wild-type gene 2.5 and pET17(b) expressing gp2.5-Y158C were amplified with
oligonucleotides
5'-CCTTTAAGTTCAAATGCTGCGCGTCTTTCCAAGACAAG-3' and
5'-CTTGTCTTGGAAAGACGCGCAGCATTTGAACTTAAAGG-3' (sequence
changes from wild-type T7 gene 2.5 have been underlined) using the
polymerase chain reaction. The resulting constructs were used to
transform XL1-Blue super-competent E. coli cells
(Stratagene). The oligonucleotides were synthesized and gel purified by
Oligos Etc. Inc.
Expression and Purification of Gene 2.5 Proteins--
Ten liters
of E. coli BL21(DE3) cells expressing wild-type gene 2.5 protein from pET17b were grown to an A595 of 1.0 in a Bioflo 2000 fermentor, in the presence of 6 µg/ml ampicillin. They were then induced with isopropyl- Expression and Purification of Gene 2.5 Histidine Fusion
Proteins--
E. coli BL21(DE3) competent cells were
transformed with pET19b 2.5PPS, pET19b 2.5PPS-Y158C, pET19b
2.5PPS-K109I, pET19b 2.5PPS-K152E, pET19b 2.5PPS-Y111C/Y158C, and
pET19b 2.5PPS In Vivo Complementation Assays--
Each variant protein was
analyzed for its ability to support bacteriophage T7 growth in
vivo. Electrocompetent E. coli HMS 262 cells were
transformed with pETGP2.5 plasmids containing each individual mutation.
Cells expressing wild-type and mutant gene 2.5 proteins were infected
with T7 bacteriophage lacking gene 2.5, and overlaid in soft agar onto
LB plates. The number of plaques formed on each plate was counted after
a 4-h incubation at 37 °C. The total amount of plaques produced by
cells expressing wild-type gene 2.5 protein was represented as a
plating efficiency of one. Therefore, plating efficiencies of the
variant proteins could be determined by comparison with wild-type.
Those mutations that could not substitute for wild-type gene 2.5 protein in vivo were examined for their ability to inhibit
the growth of wild-type bacteriophage T7. E. coli HMS 89 cells expressing each variant protein from the pET17(b) vector were
infected with wild-type bacteriophage T7 using the standard protocol.
The formation of plaques was examined after an overnight incubation at
37 °C.
Molecular Weight Approximation by Gel Filtration
Analysis--
The molecular weight of the gene 2.5 mutant proteins in
solution was approximated by gel filtration analysis on a Superdex 75 column (Amersham Biosciences) (8). Gel filtration was performed at
4 °C in buffer G at a flow rate of 0.6 ml/min. The column was initially calibrated using the following protein standards: ovalbumin (43 kDa), ribonuclease A (13.7 kDa), chymotrypsinogen (25 kDa), and
bovine serum albumin (67 kDa) (Amersham Biosciences). The elution
volumes of blue dextran and xylene cyanol determined the void volume
(V0) and total volume (Vt) of
the column, respectively. Five hundred µl of 0.2 µg/µl gene 2.5 variants, diluted in buffer S, were applied to the column and their
elution was monitored through spectrophotometric absorbance at 280 nm. The fractional retention, Kav, was calculated
for each of the standard proteins using the equation:
Kav = (Ve Surface Plasmon Resonance--
The interaction of gene 2.5 protein with T7 DNA polymerase was examined using surface plasmon
resonance (39). Initially the sensor-chip NTA (BIAcore) was activated
by passing 10 µl of running buffer (100 mM Hepes-NaOH, pH
7.5, 50 µM EDTA, 0.1 mM DTT, 100 mM NaCl) containing 0.5 mM NiCl2
over its surface at a rate of 10 µl/min. Histidine-tagged wild-type
gp2.5, gp2.5-Y158C, gp2.5-K109I, gp2.5-K152E, gp2.5-Y111C/Y158C, and
gp2.5- Affinity Chromatography--
The ability of wild type and
altered gene 2.5 proteins to physically interact with the 63-kDa form
of gene 4 protein was assessed by affinity chromatography as described
previously (9). Briefly, all proteins were dialyzed against 100 mM Hepes/NaOH, pH 7.5, 0.1 mM DTT, 0.5 mM EDTA, and 10% glycerol. Affi-Gel 15 (Bio-Rad) was
prepared according to the manufacturers instructions. Gene 2.5 protein
affinity columns were made by binding 1 mg of wild-type or altered gene
2.5 protein to 500 µl of Affi-Gel 15. Gene 4 protein (100 µg) was
passed over the column. The column was washed with 5 ml of the dialysis
buffer, then eluted in a step gradient (0-250 mM NaCl).
One-ml fractions were collected, and the amount of protein eluted from
the columns was monitored by absorbance at 280 nm.
Electrophoretic Mobility Shift Assay--
The ssDNA binding
ability of purified gene 2.5 proteins was assessed on both a 38- and
70-base length oligonucleotide using an electrophoretic mobility shift
assay (40). The oligonucleotides were 5'-end labeled with
[ Annealing Assay--
The ability of gene 2.5 protein to mediate
homologous base pairing was assayed for each altered gene 2.5 protein
using circular M13 ssDNA and a 32P-labeled linear
single-stranded fragment of M13. The labeled substrate was prepared by
initially annealing 1 µM "BCMP 206" oligonucleotide
to 0.13 µM mGPI-2 M13 in the presence of 50 mM NaCl and 25 mM Tris-Cl, pH 7.5, at 55 °C.
The primer was then partially extended using an exonuclease-deficient
T7 DNA polymerase in the presence of 8 µM dATP, dCTP,
dGTP, and dTTP, 3 µCi/µl [
Annealing of 0.09 nM of the 310-nt fragment to 2 nM circular M13 single-stranded DNA (molar ratio = 1:22) was assayed under the following conditions: 20 mM
Tris-Cl, pH 7.5, 1 mM DTT, 10 mM
MgCl2, and 50 mM NaCl, in the presence of
increasing concentrations (up to 12 µM) of wild-type and
altered gene 2.5 proteins. The reaction was incubated at 30 °C for 8 min prior to running on a 0.8% native agarose gel for 2 h at 80 V. Gels were dried under vacuum and the DNA was visualized by exposing
the gel to a Fujix phosphorimaging plate and quantitated using
ImageQuant software. For those proteins that mediated the annealing of
DNA under these conditions, i.e. wild-type gene 2.5 protein
and gp-K109I, a time course was obtained based on a 30-s interval from
0 to 7 min. To terminate the annealing activity at each time point 5 µl of stop solution (0.25% bromphenol blue, 0.25% xylene cyanol FF, 30% glycerol, and 0.5% SDS) was added to the sample.
Essential Residues in the Proposed ssDNA Binding Site--
In a
separate report (35) we described a random mutagenesis and genetic
selection to obtain lethal mutants in the cloned T7 gene 2.5. In the
present study we biochemically characterize the altered gene 2.5 proteins that have amino acid changes located in the region of the
protein predicted by the crystal structure to interact with ssDNA (16).
The location of these residues on the crystal structure of gp2.5- Amino Acid Alterations Do Not Disrupt Dimer Formation--
In
solution wild-type gene 2.5 protein forms a homodimer with molecular
weight of 51,124. One model for dimerization proposes that the
carboxyl-terminal tail of one protomer interacts with the predicted
ssDNA binding groove of another (16). Furthermore, stable dimer
formation has been shown to involve protein-protein interactions
between specific residues along the dimer interface (35). To ascertain
whether these altered proteins selected for this study were misfolded,
we assessed their ability to promote these interactions and form
dimers. Using gel filtration analysis to approximate their molecular
weights we initially calibrated a Superdex 75 column using four
standard proteins: ovalbumin, chymotrypsinogen, ribonuclease, and
bovine serum albumin. From their individual elution volumes a standard
curve was generated (Fig. 2) for the
estimation of the molecular weights of the gene 2.5 variant proteins.
Wild-type gene 2.5 protein displayed a fractional retention
(Kav), equal to 0.078. This value corresponds to
an estimated molecular weight of 47,000. Gp2.5-Y158C, gp2.5-K109I, and
gp2.5-Y111C/Y158C eluted in volumes consistent with wild-type gene 2.5 protein with a comparable molecular weight of 47,000 for each protein.
Gp2.5-K125E eluted slightly later than the native protein with its
molecular weight calculated to be 46,000. From this analysis we
conclude that each protein retains the ability to form homodimers in
solution, strongly suggesting that the overall structure of these
proteins is not affected by the amino acid substitutions.
Interaction of Gene 2.5 Proteins with Other T7 Replication
Proteins--
Gene 2.5 protein physically and functionally interacts
with T7 DNA polymerase, an interaction that is dependent on its
carboxyl-terminal amino acids (9, 17, 21). To further establish whether
the mutations affected the integrity of the protein, the interaction of
each altered gene 2.5 protein with the T7 DNA polymerase was examined
using surface plasmon resonance. In these experiments histidine-tagged
gene 2.5 proteins were immobilized on the chip surface and then T7 DNA
polymerase was flowed over the chip. The dissociation of the polymerase
from the bound gene 2.5 protein was monitored over a 10-min period. A
typical wild-type gene 2.5 binding curve is presented in Fig.
3A. Binding curves consistent with that of the native protein were obtained for mutants gene 2.5 protein-Y158C, gp2.5-Y111C/Y158C, gp2.5-K109I, and gp2.5-K152E (Fig.
3B, bottom). This result demonstrates that the amino acid substitutions do not disrupt the ability of the altered proteins to
physically interact with T7 DNA polymerase. Because this specific interaction is mediated by the carboxyl terminus of gene 2.5 (21), we
also examined the ability of the truncated protein, gp2.5-
Next we assessed the physical interaction between the wt and altered
gene 2.5 proteins and the 63-kDa form of the gene 4 helicase/primase protein. Previous studies have shown that these proteins interact physically and functionally (7, 9). The interaction is weaker than that
observed with T7 DNA polymerase and cannot be detected using surface
plasmon resonance.3
Therefore, we examined the interaction of gp2.5 with gene 4 protein using affinity chromatography. The 63-kDa gene 4 protein binds to a wt
gp 2.5 affinity column and elutes over a broad range of salt
concentrations (100 to 250 mM NaCl) (data not shown).
Similarly, the 63-kDa gene 4 protein stably binds to gp2.5-K109I,
gp2.5-K152E, gp2.5-Y158C, or gp2.5-Y111C/Y158C immobilized on a
column. Gene 4 protein also elutes from these columns over a broad
range of salt concentrations (50-250 mM NaCl) (data not
shown). We conclude that the altered proteins interact with the 63-kDa
gene 4 protein in a manner similar to wt gp2.5.
Binding of Gene 2.5 Protein to ssDNA--
In an earlier study we
assessed the binding of wild-type gene 2.5 protein to ssDNA using
either a nitrocellulose filter binding assay or by measuring
fluorescence quenching (8). In the present study we have employed an
electrophoretic mobility shift assay, using radioactively labeled
oligonucleotides of 38 and 70 nucleotides in length. In the experiment
shown in Fig. 4A a fixed
amount (3.3 nM) of the 33P-labeled
38-nucleotide oligonucleotide (38-mer) was incubated with increasing
amounts of wild-type gene 2.5 protein in the presence of 15 mM MgCl2. The 38-mer and
38-mer·protein complexes were then resolved by electrophoresis
through a nondenaturing polyacrylamide gel. The first detectable shift
for the wild-type gene 2.5 protein binding to the 38-mer occurs at a
protein concentration of 1330 nM. Using the Langmuir
isotherm the dissociation constant (Kd) was
calculated to be 7.9 × 10
Several of the genetically altered gene 2.5 proteins clearly have a
reduced ability to bind to the 38-mer oligonucleotide. Gp2.5-Y158C
binds to the 38-mer only at the highest concentration of protein
tested, thus requiring almost 10-fold more protein than the native
protein (Fig. 4B). As expected, a similar pattern of
diminished binding is also observed with the gene 2.5 protein containing the two amino acid substitutions Y111C/Y158C (Fig. 4C). Likewise gp2.5-K152E (Fig. 4D) has a lower
affinity for the 38-mer, and more protein is required to obtain a band
shift relative to the wild-type protein. Although gp2.5-K109I is not
able to support T7 growth and the amino acid change lies within the OB fold the K109I alteration has no detectable affect on binding to the
ssDNA (Fig. 4E) with a Kd comparable with
that of wild-type gene 2.5 protein. Interestingly, the truncated form of the protein, gp2.5-
The binding of the altered gene 2.5 proteins to a 70-mer
oligonucleotide was then assessed to determine whether the length of
ssDNA influenced the ssDNA binding patterns of the variant protein.
Results obtained mimicked that found with the 38-mer except for the
appearance of two individual bands in the gel shift for a number of
these gene 2.5 proteins. Wild-type gene 2.5 protein initially binds to
the 70-mer at a protein concentration of 670 nM, where two
species of 70-mer·protein complexes are observed. As the amount of
gene 2.5 protein is increased the amount of unbound 70-mer decreases
such that by 2650 nM all of the 70-mer is found in the
slower migrating species (Fig.
5A). The Kd
was calculated to be 3.3 × 10 Homologous Base Pairing Mediated by Gene 2.5 Proteins--
Gene
2.5 protein facilitates the annealing of complimentary strands of
ssDNA,2 a property that has been used in preparing
substrate for studies on the strand transfer mediated by the T7
helicase (10, 11). We have measured the ability of the altered gene 2.5 proteins to facilitate this reaction to determine the effect of the
decreased affinity for ssDNA. Wild-type gene 2.5 protein is capable of
successfully annealing the homologous ssDNA at a concentration of 4 µM (Fig. 6A). A
time course experiment revealed that the wild-type gene 2.5 protein can
accomplish this base pairing after approximately 1 min incubation at
30 °C (Fig. 6B). The genetically altered gene 2.5 proteins defective in ssDNA binding do not anneal the two species of
ssDNA at the concentrations tested (Fig. 6, C-E).
Presumably this lack of activity is because of the decreased binding
affinity of these proteins, because we have observed a small percentage (<25%) of annealed products at high protein concentrations (data not
shown). Therefore, it is likely that the defect in annealing observed
for these proteins is a reflection of the decreased affinity for ssDNA.
Interestingly, whereas gp2.5-K109I shows an equal affinity for ssDNA in
the electrophoretic mobility shift assay, it is not as efficient in
annealing homologous ssDNA requiring twice as much protein (Fig.
6F), and taking 1 min longer to form the annealed product
(compare Fig. 6, G-B).
There is a significant lack of knowledge on the structure-function
relationship of the product of bacteriophage T7 gene 2.5, a
ssDNA-binding protein. In the current study we sought to define the DNA
binding domain of gp2.5. This study provides the first experimental
evidence for the identity of the DNA binding domain of gp2.5. In a
separate report (35) we describe a genetic screen for lethal mutations
in bacteriophage T7 gene 2.5. By examining the crystal structure of the
protein (16) we noted that a subset of these generated mutations lay in
the postulated ssDNA binding domain. In this study we have
biochemically characterized these altered gene 2.5 proteins and show
that two of them, gp2.5-Y158C and gp2.5-K152E, are indeed defective in
their interaction with ssDNA. A third, gp2.5-K109I, appears to interact
differently with ssDNA when assessed by an electrophoretic mobility
shift assay. We feel confident that the defective phenotypes do not
arise from a misfolding of the protein as they physically interact with
both T7 DNA polymerase and the gene 4 protein, a helicase/primase. In
addition, all four proteins form dimers in a manner similar to the
wild-type protein.
At the onset of these studies the amino acids involved in the
interaction of gene 2.5 protein with ssDNA were unknown. Previous studies on other ssDNA-binding proteins have shown that aromatic residues have the potential to intercalate between the nucleic acid
bases thus stabilizing the protein-ssDNA interaction (27, 41-43). In
the E. coli SSB protein for example mutational studies have
implicated phenylalanine 60 (Phe60) and tyrptophan 54 (Trp54) in binding ssDNA (44-46). Similarly, in T4 gene
32, protein site-directed mutagenesis has identified numerous tyrosine
residues necessary for the proteins interaction with ssDNA (47, 48). T7
gene 2.5 protein possesses one such structurally conserved aromatic residue found in the OB fold. Based on our observation that the lethal
substitution Y158C gives rise to a gene 2.5 protein that has a higher
dissociation constant for ssDNA than that of wild-type gene 2.5 protein, we can infer that this residue plays an essential role in
binding to ssDNA. Similarly, the altered gp2.5-Y111C/Y158C is also
defective in binding ssDNA although the binding constant is similar to
that of gp2.5-Y158C alone, implying that tyrosine 111 is not essential
for ssDNA binding. This result was unexpected as an aromatic residue is
conserved at this site among other ssDNA-binding proteins, specifically
phenylalanine 60 in E. coli SSB protein and phenylalanine 90 in human replication protein A protein (16).
Basic residues, which can make electrostatic contacts with the
negatively charged phosphates in ssDNA, are also likely candidates for
DNA binding. Lysine residues have been shown to be involved in the
ability of E. coli SSB protein to bind ssDNA (27, 49). Indeed we have shown that substituting lysine 152 with glutamic acid
weakens its binding to ssDNA, as this variant protein exhibits a
10-fold decrease in its affinity for ssDNA. Interestingly, the protein·DNA complex resulting from this interaction has difficulty adopting the slower mobility complex at concentrations comparable with
wild-type gene 2.5 protein, as discussed in more detail below. Perhaps
Lys152 is involved in an interaction of gene 2.5 with ssDNA
at higher concentrations, and this interaction leads to the higher
order structure in a manner similar to the E. coli SSB
protein, which demonstrates distinct binding modes at different protein
concentrations (18). In addition, the side chain of Lys152
is oriented away from the prominent groove in the crystal structure, decreasing its direct accessibility to ssDNA bound at this site. Therefore to implicate Lys152 in binding ssDNA, the ssDNA
would somehow have to wrap around the protein, suggesting that the
interaction encompasses more residues than those lying directly within
this groove. A structural based sequence alignment of other
ssDNA-binding proteins does not reveal conservation at this particular
residue (16) and therefore, perhaps this interaction is a unique
feature of the gene 2.5 protein.
A second lethal mutation resulting in the alteration of a basic residue
was identified at Lys109. In contrast to gp2.5-K152E, this
K109I variant gene 2.5 protein failed to inhibit the ability of gene
2.5 protein to bind ssDNA despite the loss of a positively charged
residue in the OB fold. Interestingly gp2.5-K109I displayed an aberrant
binding pattern in the gel shift assay, forming only the slower
mobility complex. This binding pattern was also exhibited by
gp2.5- Based on the crystal structure a model was proposed for DNA binding
that assumes that gene 2.5 binds ssDNA as a monomeric species (16). The
hypothesis is that the negatively charged, acidic carboxyl terminus
competes with the proposed DNA binding site of an adjacent protomer
leading to the formation of dimers in the absence of ssDNA. Therefore
dissolution of the dimer would be necessary to expose the DNA binding
domain and allow for ssDNA binding. In support of this model we have
shown how a carboxyl-terminal truncated form of the protein gp
2.5- The electrophoretic mobility shift assay employed in this study
provided an insight into the mode by which wild-type gene 2.5 protein
binds ssDNA. Over a protein concentration series from 80 to 10,600 nM, upon binding a 70-mer oligonucleotide, two distinct protein·DNA complexes were resolved. Conceivably these two complexes could represent the binding of one monomer of gene 2.5 protein and a
subsequent second monomer at a higher concentration. This interpretation is supported by the absence of the slower mobility complex when wild-type gene 2.5 protein binds to a shorter
oligonucleotide of 38 bases in length as presumably only one monomer
can be accommodated on this length. However, this hypothesis does not
agree with the published site size for the protein, which is seven
nucleotides per monomer (8). This site size was calculated by assessing the binding of gene 2.5 protein to circular M13 ssDNA. In a subsequent study using surface plasmon resonance, stable binding of gene 2.5 protein required an oligonucleotide of at least 30 nucleotides in
length.4 Nonetheless this unexplained phenomenon presented
by the distinct protein·DNA complexes warrants further study
especially because other ssDNA-binding proteins have demonstrated
different modes of binding ssDNA.
As gene 2.5 protein has the capacity to mediate homologous DNA
annealing2 we examined how efficiently the altered proteins
could accomplish this activity. The altered proteins deficient in
binding to ssDNA were also defective in annealing complimentary strands
of ssDNA. However, we do observe some annealing at higher protein
concentrations. For this reason, we feel the lack of annealing we
observed is related to the affinity of the altered proteins for ssDNA
rather than reflecting a defect in the basic mechanism of homologous base-pairing. Interestingly, gp2.5- All of the proteins described in this study were expressed from lethal
mutations in gene 2.5. It is likely that gp2.5-K152E, gp2.5-Y158C, and
gp2.5-Y111C/Y158C are lethal because they have a lower affinity for
ssDNA. Given that ssDNA binding is an essential function for gene 2.5 it is not surprising that amino acid changes that reduce binding
affinity in vitro are lethal in vivo. In
addition, we have shown that these proteins can still form dimers, and
physically interact with the T7 DNA polymerase and the 63-kDa gene 4 protein. The mechanism underlying gp2.5-K109I lethality, on the other
hand, remains unclear. In cells harboring a plasmid
encoding for this genetic alteration, we detected a reduced level DNA
synthesis in vivo upon infection by T726C, binds single-stranded DNA 10-fold more tightly
than the wild-type protein. The three altered proteins defective in
single-stranded DNA binding cannot mediate the annealing of homologous
DNA, whereas gp2.5-
26C mediates the reaction more effectively than
does wild-type. Gp2.5-K109I retains this annealing ability, albeit
slightly less efficiently. With the exception of gp2.5-
26C, all
variant proteins form dimers in solution and physically interact with
T7 DNA polymerase.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1 (8) a value
that is less than one-tenth the affinity exhibited by E. coli SSB (18) and T4 gene 32 protein (19, 20). In addition, gene
2.5 protein binds ssDNA with limited, if any, cooperativity (8). Kim
et al. (8) reported that gene 2.5 protein bound to ssDNA
with a stoichiometry of 7 nucleotides per monomer of protein, although
it is not known if gene 2.5 protein binds to ssDNA as a monomer or dimer.
barrel, capped with an
helix. The location of this helix varies among DNA-binding proteins. In gene 2.5 protein this helix is found
between the second and third strands (16), whereas in both human
replication protein A and E. coli SSB protein it connects the third and forth strands (27, 30). In gene 2.5 protein loop
extensions from two of the
sheets form a prominent groove on the
surface of the fold. This groove most likely represents the ssDNA
interface and in Fig. 1 ssDNA is modeled into the crystal structure
along this position. Located within this groove of gene 2.5 protein are
two aromatic residues, tyrosine 111 and tyrosine 158. These aromatic
residues, in addition to the adjacent
stands and their connecting
loops comprise an evolutionarily conserved trinucleotide binding motif
(16) that binds three nucleotides in an orientation analogous with
other ssDNA-binding proteins (27, 30). A number of basic residues
(Lys3, Arg35, Lys107,
Lys109, Lys150, and Lys152) lie in
proximity to this trinucleotide binding motif forming a positively
charged cleft, suggesting a role in the interaction with ssDNA (16).
However, prior to the current study, there was no direct evidence that
implicates these residues in binding ssDNA.
26C, binds DNA with a greater affinity than the wild-type protein.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
prophage encoding the T7 RNA polymerase gene under the control of the
lac promoter (Novagen), were used as host strains for protein overexpression and purification. Wild-type and mutant forms of T7 gene
2.5 are expressed from the pET17b plasmid (Novagen) containing the T7
RNA polymerase promoter. DNA encoding His-tagged gene 2.5 proteins were
subcloned into the NdeI and BamHI restriction
sites of a modified pET19b vector (Novagen). The enterokinase site of this plasmid was replaced with a rhinovirus C protease (PreScission protease, Amersham Biosciences) site located upstream of the
start codon. T. Biswas (Harvard Medical School) provided this plasmid. E. coli strains HMS 262 and HMS 89 and all T7 bacteriophage
are from our laboratory collection. Growth and manipulation of
bacteriophage T7 and E. coli were performed as described
previously (36).
28. D. Johnson
and J. Lee (Harvard Medical School) supplied wild-type T7 DNA
polymerase. T7 gp2.5-
26C and His-tagged gp2.5-
26C were obtained
from E. Toth (Harvard Medical School) and J. Stattel (Harvard Medical
School), respectively. Gene 4 protein was kindly provided by D. Crampton (Harvard Medical School). All chemicals and reagents were from
Sigma unless otherwise noted.
-D-galactoside at
a concentration of 1 mM. After 4 h the cells were
harvested by centrifugation at 2,000 rpm for 40 min, resuspended in 250 ml of lysis buffer (50 mM Tris-Cl, pH 7.5, 10% sucrose,
0.1 mM EDTA), and frozen on dry ice. Prior to purification
the cells were thawed at 4 °C overnight in the presence of 50 mM
-mercaptoethanol. Cell lysis was accomplished by the
addition of lysozyme at a final concentration of 0.5 mg/ml (diluted in
lysis buffer) followed by incubation at 4 °C for 45 min. Lysed cells
were heated to 20 °C in a 37 °C water bath, then chilled on ice
and centrifuged at 100,000 × g for 45 min at 4 °C.
To precipitate T7 gene 2.5 protein, polyethylenimine, at a final
concentration of 0.1%, was added to the supernatant and the solution
incubated at 4 °C for 1 h. The suspension was centrifuged at
21,000 × g for 15 min and the resulting pellet was
resuspended in 90 ml of buffer A (50 mM Tris-Cl pH 7.5, 0.1 mM EDTA, 1 mM DTT, 10% glycerol) containing 1 M NaCl. This suspension was rotated at 4 °C for 1 h
then centrifuged at 21,000 × g for 15 min. The
resulting supernatant (~70 ml) was diluted with buffer A to 175 ml
and 84 g of (NH4)2SO3 was
added slowly over a 1-h time period. The suspension was stirred for
another hour at 4 °C followed by centrifugation at 21,000 × g for 15 min. The protein pellet was resuspended in 75 ml of
buffer A and filtered through a 0.22-µm bottle top filter. This
suspension was loaded onto a POROUS HQ column (PE Biosystems) and T7
gene 2.5 protein eluted in a 50 mM to 1 M NaCl
gradient with most of the protein eluting at ~550 mM
NaCl. The eluted protein was precipitated by the addition of
(NH4)2SO3 to 85% saturation and
the protein was collected by centrifugation at 21,000 × g for 15 min. The resulting pellet was resuspended in 500 µl of buffer G (50 mM potassium phosphate buffer, pH 7.0, 150 mM NaCl, 0.1 mM EDTA, 0.1 mM
DTT, 10% glycerol) and loaded onto a Superose 12 size exclusion column
(Amersham Biosciences). T7 gene 2.5 protein eluted from the column in
~15 ml of buffer G and was then dialyzed into buffer S (50 mM Tris-Cl, pH 7.5, 0.1 mM EDTA, 1 mM DTT, 50% glycerol). The protein was over 99% pure as
determined by SDS-polyacrylamide electrophoresis and subsequent
staining by Coomassie Blue. Gp2.5-Y158C, gp2.5-K152E, and gp2.5-K109I
were purified as described for wild-type. It was necessary to express
mutant gp2.5-Y111C/Y158C in E. coli strain HMS174 (DE3)
pLys-S (Novagen) and it was purified from a 4-liter culture. Protein
concentrations were determined by spectrophotometric absorbance at 280 nm using the extinction coefficient of the protein calculated according
to Gill and von Hippel (38).
26C. One-liter cultures were grown of each in LB media
containing ampicillin. They were induced and harvested as described
previously and the pelleted cells were resuspended in 20 ml of buffer B
(50 mM Tris-Cl, pH 7.5, 500 mM NaCl, and 70 mM imidazole). Following one freeze-thaw cycle the
cells were lysed in the presence of 0.5 mg/ml lysozyme by incubating
4 h at 4 °C. To degrade the E. coli DNA, 125 units of Benzonase nuclease (Novagen) was added to the suspension and the suspension was heated to 20 °C in a 37 °C water bath. The cell debris was collected by centrifugation at 8,000 × g for 40 min at 4 °C. The cleared lysates were introduced
onto a nickel-nitrilotriacetic-agarose column (Qiagen) with a bed
volume of 5 ml. The resin was washed with 20 column volumes of buffer B
and the protein was eluted in 20 ml of buffer B containing 500 mM imidazole. Each protein was then dialyzed against
buffer S and stored at
20 °C.
Vo)/(Vt
Vo), where Ve is the peak elution
volume of each protein. The molecular weight of each gene 2.5 variant
was approximated using a standard curve generated by plotting the Kav value versus log10
Mr.
26C were diluted in running buffer supplemented with 500 nM bovine serum albumin. To immobilize gene 2.5 proteins,
10 µl of the protein solution was injected onto individual lanes on
the surface of the chip at a rate of 10 µl/min. Running buffer was
passed over the chip for 2 min. At this time an increase in resonance
units of ~7,000 on the surface of the chip was noted, thus
establishing a baseline. Ten µl of various concentrations (up to 500 nM) of T7 DNA polymerase were injected onto the chip over 1 min, followed by 10 min of buffer. The association and dissociation of
T7 DNA polymerase with gene 2.5 protein was monitored during the
experiment by noting the change in resonance unit value. All proteins
were removed and the chip was regenerated by passing 20 µl of running buffer containing 350 mM EDTA over its surface.
-33P]ATP using polynucleotide kinase at 37 °C for
2 h and purified using Bio-Rad micro-biospin column P-30.
Oligonucleotide (3.3 nM) was incubated for 15 min on ice
with increasing concentrations (up to 10.6 µM) of
purified wild-type gp2.5, gp2.5-Y158C, gp2.5-K109I, gp2.5-K152E, and
gp2.5-Y111C/Y158C. All proteins were diluted in 20 mM
Tris-Cl, pH 7.5, 10 mM
-mercaptoethanol, 500 µg/ml
bovine serum albumin. Final concentrations of the components (in 15 µl) were 15 mM MgCl2, 5 mM DTT,
50 mM KCl, 10% glycerol, and 0.01% bromphenol blue.
Samples were loaded onto 10% TBE pre-cast gels (Bio-Rad) and run at 80 V for 2 h at 4 °C using 0.5× Tris glycine running buffer (12.5 mM Tris base, 95 mM glycine, 0.5 mM
EDTA). Gels were dried, exposed to a FujiX phosphorimaging plate, and the fraction of DNA bound by gene 2.5 protein was measured using ImageQuant software. This measurement facilitated the calculation of
the dissociation constants (Kd) for each protein
using the Langmuir isotherm formula.
-32P]dGTP, 5 mM DTT, 2 µM bovine serum albumin, and 10 mM MgCl2, at room temperature for 10 min. To
fully extend the primer 80 µM dATP, dGTP, dCTP, and dTTP
were then added and the reaction was incubated for 15 min at room
temperature. Reactions were then incubated for a further 10 min at
70 °C to denature the polymerase. Next 55 nM E. coli SSB protein was added and the DNA was digested with Acc65
I for 2 h at 37 °C. DNA was extracted with 50 µl of phenol:chloroform:isoamyl alcohol (25:24:1) and then separated from
unincorporated nucleotides using a MicroSpin S-400HR column (Amersham
Biosciences). To produce ssDNA fragments, the DNA was denatured with
100 mM NaOH at 25 °C for 5 min followed by
neutralization on ice with 100 mM HCl and 100 mM Tris, pH 7.5. Separation of the fragments was achieved
by gel electrophoresis on a 1.4% agarose gel run at 50 V for 90 min.
The radioactive band corresponding to the Acc65 1 digested a
310-nt fragment that was extracted from the gel and purified using the
gel extraction kit from Qiagen.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
26C
is shown in Fig. 1 and the phenotype of
these mutants is summarized in Table I.
The aromatic residues tyrosine 158 and tyrosine 111, structurally conserved among other ssDNA-binding proteins, lie in the core of the OB
fold and comprise the trinucleotide-binding motif (16). The
screen identified a single lethal mutation, Y158C. A mutation leading
to an amino acid change at tyrosine 111, on the other hand, was not
found alone, but rather in a clone containing two other mutations (35).
Thus, site-directed mutagenesis was used to generate a gene 2.5 protein
with a Y111C substitution. However, a plasmid expressing gp2.5-Y111C
was shown to retain the ability to support the growth of T7 phage
lacking wild-type gene 2.5 protein (Table I) and was not characterized
further. A double mutant generated with the substitutions Y111C and
Y158C was found to be unable to support the growth of T7
2.5 phage.
In addition, the original screen detected independent lethal mutations
at two lysine residues resulting in variants K109I and K152E. As well as being positively charged, both residues lie within the
barrel of
the OB fold and flank the trinucleotide binding motif making them
likely candidates to interact with ssDNA. None of these mutations could
complement the growth of T7
2.5, nor were they dominant lethal, as
they could not suppress the growth of wild-type T7 bacteriophage (Table
I).
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Fig. 1.
Proposed model for ssDNA binding in the
crystal structure of T7 gene 2.5 protein and location of the altered
amino acids. The crystal structure of gp2.5- 26,
determined at 1.9-Å resolution (16). The five-stranded anti-parallel
-barrel, depicted in purple, capped by a
-helix
(gray), comprises the conserved
oligosaccharide/oligonucleotide binding fold. Single-stranded DNA
(pink) is modeled in the structure at the predicted binding
site. The side chains of the residues characterized in this study are
highlighted in green and their position in the polypeptide
note.
Ability of gene 2.5 proteins to complement the growth of T72.5 phage
or suppress the growth of wild-type T7 phage
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Fig. 2.
Estimation of the molecular weight of
wild-type and variant gene 2.5 proteins by gel filtration
analysis. Gel filtration was performed using a Superdex 75 column
as described under "Experimental Procedures." The protein
standards, ovalbumin (43 kDa), chymotrypsinogen (25 kDa), bovine serum
albumin (67 kDa), and ribonuclease A (13.7 kDa), were used to calibrate
the column. The elution volumes of blue dextran and xylene cyanol
determined the void volume and total volume of the column,
respectively. A plot of Kav versus
the log Mr of the standard proteins was
generated and the best-fit line was determined. Wild-type gene 2.5 protein, gp2.5-K109I, gp2.5-K152E, gp2.5-Y158C, and gp2.5-Y111C/Y158C
were applied to the column in three independent experiments. The
Kav for each variant was calculated based on
their elution volumes. Their positions on the standard curve are
noted.
26C, that
lacks the 26 carboxyl-terminal amino acids to interact with the T7 DNA
polymerase (Fig. 3B, top). As expected, this
protein did not appreciably bind the polymerase. These findings,
combined with the gel filtration results presented above, increase our confidence that the variant proteins are not grossly misfolded.
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Fig. 3.
Interaction of gene 2.5 proteins with T7 DNA
polymerase. Surface plasmon resonance was used to detect the
interaction between the gene 2.5 proteins and T7 DNA polymerase. The
surface of the chip was activated by saturating the nitrilotriacetic
sites with running buffer (100 mM Hepes-NaOH, pH 7.5, 50 µM EDTA, 0.1 mM DTT, 50 mM NaCl)
containing 0.5 mM NiCl2. Five nmol of the
histidine-tagged gene 2.5 proteins were then immobilized on the chip
surface. Following a period of stabilization, 5 nmol of DNA polymerase
was passed over the chip. Its association and dissociation with the
gene 2.5 proteins was monitored for 10 min through changes in the
refractive index, measured in arbitrary resonance units
(RU). A, an overlay plot where either 0 or 500 nM DNA polymerase was injected onto a nitrilotriacetic chip
in which wild-type gene 2.5 had been immobilized. The injection point
of T7 DNA polymerase and its dissociation period is noted.
B: top, an overlay plot depicting the interaction
of 500 nM T7 DNA polymerase with either wild-type gp2.5
(solid line) or gp.2.5 26C (dashed line)
immobilized to the chip. Bottom, an overlay plot depicted
the interaction of 500 nM T7 DNA polymerase with
gp2.5-Y158C (closed circles), gp2.5-K152E (dashed
line), gp2.5-Y111C/Y158C (solid line), or gp2.5-K109I
(dotted line).
6 M. The
Kd values are listed in Table
II for the wild-type protein as well as
for the altered gene 2.5 proteins discussed below.
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Fig. 4.
Binding of gene 2.5 proteins to a
38-nucleotide oligonucleotide. An electrophoretic mobility shift
assay was employed to assess the binding of the gene 2.5 proteins to
ssDNA. 3.3 nM 5'-33P-Labeled oligonucleotide
was incubated for 15 min on ice with increasing amounts of gene 2.5 proteins, as described under "Experimental Procedures." Reaction
products were resolved on a 10% nondenaturing polyacrylamide gel. The
proteins examined were wild-type gene 2.5 protein (wt) (A),
gp2.5-Y158C (Y158C) (B), gp2.5-Y111C/Y158C (Y111C/Y158C)
(C), gp2.5-K152E (K152E) (D), gp2.5-K109I (K109I)
(E), and gp2.5- 26C (F).
Dissociation constants of gene 2.5 proteins to ssDNA
26C, binds much tighter to the 38-mer compared with the native protein (Fig. 4F) with a corresponding
Kd that is 10-fold lower.
6 for the more rapidly
migrating complex and 5.4 × 10
7 for the slower
migrating complex. No bandshift was observed in the case of gp2.5-Y158C
until a protein concentration of 2650 nM was reached, which
was 4-fold the concentration required for the native protein (Fig.
5B). Furthermore, the slower migrating complex is observed
only at the highest concentration of protein. The Kd
values for both complexes are ~10-fold higher than that observed with
the wild-type protein (Table II). The variant gp2.5-Y111C/Y158C has a
similar affinity to that of gp2.5-Y158C (Fig.
5C) but with this altered protein the slower migrating
complex is never observed. In addition gp2.5-K152E has a lower affinity for the 70-mer and considerably higher concentrations of this altered
protein are required to bind all of the oligonucleotide and to achieve
the slower migrating complex (Fig. 5D and Table II). As with
the 38-mer, gp2.5-K109I appears to bind to the 70-mer with the same
affinity as the native protein. However, no rapidly migrating complex
is observed with gp2.5-K109I (Fig. 5E). We have also
examined the binding of the altered protein gp2.5-
26C, where the
protein is found to have a higher affinity for the 70-mer and, like
gp2.5-K109I, fails to produce the rapidly migrating complex (Fig.
5F).
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Fig. 5.
Binding of gene 2.5 proteins to a
70-nucleotide oligonucleotide. The experiment in Fig. 4 was
repeated with a 70-mer oligonucleotide and the same abbreviations are
used.
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Fig. 6.
Homologous base pair annealing facilitated by
gene 2.5 proteins. Circular M13 ssDNA was incubated with
a 310-nt 32P-labeled complimentary ssDNA fragment, in the
presence of increasing concentrations of gene 2.5 proteins as described
under "Experimental Procedures." The annealing products were
fractionated on a nondenaturing agarose gel and visualized by
autoradiography. A, a concentration series of wild-type gene
2.5 protein illustrating the ability of this protein to anneal a 310-nt
fragment to M13 ssDNA. B, a time course experiment using 8 µM wild-type gene 2.5 protein. Time points were taken at
30-s intervals and the reaction was terminated by adding 5 µl of stop
solution (0.25% bromphenol blue, 0.25% xylene cyanol FF, 30%
glycerol, and 0.5% SDS). C-E, annealing reactions were
performed using increasing amounts of gp2.5-Y158C (C),
gp2.5-K152E (D), and gp2.5-Y111C/Y158C
(E). F, the annealing activity of gp2.5-K109I was
assessed over a range of protein concentrations. G, the rate
of homologous base pair annealing of a 310-nt fragment to M13 ssDNA
mediated by gp2.5-K109I (16 µM). Reactions were performed
as described above.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
26C, which likewise failed to form the rapidly migrating
complex. Originally this binding pattern was thought to indicate a
level of cooperative binding to ssDNA not characteristic of the native
protein. However, upon closer examination, the appropriate kinetic
calculations, i.e. Hill coefficients, could not support this
theory. Further dissection of the binding mode(s) of the native protein
may lead to an explanation of this observation.
26C that exists as a monomer in solution (35) has a 10-fold
greater affinity for ssDNA as compared with wild-type gene 2.5 protein.
Similar results have been seen with gp2.5-
21C, also a monomer in
solution, where ssDNA binding was analyzed by surface plasmon
resonance.4
26C facilitates strand annealing more efficiently than the wild-type protein,3 further
supporting a relationship between ssDNA binding affinity and homologous
base pair annealing. Furthermore, despite its unaltered affinity for
ssDNA the variant protein K109I is defective in base pairing, requiring
2-fold more protein to completely anneal all the substrate. We are
currently pursuing the mechanism of this reaction by studying another
altered protein that binds ssDNA but cannot facilitate the annealing of
homologous strands of ssDNA.3 From the data presented here,
we conclude that gp2.5 must be able to bind ssDNA to facilitate DNA annealing.
2.5
bacteriophage,5 suggesting
that the function of this residue is important in the overall DNA
replication of the bacteriophage. It is conceivable that both the
impaired ability to facilitate annealing and the altered manner in
which it binds ssDNA accounts for the in vivo phenotype.
However, further investigation is necessary to probe the precise
molecular basis of its involvement in the life cycle of bacteriophage T7.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Tomas Hollis for preparation of Fig. 1, Stanley Tabor for assistance with the DNA annealing assay, and Don Crampton for critically reading the manuscript.
![]() |
FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grant GM54397-39 (to C. C. R.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Dept. of Biological
Chemistry and Molecular Pharmacology, Harvard Medical School, 240 Longwood Ave., Boston, MA 02115. Tel.: 617-432-1864; Fax: 617-432-3362;
E-mail: ccr@hms.harvard.edu.
Published, JBC Papers in Press, December 20, 2002, DOI 10.1074/jbc.M210605200
2 S. Tabor and C. C. Richardson, unpublished data.
3 L. F. Rezende and C. C. Richardson, unpublished data.
4 J. M. Stattel and C. C. Richardson, unpublished data.
5 E. M. Hyland, L. F. Rezende, and C. C. Richardson, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: ssDNA, single-stranded DNA; OB fold, oligosaccharide/oligonucleotide binding fold; nt, nucleotide; DTT, dithiothreitol; gp, gene product.
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REFERENCES |
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