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INTRODUCTION |
ATP-dependent DNA ligases catalyze the joining of a
5'-phosphate-terminated strand to a 3'-hydroxyl-terminated strand via three sequential nucleotidyl transfer reactions (1, 2). In the first
step, nucleophilic attack on the
-phosphorus of ATP by ligase
results in release of pyrophosphate and formation of a covalent
intermediate (ligase-adenylate) in which AMP is linked via a
phosphoamide (P-N) bond to the
-amino group of a lysine. In the
second step, the AMP is transferred to the 5'-end of the
5'-phosphate-terminated DNA strand to form a DNA-adenylate intermediate, A(5')pp(5')N. In this reaction, a 5'-phosphate oxygen of
the DNA strand attacks the phosphorus of ligase-adenylate; the active
site lysine side chain is the leaving group. In the third step, ligase
catalyzes attack by the 3'-OH of the nick on the DNA-adenylate to join
the two polynucleotides and release AMP.
ATP-dependent ligases are ubiquitous in eukaryotes. They
are also encoded by certain eubacteria, bacteriophages, and eukaryotic DNA viruses. Sequence comparisons suggest that a catalytic domain common to all ATP-dependent ligases is embellished by
additional isozyme-specific protein segments at the N or C termini (3). The catalytic domain includes a set of six collinear motifs (I, III,
IIIa, IV, V, and VI) that define a superfamily of covalent nucleotidyl
transferases encompassing the ATP-dependent polynucleotide ligases and GTP-dependent mRNA capping enzymes (4, 5).
The crystal structure of T7 DNA ligase with bound ATP shows that the nucleotide binding pocket is composed of motifs I, III, IIIa, IV, and V
(6). The lysine in motif I (KXDGXR) is the site
of covalent attachment of AMP to the enzyme (7).
Crystallography and mutagenesis have illuminated enzymic functional
groups that are involved in nucleotide binding and nucleotidyl transfer
(6, 8-10). What remains obscure is the structural basis for DNA nick
recognition by DNA ligases. Conversion of nicks into phosphodiester
bonds is the common final step in the DNA repair and replication
pathways. Nicks are potentially deleterious DNA lesions that, if not
corrected, may give rise to lethal double-strand breaks. One imagines a
mechanism to ensure that ligases are directed to sites where their
action is required, either via the interaction of ligase with other
replication or repair proteins assembled at a nick (3) or via an
intrinsic capacity to discriminate nicks from other DNA structures.
We are examining the interaction of eukaryotic ligases with defined
DNAs using virus-encoded enzymes as models. Vaccinia virus DNA ligase
and Chlorella virus DNA ligase each forms a discrete complex
with a singly nicked DNA ligand in the absence of magnesium that can be
resolved from free DNA by native polyacrylamide gel electrophoresis
(11, 12). The viral ligases do not form stable complexes with the
following ligands: (i) DNA containing a 1- or 2-nucleotide gap, (ii)
the sealed duplex DNA product of the ligation reaction, (iii) a singly
nicked duplex containing a 5'-OH terminus at the nick instead of a
5'-phosphate, or (iv) a singly nicked duplex containing an RNA strand
on the 5'-phosphate side of the nick (10-15). These DNA ligases
apparently do have an intrinsic nick sensing function. Discrimination
at the substrate binding step may account for the feeble activity of
the eukaryotic viral DNA ligases in sealing across gaps and in joining
nicked molecules containing a 5'-phosphate-terminated RNA strand (14,
15). Nick recognition by vaccinia virus DNA ligase and
Chlorella virus DNA ligase also depends on occupancy of the
AMP binding pocket on the enzyme, i.e. mutations of the
ligase active site that abolish the capacity to form the
ligase-adenylate intermediate also eliminate nick recognition, whereas
a mutation that preserves ligase-adenylate formation, but inactivates
downstream steps of the strand joining reaction, has no effect on
binding to nicked DNA (10, 13).
To physically map the ligase-DNA interface, we sought to obtain
footprints of the ligase binding site on DNA and the DNA binding site
on ligase. We conducted this analysis using the 298-amino acid
Chlorella virus PBCV-1 DNA ligase, which is the smallest eukaryotic DNA ligase known and likely constitutes the minimal catalytic unit (12). A confounding technical factor in
nuclease-footprinting the DNA side of the ligase-DNA interface was the
requirement for a divalent cation cofactor at the nuclease digestion
step, the problem being that the ligase-adenylate, which binds to
nicked DNA in the absence of divalent cations, immediately catalyzes strand joining upon addition of a divalent cation and then dissociates from the closed duplex ligation product. The enabling factor in obtaining a nuclease footprint of the ligase-DNA complex was the finding that an alanine substitution for Asp-29 in motif I of Chlorella virus DNA ligase
(27KXDGXR32) blocks step
2 of the ligation pathway without affecting enzyme-adenylate formation
and nick recognition (10). This "step-arrest" mutant ligase remains
bound at the nick in the presence of magnesium; consequently, the
D29A-DNA complexes could be footprinted with exonuclease III. We report
that the ligase footprint extends from 8-9 nucleotides on the 3'-OH
side of the nick to 11-12 nucleotides on the 5'-phosphate side of the
nick. To examine the protein component of the ligase-DNA interface, we
used the classical approach of limited proteolysis to probe the
structure of Chlorella virus DNA ligase in the free and
DNA-bound states. The Chlorella virus ligase consists of two
protease-resistant domains connected by a protease-sensitive
interdomain segment located between motifs V and VI. Protection of the
interdomain segment from proteolysis upon binding to nicked DNA
implicates this region as a component of the DNA binding surface of the enzyme.
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EXPERIMENTAL PROCEDURES |
Enzyme Purification--
Recombinant wild type
Chlorella virus DNA ligase (12) was a gift of Dr. C. K. Ho. Mutant enzyme D29A (10) was expressed in bacteria without an
affinity tag and purified from soluble cell lysates by S-Sepharose and
blue-Sepharose column chromatography. The chromatographic behavior of
D29A was monitored by
SDS-PAGE1 analysis of the
column fractions. The lysate was applied to S-Sepharose that had been
equilibrated with 0.2 M NaCl in Buffer A (50 mM Tris-HCl (pH 7.5), 2 mM EDTA, 5 mM DTT, 10%
glycerol). The column was step-eluted with Buffer A containing 0.4, 0.6, 0.8, and 1 M NaCl. D29A was recovered in the 0.6 M NaCl step. The peak fractions were pooled, diluted to 0.2 M NaCl, and then adsorbed to a blue-Sepharose column that
had been equilibrated with 0.2 M NaCl in Buffer A. The
column was developed with a linear gradient of 0.2-1 M
NaCl. D29A was eluted at ~0.6 M NaCl. The peak fractions
were pooled and concentrated by centrifugal ultrafiltration. Protein
concentration was determined with the Bio-Rad dye reagent. The D29A
enzyme preparation was a generous gift of Dr. Aidan Doherty.
Preparation of Labeled DNA Ligands--
DNA ligands used in DNA
binding and footprinting experiments were prepared by annealing a 5'
32P-labeled oligonucleotide to unlabeled complementary
strands to form the nicked and gapped structures depicted in the
figures. End-labeling reactions and gel purification of the
32P-labeled strands were performed as described (9).
Labeled oligonucleotides were annealed to an ~4-fold molar excess of
unlabeled strands in a solution of 0.2 M NaCl by heating
the mixture for 5 min at 65 °C and then cooling slowly to
22 °C.
Preparation of 2',3'-Dideoxy and 2'-OH DNA Strands--
A
2',3'-dideoxy-terminated 19-mer strand was prepared in two stages.
First, an 18-mer strand d(ACATATCCGTGTCGCCCT) was 5' 32P-labeled and gel-purified. Then, the labeled 18-mer was
reacted for 30 min at 37 °C with 1 mM dideoxythymidine
triphosphate and calf thymus terminal transferase (57 units; Promega)
in primer extension buffer containing 0.1 M potassium
cacodylate (pH 7.2), 10 mM CoCl2, and 1 mM DTT. The 19-mer dideoxy product was separated from
residual 18-mer substrate by preparative electrophoresis through a 20%
polyacrylamide gel containing 7 M urea. The labeled strands
were localized by autoradiography, and the 19-mer was eluted from an
excised gel slice and recovered by ethanol precipitation.
A 3'-deoxy, 2'-hydroxyl-terminated 19-mer strand was prepared in
similar fashion. First, an 18-mer strand d(CATATCCGTGTCGCCCTT) was 5'
32P-labeled and gel-purified. Then, the labeled 18-mer was
reacted for 30 min at 37 °C with 1 mM 3' dATP
(cordycepin triphosphate) and calf thymus terminal transferase. The
19-mer 2'-OH product was separated from residual 18-mer substrate by
preparative electrophoresis through a 20% polyacrylamide gel and
recovered as described above.
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RESULTS |
Exonuclease III Footprint of DNA Ligase Bound at a Nick--
To
footprint the ligase-DNA interface on the 5'-phosphate side of a DNA
nick, we incubated the purified D29A enzyme with a singly nicked ligand
consisting of a 42-nucleotide 3'-OH-terminated hairpin DNA and a 5'
32P-labeled 18-mer strand annealed to the 5'-tail of the
hairpin strand (Fig. 1A). The
binding reaction mixtures contained 50 nM nicked DNA and up
to 0.8 µM ligase. At these concentrations of DNA and
protein, it is the adenylated form of Chlorella virus ligase
that binds to the nick to form a discrete complex (10). Analysis of the
binding reaction mixtures by native gel electrophoresis showed that
nearly all of the 32P-labeled nicked DNA was bound at 0.2 µM D29A, which suggests that about one-fourth of the
enzyme molecules in the preparation were adenylated (Fig.
1B). The D29A-adenylate is defective in the transfer of AMP
to the 5'-phosphate of the nick (10). Hence, the enzyme remained bound
at the nick when magnesium was included in the reaction mixtures (not
shown). Analysis by denaturing gel electrophoresis confirmed that the
32P-labeled 18-mer strand was unreactive with a molar
excess of D29A in the presence of magnesium during a 10-min incubation
at 22 °C (Fig. 1C, lane + and data not
shown).

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Fig. 1.
Exonuclease III footprint of
Chlorella virus DNA ligase bound at a nick.
A, the structure of the nicked duplex ligand used in the DNA
binding and footprinting reactions is shown. The
32P-labeled 5'-phosphate at the nick is indicated by the
asterisk. The vertical arrows indicate the
margins of the exonuclease III footprint of ligase on the 5'-phosphate
strand. B, DNA binding. Reaction mixtures (10 µl)
containing 50 mM Tris-HCl (pH 8.0), 5 mM DTT,
0.5 pmol of nicked ligand, and 0.25, 0.5, 1, or 2 pmol of D29A ligase
(proceeding left to right) were incubated for 5 min at 22 °C. A control mixture (lane ) contained no
ligase. The samples were adjusted to 8% glycerol and then analyzed by
electrophoresis through a native 8% polyacrylamide gel containing TBE
(90 mM Tris borate, 2.5 mM EDTA). An
autoradiogram of the gel is shown. C, exonuclease III
footprint. Reaction mixtures (10 µl) contained 50 mM
Tris-HCl (pH 8.0), 5 mM DTT, 0.66 mM
MgCl2, 0.5 pmol of nicked ligand, and D29A ligase as
specified. Control reactions containing 8 pmol of ligase
(lane +) or no ligase (lane ) were incubated
for 10 min at 22 °C and then quenched in 45 mM EDTA and
20% formamide. Footprinting reaction mixtures (lanes Exo
III) containing no ligase (lane ) or 1, 2, 4, or 8 pmol of ligase (proceeding left to right) were
incubated for 5 min at 22 °C and then supplemented with 10 units of
exonuclease III (New England Biolabs). After digestion for 10 min at
22 °C, the reactions were quenched in EDTA and formamide. The
reaction products were analyzed by electrophoresis through a 20%
polyacrylamide gel containing 7 M urea. An autoradiogram of
the gel is shown. A 5' 32P-labeled 12-mer oligonucleotide
(pATTCCGATAGTG) was analyzed in parallel (lane M).
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The ligase-DNA mixtures and a control sample containing DNA alone were
treated with exonuclease III. The 5'-labeled digestion products were
resolved by denaturing gel electrophoresis and detected by
autoradiography. The amount of exonuclease III included in the
reactions was sufficient to convert all of the 32P-labeled
18-mer into a rapidly migrating species of less than 5 nucleotides
(Fig. 1C). Ligase binding to the DNA resulted in the
appearance of a nuclease resistant doublet at the expense of the
complete digestion product. The upper band comigrated with a 5'
32P-labeled 12-mer pATTCCGATAGTG identical in sequence to
the first 12 nucleotides of the input 18-mer strand (lane
M). The ligase concentration dependence of exonuclease III
protection was similar to the ligase dependence of the protein-DNA
complex formation (Fig. 1 and data not shown). We conclude that the
ligase footprint on the 5'-phosphate strand extends 11-12 nucleotides
downstream of the nick.
The ligase footprint on the 3'-OH side of the nick was determined using
a different DNA ligand composed of three strands annealed to form the
structure shown in Fig. 2. This DNA was
32P-labeled at the 5'-end of the 60-mer template strand.
Incubation of the nicked DNA with a molar excess of D29A ligase did not
alter the size of the labeled strand. Exonuclease III digestion of the DNA in the absence of ligase converted all of the 60-mer into shorter
5'-labeled oligonucleotides, principally a cluster of three species
migrating at ~11-13 nucleotides and a minor product of ~23
nucleotides. Binding of D29A ligase resulted in the appearance of an
exonuclease III-resistant doublet migrating at 38-39 nucleotides (denoted by the asterisk in Fig. 2). Hence, the ligase
footprint on the 3'-OH side of the nick spans 8-9 nucleotides on the
template strand.

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Fig. 2.
Footprinting the ligase-DNA interface on the
3'-OH side of the nick. The structure of the nicked duplex ligand
used in the footprinting reactions is shown. The
32P-labeled 5'-phosphate on the 60-mer template strand is
indicated by the asterisk. The vertical arrows
indicate the margins of the exonuclease III footprint of ligase on the
60-mer strand. The 5'-phosphate at the nick is unlabeled. Footprinting
was performed as described in Fig. 1. Control reactions containing 8 pmol of ligase (lane +) or no ligase (lane )
were not subjected to nuclease treatment. Reaction mixtures containing
no ligase (lane ) or 1, 2, 4, or 8 pmol of ligase
(proceeding left to right) were digested with
exonuclease III. The reaction products were analyzed by electrophoresis
through a denaturing 12% polyacrylamide gel. An autoradiogram of the
gel is shown. A mixture of 5' 32P-labeled oligonucleotides
(41-, 30-, 18-, and 12-mer) was run in parallel in lane
M.
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Catalytic Contribution of the Motif I Aspartate to DNA
Ligation--
The D29A protein is impaired in the catalysis of
DNA-adenylate formation and, consequently, in strand joining. To gauge
the magnitude of the catalytic defect, we performed a kinetic analysis of the reaction of D29A with the nicked hairpin substrate under single-turnover conditions (i.e. at an 8:1 molar ratio of
enzyme to DNA). We found that D29A catalyzed progressive strand joining over a period of 20 h, attaining an end point with 84% of the input nicked substrate being sealed (Fig.
3). The reaction proceeded with apparent
first-order kinetics. A fit of the data to a single exponential yielded
a rate constant of 3.5 × 10
5 s
1. A
comparison of this value to the rate constant of 0.23 s
1
for single-turnover ligation by the wild type Chlorella
virus ligase (10) indicates that the conserved motif I Asp residue contributes about a 6000-fold enhancement of the reaction rate. Note
that there was no detectable accumulation of DNA-adenylate during the
D29A reaction (Fig. 3), implying that DNA-adenylate formation (step 2),
not the attack of the 3'-OH on DNA-adenylate to form the phosphodiester
(step 3), is rate-limiting for D29A.

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Fig. 3.
Kinetic analysis of strand joining by
D29A. Reaction mixtures containing (per 10 µl) 50 mM
Tris-HCl (pH 8.0), 5 mM DTT, 10 mM
MgCl2, 250 fmol of nicked hairpin DNA (Fig. 1A),
and 2 pmol of D29A ligase were incubated at 22 °C. Aliquots (10 µl) were withdrawn at the times indicated and quenched immediately
with EDTA and formamide. The reaction products were analyzed by
electrophoresis through a denaturing 20% polyacrylamide gel. An
autoradiogram of the gel is shown in the top panel. The
extent of ligation (fmol of 60-mer product formed) is plotted as a
function of reaction time in the bottom panel.
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The 3'-OH Is Not Required for the Binding of Ligase to Nicked
DNA--
Chlorella virus DNA ligase binds selectively to
DNA molecules containing a nick. The 5'-phosphate group and the
3'-nucleotide at the nick are critical for DNA recognition (10, 12).
The role of the 3'-OH moiety in DNA binding has not been evaluated. In
the experiment shown in Fig. 4, we
analyzed by native gel electrophoresis the binding of wild type ligase
to a DNA ligand containing a standard 3'-OH/5'-phosphate nick
versus ligands containing variant structures at the termini
of the discontinuous top strands. The ligase bound to nicked DNA, but
it did not form a complex with nicked DNA lacking a 5'-phosphate or
with DNA containing a 1-nucleotide gap between the 5'-phosphate and the
3'-OH groups (Fig. 4B). To test specifically the role of the
3'-OH group, we prepared a ligand containing a single
3'-dideoxynucleotide at the nick (Fig. 4A). The instructive finding was that ligase did bind to a 3'-deoxy/5'-phosphate nick (Fig.
4B). Similar experiments performed with purified recombinant vaccinia virus DNA ligase showed that it was also capable of binding to
a 3'-deoxy/5'-phosphate nick (not shown). Thus, the 3'-OH moiety is not
required for nick recognition by the two virus-encoded enzymes.

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Fig. 4.
Nick recognition by ligase requires the
5'-phosphate, but not the 3'-OH, at the nick. A, the
structures of the ligands used in the DNA binding assays are shown. The
sites of 32P-labeling are denoted by dots. The
two top strands were annealed to a continuous 60-mer
template strand. The sequences of the single-strand tails of the
template strand are not shown but are depicted, for simplicity, as
horizontal lines flanking the duplex regions. B,
reaction mixtures (10 µl) containing 50 mM Tris-HCl (pH
8.0), 5 mM DTT, 200 fmol of 32P-labeled DNA,
and 0, 100, 200, 400, or 800 fmol of ligase were incubated for 5 min at
22 °C. The mixtures were adjusted to 10% glycerol and then analyzed
by electrophoresis through a nondenaturing 12% polyacrylamide gel in
Tris borate/EDTA. An autoradiograph of the dried gel is shown.
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The 3'-OH of the Nick Is Required for DNA-Adenylate
Formation--
The second step of the ligase pathway is the reaction
of DNA-bound ligase-adenylate with the 5'-phosphate at the nick to form the nicked DNA-adenylate intermediate. DNA-adenylate is normally undetectable during reactions catalyzed by eukaryotic viral ligases (11, 12); this is because the enzyme remains bound to the nicked
DNA-adenylate after step 2 and rapidly catalyzes the attack of the
3'-OH on DNA-adenylate to form a phosphodiester. By definition, the
3'-OH is required for step 3. But does the 3'-OH participate in step 2?
We have shown here that a 3'-OH is not required for the binding of
ligase to the nicked substrate. If the 3'-OH plays no role in step 2 chemistry, we would expect to detect the accumulation of DNA-adenylate
when ligase reacts with the 3'-deoxy/5'-phosphate nicked DNA substrate.
3'-OH/5'-Phosphate-nicked DNA and 3'-deoxy/5'-phosphate-nicked DNA
substrates (both 5' 32P-labeled at the nick) were incubated
with an 8-fold molar excess of wild type Chlorella virus DNA
ligase in the presence of magnesium and ATP. Reaction with the standard
nick resulted in near quantitative joining of the 5'
32P-labeled 18-mer strand to the unlabeled 42-mer hairpin
strand to form a 60-mer ligation product (Fig.
5B). Chlorella
virus ligase formed no 37-mer ligation product with the 3'-deoxy
substrate (as expected), but we also detected no adenylate transfer to
the 5' 32P-labeled 18-mer strand. A positive control for
DNA-adenylate formation was provided by the reaction of vaccinia virus
DNA ligase with a 1-nucleotide gapped substrate. The vaccinia ligase
binds weakly to the gapped substrate, but it can catalyze step 2 under conditions of enzyme excess (11). The ligase tends to dissociate from
the gapped DNA-adenylate step 2 product without executing step 3. This
leads to the accumulation of high levels of an adenylated DNA strand,
AppDNA, and very little formation of the expected 36-mer ligation
product (Fig. 5B). Note that reaction of vaccinia ligase
with the 3'-deoxy nicked DNA substrate yielded only trace amounts of
DNA-adenylate (<1% of the input 5'-phosphate strand was adenylated)
(Fig. 5B). We surmise from this experiment that a 3'-OH
moiety is critical for DNA-adenylate formation by the two virus-encoded
DNA ligases.

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Fig. 5.
The 3'-OH is required for DNA-adenylate
formation. A, the structures of the substrates used in
the ligation assays are shown. The sites of 32P-labeling
are denoted by dots. B, ligation reaction
mixtures (10 µl) containing 50 mM Tris-HCl (pH 8.0), 5 mM DTT, 10 mM MgCl2, 1 mM ATP, 100 fmol of 3'-OH nicked DNA, 3'-deoxy nicked DNA,
or a 1-nucleotide (nt) gap DNA as specified, and 800 fmol of
wild type Chlorella virus (ChV) or vaccinia virus
(VV) ligase were incubated for 5 min at 22 °C. Ligase was
omitted from control mixtures (lanes ). The reactions were
quenched with EDTA and formamide, and the products were resolved by
denaturing polyacrylamide gel electrophoresis. An autoradiogram of the
gel is shown. The positions of the 5' 32P-labeled 18-mer
substrate oligonucleotide (pDNA) and DNA-adenylate
intermediate (AppDNA) are shown on the
right.
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A 2'-OH Is Unable to Substitute for the 3'-OH in the Second and
Third Steps of the Ligation Reaction--
We performed an additional
experiment in which Chlorella virus DNA ligase was reacted
in stoichiometric amounts with a nicked substrate consisting of a 5'
32P-labeled 18-mer (pTTCCGATAGTGACTACAG) and a
2'-OH-terminated 19-mer strand (pCATATCCGTGTCGCCCTTA-2'-OH) annealed to
a complementary 36-mer template strand. (The 2'-OH-terminated 19-mer
strand was prepared by 3' extension of an 18-mer with cordycepin
triphosphate.) The ligase catalyzed no detectable joining of the two
strands to form an expected 37-mer ligation product and there was only a trace conversion of the 5' 32P-labeled 18-mer to
DNA-adenylate (data not shown). Thus, we conclude that the essential
role of the 3'-OH at the nick in DNA adenylate formation cannot be
fulfilled by a 2'-OH moiety. Note that the presence of a 2'-OH has no
inhibitory effects on the action of Chlorella virus DNA
ligase, which is perfectly capable of binding and sealing a nicked
duplex in which the 3'-OH strand is all RNA (15).
Can a 2'-OH serve as the nucleophile in step 3 of the ligation
reaction? To answer this question, we tested the ability of Chlorella virus ligase to seal a preadenylated nicked duplex
DNA containing a 2'-OH at the nick (Fig.
6). The use of a preadenylated nicked
molecule bypasses the requirement for the 3'-OH during step 2 and
allows us to focus exclusively on the reactivity of a 2'-OH as the
nucleophile in the strand closure step. The preadenylated 2'-OH
substrate and a preadenylated 3' OH control nicked DNA substrate were
reacted with an 8-fold excess of ligase for 60 min. The 3'-OH substrate
was efficiently joined to the adenylated strand to yield the expected
ligation product, and the reaction was complete in 15 min (Fig. 6). In
contrast, the 2'-OH strand was completely unreactive with the
DNA-adenylate (Fig. 6). We conclude that Chlorella virus
ligase displays a strict requirement for a 3'-OH during the step of
phosphodiester bond formation.

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Fig. 6.
A 2'-OH cannot replace the 3'-OH in strand
closure at a preadenylated nick. The adenylated 18-mer
oligonucleotide used to form the preadenylated nicked substrate was
synthesized by ligase-mediated AMP transfer to the 5'
32P-labeled 18-mer strand of a DNA molecule containing a
1-nucleotide gap (13). The radiolabeled DNA-adenylate strand was
gel-purified and annealed to an unlabeled 36-mer template in the
presence of a 2'-OH 19-mer strand to form the 2'-OH structure shown in
the figure. The molar ratio of 18-mer DNA-adenylate to 36-mer to 19-mer
strands in the annealing reaction was 1:2:3. The 19-mer 2'-OH strand
was 5' 32P-labeled at 1/400 the specific activity of the
18-mer DNA-adenylate strand. A control 3'-OH substrate was prepared by
using an unlabeled 3'-OH 18-mer strand in place of the 19-mer 2'-OH
strand. The structure of the 3'-OH substrate is shown. Ligation
reaction mixtures containing (per 10 µl) 50 mM Tris-HCl
(pH 8.0), 5 mm DTT, 10 mM MgCl2, 80 fmol of
preadenylated nicked DNA, and 640 fmol of ligase were incubated at
22 °C. Aliquots (10 µl) were removed after 15, 30, and 60 min and
quenched immediately with EDTA and formamide. The products were
resolved by denaturing polyacrylamide gel electrophoresis. An
autoradiogram of the gel is shown. The positions of the input
32P-labeled DNA-adenylate strand (AppDNA), the
nonadenylated 5' 32P-labeled 18-mer (pDNA,
analyzed in lane M), and the 36-mer ligation product formed
with the 3'-OH substrate are indicated on the right.
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Probing the Structure of DNA Ligase by Limited Proteolysis with
Trypsin--
Purified D29A ligase was subjected to proteolysis with
increasing amounts of trypsin. SDS-PAGE analysis of the undigested enzyme preparation revealed an ~40 kDa polypeptide corresponding to
the Chlorella virus ligase (Fig.
7). The electrophoretic mobility of the
298-amino acid PBCV-1 ligase (predicted size of 34 kDa) is anomalously
slow (12). Sequencing of the 40-kDa species by automated Edman
chemistry after transfer to a polyvinylidene difluoride membrane
confirmed that the N-terminal sequence (AITKPL) corresponded to that of
the Chlorella virus enzyme beginning at residue Ala-2 of the
ligase polypeptide. Apparently, the ligase suffered removal of the
initiating methionine during expression in Escherichia coli.
Initial scission of the ligase at low concentrations of trypsin yielded
two clusters of digestion products: a set of 3 polypeptides migrating
at ~26-29 kDa and a smaller set of polypeptides migrating at
~10-12 kDa (Fig. 7). Sequencing of these cleavage products revealed
that each of the polypeptides in the larger cluster derived from the N
terminus of the enzyme. The lower molecular mass cluster was a mixture
of at least 4 polypeptides. The N-terminal sequences of the two major
species, RSTHKSGKV and STHKSGKVE, indicated that these products arose
by tryptic cleavage at Lys-219/Arg-220 and Arg-220/Ser-221. Two less
abundant peptides within the low molecular mass cluster derived from
tryptic cleavages at Lys-210/Thr-211 and Lys-224/Ser-225. The
trypsin-accessible sites fall within a 16-amino acid segment between
nucleotidyl transferase motifs V and VI (see Fig. 10). The finding that
the two clusters of tryptic products persisted at trypsin
concentrations in excess of the amount sufficient to cleave all of the
input protein suggested that these species corresponded to folded
protein domains. The sensitivity of wild type ligase to tryptic
proteolysis was identical to that observed with the D29A enzyme (not
shown). These results are consistent with a bipartite ligase structure
consisting of a larger N-terminal domain and a smaller C-terminal
domain connected by a linker segment distal to motif V.

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Fig. 7.
Limited proteolysis of Chlorella
virus DNA ligase with trypsin. Reaction mixtures (10 µl)
containing 50 mM Tris-HCl (pH 8.0), 5 mM DTT, 7 µg of D29A ligase, and 1, 2, 4, or 8 ng of trypsin (proceeding from
left to right) were incubated for 10 min at
22 °C. Trypsin was omitted from a control reaction (lane
). The reactions were quenched by the addition of SDS sample buffer,
and the digests were analyzed by electrophoresis through a 16%
polyacrylamide gel containing 0.1% SDS. A photograph of the Coomassie
Blue-stained gel is shown. The positions and sizes (kDa) of marker
proteins are indicated on the right. A parallel set of
reaction mixtures was resolved by SDS-PAGE, and the indicated
polypeptide bands were subjected to automated N-terminal sequencing as
described (17). The N-terminal sequences are denoted in
single-letter code.
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Nicked DNA Affords Protection of Ligase from Tryptic
Proteolysis--
The tryptic sensitivity of free ligase was compared
with that of ligase preincubated with an equimolar concentration (15 µM) of DNA ligand containing a single 3'-OH/5'-phosphate
nick. The trypsin concentration dependence of cleavage of the free
enzyme to yield the N-terminal cluster (Fig.
8) was similar to that seen in Fig. 7.
Nicked DNA afforded substantial (~8-fold) protection of the ligase
from tryptic digestion (Fig. 8). We did not observe protection of
ligase from trypsin digestion when the ligase was preincubated with the
42-mer hairpin strand alone (not shown). These results suggest that the
trypsin-sensitive segment of the free ligase that is protected in the
presence of DNA may comprise part of the DNA binding site on the
enzyme. Alternatively, protection from proteolysis may occur because of
a ligand-induced conformational change involving a site on the enzyme
other than the DNA-binding surface. A ligand-induced conformational
change would be supported by a finding of DNA-induced acquisition of
trypsin sensitivity at novel sites not cleaved in the free enzyme.
However, we found that the electrophoretic mobility of the digestion
products generated in the presence of nicked DNA did not differ from
those produced in the absence of DNA.

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Fig. 8.
DNA protects Chlorella virus
ligase from digestion by trypsin. Reaction mixtures (10 µl)
containing 50 mM Tris-HCl (pH 8.0), 5 mM DTT,
and 5 µg (150 pmol) of D29A ligase were preincubated for 10 min at
22 °C in the presence (lanes 6-10) or absence
(lanes 1-5) of 150 pmol of singly nicked DNA. The
structure of the DNA ligand is shown above the right
panel. The samples were then digested for 10 min at 22 °C with
1, 2, 4, or 8 ng of trypsin (proceeding from left to
right within each titration series). Trypsin was omitted
from control reactions (lanes 1 and 6). The
digestion products were analyzed by SDS-PAGE. A Coomassie Blue-stained
gel is shown. Polypeptides corresponding to the full size ligase and
N-terminal tryptic fragments are denoted on the right.
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Limited Proteolysis of DNA Ligase with Chymotrypsin--
To
further probe the structure of Chlorella virus DNA ligase
and its binding site for DNA, we carried out limited proteolysis with
chymotrypsin. Digestion of the enzyme in the absence of DNA yielded two
sets of cleavage products: a set of 3 polypeptides migrating at
~26-29 kDa and a smaller set of polypeptides migrating at ~10-12
kDa (Fig. 9). The polypeptides in the
larger cluster derived from the N terminus of the enzyme. The lower
molecular mass cluster was of a mixture of at least 5 polypeptides. The N-terminal sequences of two of these small polypeptides, KSGKV and
SKRST, suggested that initial scission by chymotrypsin occurred at
His-223/Lys-224 and Tyr-217/Ser-218. These two chymotryptic cleavage
sites fall within the same segment of the ligase between motifs V and
VI that is accessible to trypsin (Fig.
10). Other peptides within the low
molecular mass cluster of the chymotryptic digest derived from the N
terminus or from proteolytic cleavages at Tyr-89/Asn-90 and
Met-83/Thr-84 (Fig. 7). The latter chymotryptic sites are located
between motifs III and IIIa (Fig. 10).

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Fig. 9.
Limited proteolysis of Chlorella
virus DNA ligase with chymotrypsin. Reaction mixtures (10 µl) containing 50 mM Tris-HCl (pH 8.0), 5 mM
DTT, and 5 µg of D29A ligase were incubated for 10 min at 22 °C in
the presence (right panel) or absence (left
panel) of 150 pmol of nicked DNA. The mixtures were then digested
for 10 min at 22 °C with 50, 100, 200, or 400 ng of chymotrypsin
(proceeding from left to right within each
titration series). Chymotrypsin was omitted from control reactions
(lanes ). The samples were analyzed by SDS-PAGE. A
Coomassie Blue-stained gel is shown. The positions of 29- and 14-kDa
marker proteins are indicated on the left. A parallel set of
digests was resolved by SDS-PAGE, and the indicated polypeptide bands
were subjected to automated N-terminal sequencing. The N-terminal
sequences are denoted in single-letter code.
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Fig. 10.
Sites of protease cleavage within native
Chlorella virus DNA ligase. The amino acid
sequence of Chlorella virus DNA ligase is shown. Nucleotidyl
transferase motifs I, III, IIIa, IV, V, and VI are
highlighted in boxes. The sites of proteolysis by
trypsin are denoted by the arrows above the sequence, and
sites cleaved by chymotrypsin are indicated by the arrows
below the sequence.
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Prior incubation with nicked DNA protected the ligase from initial
scission by chymotrypsin and altered the distribution of cleavage
products within the high molecular mass cluster (Fig. 9). The major
species of ~29 kDa generated by chymotrypsin in the presence of DNA
comigrated with the least abundant of the 3 polypeptides in the high
molecular mass cluster seen in the absence of DNA. Sequencing of the
29-kDa polypeptide showed that it derived from the N terminus of the
ligase. This N-terminal polypeptide was relatively refractory to
further proteolysis. The lower molecular mass cluster of chymotryptic
products formed in the presence of nicked DNA consisted of the same set
of peptides detected in the digest of the free enzyme, except that the
NAKFSY product of cleavage at Tyr-89/Asn-90 was relatively more
abundant in the DNA-containing digests than in the digest of the free
ligase. Although the C termini of the chymotryptic fragments are not
known, the findings suggest that the 29-kDa N-terminal fragment is the initial chymotryptic product (a consequence of scission at
His-223/Lys-224) and that nearby secondary cleavages (e.g.
at Tyr-217/Ser-218 and at least one other site) generate the slightly
smaller products in the high molecular mass cluster. In some
chymo-tryptic digests, we detected among the low molecular mass
products a peptide with the sequence KNTNT, which indicated cleavage at
Phe-204/Lys-205. Nicked DNA apparently protects the ligase from primary
and secondary cleavages at these sites.
Effect of Limited Proteolysis on DNA Binding--
The D29A ligase
was subjected to limited proteolysis with increasing amounts of trypsin
or chymotrypsin in the absence of DNA. Aliquots of the proteolyzed
enzyme were then incubated with 32P-labeled nicked duplex
DNA, and DNA-protein complex formation was assayed by native gel
electrophoresis (Fig. 11A).
The remainder of the digest was analyzed by SDS-PAGE to ascertain the
extent of scission of the ligase polypeptide (Fig. 11B). We
envisioned three outcomes of the effect of limited proteolysis on DNA
binding: (i) if scission of the ligase between motifs V and VI did not disrupt the physical association of the two domains or the capacity of
the cleaved enzyme to recognize the nick, then the proteolyzed enzyme
should form a shifted complex on the DNA with little perturbation of
the mobility of that complex compared with undigested ligase; (ii) if
proteolytic scission dissociated the domains and either domain retained
the capacity to bind to the nick, then we expected to see the
appearance of a protein-DNA complex with novel electrophoretic mobility; (iii) if scission within the protease-sensitive segment abolished nick recognition, then all DNA binding ability should decay
in relation to the reduction in uncut ligase and no novel complex
should be evident. The results of Fig. 11 are consistent with the third
scenario. We conclude that the integrity of the protease-sensitive
linker segment is critical for nick recognition.

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Fig. 11.
Proteolysis of ligase results in loss of DNA
binding. Reaction mixtures (20 µl) containing 50 mM
Tris-HCl (pH 8.0), 5 mM DTT, 9.4 µg of D29A ligase, and
either 1, 2, 4, or 8 ng of trypsin (from left to
right) or 50, 100, or 200 ng of chymotrypsin
(Chymo, from left to right) were
incubated at 22 °C. A, after 5 min, an aliquot (1 µl)
was withdrawn from each mixture and diluted in 50 mM Tris
(pH 8.0), 10% glycerol to a final concentration of 1 µM
ligase. DNA binding reaction mixtures (10 µl) containing 50 mM Tris-HCl (pH 8.0), 5 mM DTT, 0.5 pmol of
32P-labeled nicked DNA ligand, and 1 µl of the diluted
ligase digests were incubated for 5 min at 22 °C. The binding
reaction products were analyzed by native gel electrophoresis. An
autoradiogram of the gel is shown. A control DNA binding reaction
mixture containing nicked DNA but no ligase was run in the
leftmost lane. B, the proteolysis reactions were
terminated with SDS after 10 min, and the digests were analyzed by
SDS-PAGE. A Coomassie Blue-stained gel is shown. The positions of 29- and 18-kDa marker proteins are indicated on the left.
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DISCUSSION |
The experiments presented in this study enhance our understanding
of nick recognition and catalysis by ATP-dependent DNA
ligases in four respects: (i) we provide the first physical
characterization of the enzyme binding site on a singly nicked duplex
DNA; (ii) we identify a protease-sensitive interdomain linker that may
comprise part of the substrate binding site on the ligase; (iii) we
show that the 3'-OH group at the nick is not required for nick
recognition but is essential for DNA-adenylate formation by
Chlorella virus ligase; and (iv) we find that a 2'-OH cannot
substitute for the 3'-OH during steps 2 or 3 of the ligase reaction.
DNA Footprint of Ligase Bound at a Nick--
We used exonuclease
III to footprint Chlorella virus D29A ligase bound at a
single nick in duplex DNA. The size of the binding site was 19-21
nucleotides. The ligase footprint was slightly asymmetric, extending
8-9 nucleotides on the 3'-OH side of the nick and 11-12 nucleotides
on the 5'-phosphate side. The margins of the footprint were remarkably
discrete. Also, there was nearly complete exonuclease arrest at these
margins when ligase was present at concentrations sufficient to bind
nearly all of the input ligand. We surmise that the
Chlorella virus ligase does not dissociate from the DNA or
diffuse linearly away from the nick during the time frame of the
in vitro assays. Given that Chlorella virus ligase is a monomer in solution and that ligase binding to nicked DNA
yields a single discrete complex when analyzed by native gel electrophoresis, we attribute the observed footprint to the binding of
a single enzyme molecule at the nick. The use of the D29A enzyme for
the nuclease footprint analysis was dictated by the need to prevent
ligase-adenylate from immediately catalyzing strand joining once bound
to the nicked DNA in the presence of magnesium. The footprint observed
for the D29A enzyme is likely to mimic that of the wild type
Chlorella virus ligase, insofar as: (i) the affinity of D29A
for the nick is equivalent to that of wild type ligase, (ii) the
mobility of the D29A-nicked DNA complex during native gel
electrophoresis is identical to that of the wild type ligase-nicked DNA
complex, and (iii) the tertiary structures of the D29A and wild type
ligase are grossly similar as gauged by partial proteolysis.
The 298-amino acid Chlorella virus ligase represents a
minimal catalytic domain of an ATP-dependent ligase. As
such, its footprint on nicked DNA may represent a minimal interface
required for intrinsic nick sensing. Other eukaryotic DNA ligases,
which are much greater in size (552-922 amino acids), may well cover a
larger segment of DNA than does the Chlorella virus enzyme.
To examine this point, we attempted to footprint the 552-amino acid
vaccinia virus DNA ligase bound at a nick. We engineered and purified a
mutated version of vaccinia ligase (D233A) in which the motif I Asp was
replaced by alanine with the intent of recapitulating the step-arrest
phenotype of the Chlorella virus D29A mutant. Vaccinia
ligase D233A was active in enzyme-adenylate formation, and the mutant
protein bound to nicked DNA with normal affinity. Unfortunately, a low
level of residual strand joining activity (1% of that of the wild type vaccinia ligase) was sufficient to allow single-turnover ligation within the time frame of the footprinting assays once the D233A ligase-DNA complexes were supplemented with magnesium. We conclude that
a very tight step-arrest mutation is required to obtain a satisfactory
nuclease footprint in the presence of a divalent cation cofactor.
Proteolytic Footprinting of Free and DNA-bound Ligase--
Limited
proteolysis of native and DNA-bound Chlorella virus ligase
provides a low resolution view of the enzyme's domain structure and a
clue to the location of the DNA binding site on the enzyme. The peptide
bonds in the native enzyme that were most accessible to cleavage by
trypsin and chymotrypsin cluster within a short segment located distal
to motif V. This suggests that Chlorella virus ligase
consists of an N-terminal domain that includes nucleotidyl transferase
motifs I, III, IIIa, IV, and V, plus a smaller C-terminal domain. The
domain structure inferred from the proteolysis experiments is
consistent with the actual domain structure of T7 DNA ligase determined
by x-ray crystallography (6).
It is worth noting that the principal sites of protease accessibility
in the T7 and vaccinia ligases are situated between motifs III and IIIa
(16, 17) rather than at the interdomain boundary deduced from the
crystal structure. The protease-accessible region between motifs III
and IIIa corresponds to an exposed surface loop that is disordered in
the T7 ligase crystal (6). The Chlorella virus ligase is
cleaved by chymotrypsin between motifs III and IIIa, but this site is
less accessible than the sites located distal to motif V. The segment
between motifs III and IIIa is shorter in Chlorella virus
ligase (24 amino acids) than in T7 ligase (49 amino acids) and is less
rich in potential tryptic sites than the corresponding segments of the
T7 and vaccinia enzymes. Thus, the relative protease-resistance of the
putative loop within domain 1 of Chlorella virus ligase
fortuitously permitted the delineation of a protease-sensitive
interdomain linker elsewhere in the protein. The amino acid sequence of
the linker segment of the Chlorella virus ligase
(FGYSKRSTHKSG) is loosely conserved at an analogous location distal to
motif V in the DNA ligases of Schizosaccharomyces pombe
(YGKGKRTSVYGA), Saccharomyces
cerevisiae (YGRGKRTGTYGG),
Desulfurolobus ambivalens
(HGKGRKGGKYSS), human ligase I
(LGRGKRAGRYGG), and bacteriophage
T4
(YPHRKDPTKAGG)
(18).
The interdomain linker the Chlorella virus ligase was less
accessible to proteolysis when the enzyme was bound to nicked duplex DNA. The simple interpretation of this result is that the linker comprises part of the DNA binding site and protection from proteolysis reflects steric hindrance by the bound substrate. The proposition that
the interdomain segment might be directly involved in DNA binding is
consonant with the suggestion that duplex DNA might bind within a deep,
positively charged cleft separating domains 1 and 2 of the T7 ligase
(6). Because there is no obvious sequence conservation between the
Chlorella virus ligase linker and domain 2 of T7 ligase, we
are unable to model it on the T7 structure. We do not exclude an
alternative interpretation of the data whereby DNA binding to ligase
elicits a conformational change in the linker segment. Conformational
flexibility of a related nucleotidyl transferase is revealed in the
crystal structure of Chlorella virus RNA capping enzyme,
which can adopt an open conformation with a wide interdomain groove and
a closed form with a narrow groove (19). The conformational transition
in the capping enzyme entails local changes in the secondary structure
of motif V and distal elements within domain 2.
Catalytic Role of the 3'-OH of the Nick--
The 5'-phosphate and
3'-OH groups at the nick act sequentially as nucleophiles during steps
2 and 3 of the strand joining pathway. The 5'-phosphate is also
essential for the binding of ligase-AMP to nicked DNA, implying that
functional groups on the enzyme make direct contact with the
5'-phosphate prior to step 2 chemistry. Here, we have shown that the
3'-OH moiety is not required for nick recognition. The
Chlorella virus ligase and the vaccinia enzyme bind to a
nicked ligand containing 3'-deoxy and 5'-phosphate termini. An
instructive finding was that such nicks are not adenylated by the viral
ligases, i.e. that the 3'-OH is important for step 2 chemistry even though it is not itself chemically transformed during
DNA-adenylate formation.
Our results confirm and extend the early findings of Modrich and Lehman
(20) in their classic paper describing the steady state kinetic
analysis of the composite ligation reaction and the partial reactions
catalyzed by the NAD-dependent E. coli DNA ligase. Using homopolymeric substrates consisting of reactive poly(dT)
strands annealed to poly(dA), they were able to detect the addition of
AMP to the 5'-phosphate ends of poly(dT) that was modified by the
addition of a single 3'-dideoxythymidylate residue. However, the rate
of synthesis of DNA-adenylate on the homopolymeric 3'-deoxy substrate
was at least three orders of magnitude slower than the rate of strand
joining with a 3'-OH substrate. Modrich and Lehman (20) proposed, as we
do, that the 3'-OH is essential for step 2. A more recent study by Yang and Chan (21) demonstrated that 3'-dideoxynucleotide substitution at
the nick completely blocked DNA-adenylate formation and strand joining
by human DNA ligases I and II, but the issue of whether the human
ligases could bind to the 3'-deoxy nicks was not addressed. The present
study of Chlorella virus ligase clearly excludes a requirement for the 3'-OH in nick recognition as an explanation for the
step 2 block. We suggest that the 3'-OH at the nick interacts with an
essential step 2 catalyst, either an amino acid on the Chlorella virus enzyme or the divalent cation cofactor. The
key role of the 3'-OH during step 2 of the ligation reaction of
Chlorella virus DNA ligase cannot be fulfilled by a 2'-OH group.
A need for the 3'-OH during step 2 may not apply to every DNA ligase.
Tomkinson et al. (22) have shown that the S. cerevisiae DNA ligase Cdc9p is able to catalyze DNA-adenylate
formation at a 3'-dideoxy/5'-phosphate nick. This is remarkable, given
that yeast Cdc9p is regarded as the structural and functional
counterpart of mammalian DNA ligase I (22), which as noted above,
stringently requires the 3'-OH for catalysis of DNA-adenylate formation.
The next milestone in understanding ligase action in molecular detail
will be the determination of the structure of the enzyme bound to DNA.
The present findings inform such efforts, insofar as (i) knowledge of
the ligase footprint can direct the design of nicked substrates for
cocrystallization trials and (ii) stable nick binding by step-arrest
mutants makes such mutants good candidates for crystallization.