From the Department of Chemistry, University of North
Carolina, Chapel Hill, North Carolina 27599, the
§ Biostructures Group, deCODE Genetics, Inc., Bainbridge
Island, Washington 98110, the ¶ Laboratory of Molecular
Pharmacology, NCI, National Institutes of Health, Bethesda, Maryland
20892, and the
Department of Biochemistry & Biophysics, and
the Lineberger Comprehensive Cancer Center, University of North
Carolina, Chapel Hill, North Carolina 27599
Received for publication, December 18, 2002, and in revised form, January 8, 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1- Human topoisomerase I solves the DNA topological problems that
arise from a wide variety of nuclear processes including replication, transcription, and recombination (1, 2). The enzyme nicks one strand of
duplex DNA using a transesterification reaction that produces a
transient 3'-phosphotyrosine linkage and guides the relaxation of
either positive or negative superhelical tension by a proposed
"controlled rotation" mechanism (3). The enzyme then catalyzes a
second transesterification in which the free hydroxyl at the 5'-end of
the nicked DNA strand attacks the phosphotyrosine bond, resealing the
nick, and releasing a more relaxed DNA molecule. Topoisomerase I plays
a vital role in maintaining DNA stability and is known to travel with
active replication and transcription complexes in human cells (4,
5).
Human topoisomerase I is the sole target of the camptothecins
(CPT), a potent class of anticancer drugs used to treat late-term solid
malignancies (3, 4, 6). Camptothecin effectively targets the religation
phase of topoisomerase I catalysis by stabilizing the covalent
protein-DNA complex and trapping the enzyme on DNA (7, 8). In this way,
CPT converts topoisomerase I into a cellular poison. Human
topoisomerase I is also affected by several forms of DNA damage,
including abasic lesions, wobble base pairs, and base pair mismatches
(9-13). Such lesions can impact each stage of topoisomerase I's
catalytic cycle, including DNA binding, single-strand DNA cleavage, and religation.
1--D-Arabinofuranosylcytosine
(Ara-C) is a potent antineoplastic drug used in the treatment of acute
leukemia. Previous biochemical studies indicated the
incorporation of Ara-C into DNA reduced the catalytic activity of human
topoisomerase I by decreasing the rate of single DNA strand religation
by the enzyme by 2-3-fold. We present the 3.1 Å crystal structure of
human topoisomerase I in covalent complex with an oligonucleotide
containing Ara-C at the +1 position of the non-scissile DNA strand. The
structure reveals that a hydrogen bond formed between the 2'-hydroxyl
of Ara-C and the O4' of the adjacent
1 base 5' to the damage site stabilizes a C3'-endo pucker in the Ara-C arabinose ring. The structural distortions at the site of damage are translated across the
DNA double helix to the active site of human topoisomerase I. The free
sulfhydryl at the 5'-end of the nicked DNA strand in this trapped
covalent complex is shifted out of alignment with the
3'-phosphotyrosine linkage at the catalytic tyrosine 723 residue, producing a geometry not optimal for religation. The subtle
structural changes caused by the presence of Ara-C in the DNA duplex
may contribute to the cytotoxicity of this leukemia drug by prolonging the lifetime of the covalent human topoisomerase I-DNA complex.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
Discussion
REFERENCES
-D-Arabinofuranosylcytosine
(Ara-C)1 is a nucleoside
analogue used in the treatment of acute leukemia (14, 15). Ara-C and
the standard cytosine DNA base differ by the presence of a 2'-hydroxyl
on the arabinose ring of the drug (Fig.
1). Ara-C is thought to inhibit DNA
polymerases central to replication and repair processes, and thus to
slow the growth of malignant cells (16, 17). The detailed impact of
Ara-C on human cells, however, is poorly understood. Incorporation of
Ara-C into DNA causes localized alterations in the DNA duplex,
including changes in sugar pucker, base stacking, and
backbone torsion angles (17, 18). Biochemical studies using human
topoisomerase I have revealed that Ara-C incorporation at the +1
position of the intact (non-scissile) strand adjacent to the site of
single-strand DNA cleavage induces a 4-6-fold increase in covalent
topoisomerase I-DNA complexes caused by a 2-3-fold decrease in the
rate of religation by the enzyme (12). Because the stabilization of
covalent topoisomerase I-DNA complexes converts the enzyme into a
cellular poison, the cytotoxicity of Ara-C may be enhanced by this
ability to impact the action of topoisomerase I.
View larger version (17K):
[in a new window]
Fig. 1.
Chemical structures of Ara-C and
ribocytosine nucleosides, with the O2, O3', and O5' oxygens labeled in
each.
Human topoisomerase I is 765 amino acids (91 kDa) and is composed of four domains: N-terminal (residues 1-200), core (201-635), linker (636-712), and C-terminal domain (713-765). Several crystal structures of human topoisomerase I DNA complexes have been determined (3, 8, 13, 19-21). Core subdomains I and II form the "CAP" of human topoisomerase I that contacts one side of the DNA, while core subdomain III, the "CAT," and the C-terminal domain contact the opposite side of the DNA. The CAP and CAT regions of the enzyme together wrap completely around the DNA duplex and position the active site residues within hydrogen bonding distance of the scissile DNA phosphate group. Four of five active site residues, Arg-488, Lys-532, Arg-590, and His-632, are located in core subdomain III, while the catalytic Tyr-723 resides in the C-terminal domain (3, 19, 20).
Structures of covalent human topoisomerase I-DNA complexes containing an intact 3'-phosphotyrosine linkage have also been reported (8, 19). In these trapped covalent protein-DNA complexes, the scissile phosphate contained a bridging phosphorothiolate linkage, which, upon cleavage by topoisomerase I, generates a free 5'-sulfhydryl unable to participate in strand religation (8, 19, 22). The use of 5'-bridging phosphorothiolate linkages to trap covalent complexes has been successfully employed to examine several enzymes that form transient 3'-phosphotyrosine linkages, including eukaryotic type IB topoisomerases, viral topoisomerases, and bacterial and phage tyrosine recombinases and integrases (8, 19, 22-28). Detailed biochemical studies have shown that the presence of a 5'-bridging phosphorothiolate linkage has a marginal effect on the rate of cleavage by such enzymes (down ~2-fold), but lowers the rate of religation by at least 10,000-fold (24). In addition, x-ray crystallographic studies have revealed that when the active form of human topoisomerase I (with the Tyr-723 residue intact) is used for crystallization, a bridging phosphorothiolate linkage is required to obtain crystals (8, 19).
We determined the 3.1 Å crystal structure of a human
topoisomerase I in covalent complex with a 22-base pair
oligonucleotide containing Ara-C at the +1 position of the non-scissile
DNA strand to elucidate the structural impact of Ara-C on this enzyme.
This is only the third structure of a covalent topoisomerase I-DNA complex reported to date (8, 19). We find that Ara-C introduces numerous subtle structural changes, including changes in sugar pucker
and base position, that contribute to a new positioning of the free
5'-sulfhydryl away from the 3'-phosphotyrosine linkage. Thus, the
single-strand religation reaction catalyzed by the enzyme is decreased,
producing a longer lived covalent protein-DNA complex.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Protein Purification and DNA Oligonucleotides-- The core domain of human topoisomerase I (58 kDa, residues 175-659 including a nuclear localization signal) was expressed using baculovirus in Spodoptera frugiperda (Sf9) insect cells as described (29). The 6.3-kDa C-terminal domain of human topoisomerase I (residues 713-765) was expressed as a GST fusion protein in Escherichia coli and immobilized on glutathione S-Sepharose beads (Amersham Biosciences). Nuclear extracts of Sf9 cells expressing the 58 kDa core domain were mixed in batch with the glutathione beads containing immobilized C-terminal domain, producing a reconstituted topoisomerase I complex. This complex was then purified to homogeneity as described (29).
HPLC-purified DNA oligonucleotides were purchased from Oligos Etc
(Midlands, TX) and annealed as described. The following 22-bp DNA
duplex was created,
|
where corresponds to the site of cleavage and the bold,
underlined C indicates the position of the Ara-C base. The G base in
italics contained a bridging phosphorothiolate linkage, which upon
cleavage by human topoisomerase I generates a free 5'-sulfhydryl rather
than a hydroxyl (22-28). This method of trapping human topoisomerase I
in covalent complexes with duplex DNA has been used previously in
structural studies of this enzyme (8, 19), as well as in several
detailed mechanistic and biological studies of topoisomerases and
related tyrosine recombinases (22-28). Note that the Ara-C nucleotide
is in the intact DNA strand, and the G nucleotide containing the
5'-bridging phosphorothiolate linkage is in the scissile DNA strand.
Crystallization and Data Collection-- Crystals of reconstituted human topoisomerase I in covalent complex with the 22-bp DNA duplex containing an Ara-C site of damage at the intact +1 position were grown by sitting drop vapor diffusion and cryoprotected as described (19). Crystals belong to space group P32. Data were collected at Brookhaven National Laboratory, beamline X12B, using x-ray radiation of 1.1 Å wavelength, and a MAR CCD detector. Data were processed, scaled, and reduced using HKL2000 (30).
Structure Determination and Refinement--
The structure was
determined by molecular replacement with AMoRe (31) using the crystal
structure of the human topoisomerase I reconstituted covalent complex
as a search model (PDB code 1A31; Ref. 19). 10% of the diffraction
data were set aside for the Rfree statistic
prior to any structural refinement. Partial hemihedral twinning was
detected, with a twinning fraction of 8.8% and a twinning operation of
(h, h
k,
l). The data were detwinned readily using the method of
Yeates (32) as implemented in CNS. The model was refined by rigid body,
positional, thermal displacement parameter, and simulated annealing
methods in CNS using torsion angle dynamics and the maximum likelihood
function target. Model rebuilding and manual adjustments into the
electron density maps were performed using the program O (33). Final
refinement included an overall B-factor, bulk solvent correction, and
33 well-ordered water molecules. The final structure yielded no
Ramachandran outliers as calculated by PROCHECK (34). Molecular
graphics figures were created with Molscript, Raster3D, DINO, and
Povray (35-37).2 Coordinates
have been deposited with the RCSB and assigned code 1NH3.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Overall Structure--
The crystal structure of reconstituted
human topoisomerase I in covalent complex with a 22-base pair
oligonucleotide containing Ara-C at the +1 position of the non-scissile
strand was determined to 3.1 Å resolution and refined to
crystallographic R and Rfree values
of 0.24 and 0.31, respectively (Table I).
Human topoisomerase I-DNA complex crystals are notoriously difficult to
handle (3, 13, 19-21). Indeed, in this case crystal fragility allowed
the collection of diffraction data with only 70% completeness overall to 3.1 Å resolution. In addition, because the trigonal space group P32 supports the possibility of hemihedral twinning, we
examined our data and found that twinning was present. A twinning
fraction of 0.088 and twinning operation of (h, h
k,
l) were
identified and used in refinement as implemented in CNS. Despite these
challenges, the final refined model includes all 22 base pairs of the
DNA duplex, amino acid residues 203-626 of the core domain of human topoisomerase I, and residues 719-765 of the C-terminal domain.
|
The overall structure of this complex (the Ara-C structure) is similar
to the previously reported reconstituted human topoisomerase I
enzyme in covalent complex with a non-damaged DNA duplex (PDB code
1A31; Ref. 19; Fig. 2). The quality of
the electron density is good given the limited resolution and
completeness of the diffraction data (Fig.
3). The CAP region of the enzyme's core
domain (residues 203-432) sits above the DNA and projects two
-helices out along the DNA helical axis, while the CAT (residues 433-635, 719-765) sits below the DNA and assembles the active site
around the scissile DNA phosphate. Together, the CAP and CAT wrap
completely around the DNA duplex. The C-terminal domain is packed
between the CAT and the DNA and contains the fifth active site residue,
the nucleophilic Tyr-723. The linker domain (residues 636-712) has
been observed in previous crystal structures to extend from the CAT and
C-terminal domains in the direction of the downstream region of the DNA
(3, 8, 20). This domain is not present in the reconstituted protein
construct used for these studies; the reconstituted complex is composed
of the human topoisomerase I's core domain (residues 175-659) and
C-terminal domain (713-765). The root mean square deviation (r.m.s.d.)
between the Ara-C structure and 1A31 is 0.89 Å over all protein atoms
(Table II). Shifts in position of up to
2.6 Å between individual protein residues (e.g. Ala-326 in
the CAP region) were observed when the Ara-C and 1A31 structures are
compared. Such shifts are within the range observed when the protein
regions of other human topoisomerase I DNA complexes are compared
(13, 20, 21).
|
|
|
The DNA duplex in the Ara-C structure is similar in overall structure to that observed in 1A31 (Fig. 2). In addition, like all previously described topoisomerase I-DNA complexes, the DNA molecules pack head-to-tail in a pseudocontinuous helix. The r.m.s.d. over all nucleic acid atoms between the two structures is 1.2 Å (Table II). The ends of the DNA duplex shift in position by up to 5.4 Å (Fig. 2); similar shifts in position of the ends of the DNA duplexes have been observed in previous human topoisomerase I structures (13, 20, 21).
Impact of Ara-C on DNA Base Pairing--
An Ara-C:G base pair at
the +1 intact: scissile position in the Ara-C structure replaces the
A:T base pair at this position in 1A31, a previous covalent human
topoisomerase I DNA complex (19). The presence of the Ara-C base causes
subtle changes in the DNA duplex that are localized largely at the +1
base pair (Fig. 4A). The Ara-C
and G bases shift by 1.4 and 0.75 Å, respectively, from the positions
of the A and T bases in the 1A31 structure. The arabinose ring of the
Ara-C nucleotide and the ribose ring of the G nucleotide shift in
position by up to 3.7 and 1.7 Å, respectively, relative to the
positions of the ribose rings in 1A31. The DNA bases surrounding the +1
base pair remain relatively fixed in position, however, particularly in
the scissile strand and the upstream region of the intact strand (Fig.
4A).
|
The 2'-hydroxyl of the Ara-C arabinose sugar ring extends in the
opposite direction from the equivalent group in a standard ribose sugar
(Fig. 1). The Ara-C arabinose ring exhibits a C3'-endo pucker, as
predicted based on thermodynamic/energetic considerations (Ref. 39;
Fig. 4A). The deoxyribose ring of the +1 adenine base in the
intact strand of 1A31, in contrast, exhibits the C2'-endo pucker
typical of B-form DNA bases. The C3'-endo pucker in the Ara-C arabinose
ring is stabilized by a 2.5 Å intrastrand hydrogen bond between the
Ara-C 2'-hydroxyl and the O4' of the 1 adenine (Fig. 4B).
Thus, the Ara-C base exhibits a distinct sugar pucker that alters the
+1 base pair of the DNA duplex in the Ara-C structure. This change
impacts the position of the 5'-end of the cleaved DNA strand (see
below), which helps to explain the decrease in the rate of religation
by human topoisomerase I observed with this form of DNA lesion
(12).
Two structures of DNA duplexes containing an Ara-C lesion have been
reported, one solution structure by nuclear magnetic resonance spectroscopy (NMR) and one crystal structure (17, 18). While the atomic
coordinates of the crystal structure are not available, we compared the
DNA duplex from our Ara-C complex to the 12-base pair DNA duplex
containing Ara-C determined by NMR (Fig.
5). The r.m.s.d. over all 12 equivalent
base pairs between these two duplexes is 2.3 Å. The DNA bases
superimpose well, while the backbone is more deviant between the two
structures. The ribose at the 2 scissile position in the Ara-C
complex shifts in position by 0.7 Å from the equivalent ribose in the
NMR structure; this base is at the terminus of the NMR duplex, however.
The ribose ring of the +1 scissile strand in the Ara-C structure shifts
by 2.6 Å relative to the equivalent ribose in the NMR structure, a
change likely caused by the break in the DNA backbone at this position. The arabinose sugar of the Ara-C base shifts in position by 0.9 Å.
While both rings exhibit a C3'-endo pucker, it is more extreme in the
topoisomerase I complex. Intrastrand hydrogen bonds between the Ara-C
2'-OH and the ribose O4' of the adjacent DNA base are also formed in
both structures. Thus, it appears that the nick in the DNA backbone of
the Ara-C topoisomerase I covalent DNA complex stabilizes the C3'-endo
pucker in the Ara-C arabinose sugar ring.
|
Active Site--
The active site in the Ara-C structure is similar
overall to those observed in human topoisomerase I-DNA crystal
structures elucidated previously (3, 13, 19-21). Arg-488, Lys-532,
Arg-590, and Tyr-723 share an overall r.m.s.d. of 0.6 Å with the
equivalent residues in 1A31 (Fig. 6).
His-632 is not ordered in this structure, similar to the structure of a
reconstituted covalent human topoisomerase I complex reported
previously (19). The free 5'-sulfhydryl group exhibits the largest
structural change at the active site, shifting in position by 2.6 Å relative to the 1A31 structure. Thus, in the Ara-C complex, the
5'-sulfhydryl is 3.7 Å from the 3'-phosphotyrosine linkage, while in
the 1A31 structure, the equivalent group is only 2.5 Å from the
3'-phosphotyrosine linkage. Indeed, a rotation of the 5'-sulfhydryl by
120 degrees in 1A31 places it in an ideal position to perform an
in-line attack on the covalent phosphate-tyrosine bond. In the Ara-C
complex, a similar alignment cannot be formed without more significant
structural changes. This observation helps to explain the decreased
rate of religation by human topoisomerase I using DNA substrates
containing an Ara-C lesion at the intact +1 position (12).
|
The shift in the position of the free 5'-end of the nicked DNA strand
in the Ara-C complex appears to be caused by the subtle alteration in
the +1 DNA base mediated by the arabinose Ara-C sugar. The new position
of the free 5'-sulfhydryl in the Ara-C complex brings it within 3.5 Å of the side chain of Asn-722, the amino acid immediately N-terminal in
sequence to the catalytic Tyr-723. Asn-722 shifts in position by 0.5 Å relative to the 1A31 structure; indeed, in the 1A31 complex, Asn-722 is
4.5 Å from the free 5'-sulfhydryl (19). The proximity of the Asn-722
side chain to the free 5'-end of the nicked DNA strand in the Ara-C complex indicates that a hydrogen bond is possible between these two
groups. Asn-722 is conserved in all known sequences of eukaryotic topoisomerases I and has been implicated in the catalytic action of the
enzyme. For example, mutations of this residue in both human and
Saccharomyces cerevisiae topoisomerase I impact the enzyme's sensitivity to CPT and phases of its catalytic cycle (40).
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The leukemia drug Ara-C contains a arabinose sugar ring rather
than the ribose standard to DNA and RNA bases. As such, its 2'-hydroxyl
group is oriented in a manner distinct from the equivalent RNA cytosine
base (Fig. 1). Ara-C is thought to elicit its antineoplastic effects by
acting as a competitive inhibitor of DNA polymerases and
(12,
17, 41). Even at low concentrations, however, the drug becomes
incorporated into DNA and disrupts DNA metabolism (12, 17, 41).
Pourquier et al. (12) have shown that the presence of an
Ara-C base at the +1 position of the intact strand (opposite the site
of single-strand cleavage) slows the rate of DNA strand religation by
human topoisomerase I 2-3-fold (12). The extended lifetime of the
covalent topoisomerase I-DNA complex may contribute to antineoplastic
effects of Ara-C by enhancing chromosomal instability. Indeed, human
leukemia cells that lack detectable levels of topoisomerase I are
resistant to the effects of Ara-C (12).
We determined the 3.1 Å resolution crystal structure of human
topoisomerase I in covalent complex with a 22-base pair DNA duplex
containing Ara-C at the +1 position of the intact strand (Fig. 2). The
structure reveals that the Ara-C non-standard 2'-hydroxyl introduces
numerous subtle structural changes, particularly the +1 base pair (Fig.
4A). The 2'-hydroxyl of Ara-C forms a hydrogen bond with the
O4' of the 1 sugar, which stabilizes the C3'-endo pucker exhibited by
the arabinose ring of Ara-C (Fig. 4B). These structural
changes cause the +1 base pair of the duplex to shift in position
relative to the equivalent base pair in a covalent topoisomerase I DNA
complex without a site of damage reported previously (1A31; Ref. 19).
This, in turn, appears to cause the free 5'-sulfhydryl (which replaces
the 5'-hydroxyl in this trapped covalent complex; 8, 19, 23-28) in the
nicked DNA strand to shift away from the covalent phosphotyrosine
linkage and form a hydrogen bond with the side chain of Asn-722, an
interaction not observed in previous topoisomerase I covalent complexes
(Figs. 4A and 6). Taken together, these results indicate
that the subtle change of the duplex opposite the single-strand DNA
break shifts the free 5'-end of the nicked strand away from the
covalent 3'-phosphotyrosine linkage. These results likely
explain the impact on topoisomerase I activity reported by
Pourquier et al. (12).
This Ara-C structure provides additional insight into the catalytic mechanism of human topoisomerase I. As the active site residues are brought into place upon DNA binding, Asn-722 does not appear to contact the DNA, as observed in several non-covalent topoisomerase I DNA complexes (3, 13, 20, 21). However, as the downstream region of DNA undergoes relaxation by the proposed controlled rotation mechanism, Asn-722 may have ample opportunity to hydrogen bond with the free 5'-hydroxyl of the nicked strand. Indeed, after relaxation slows, Asn-722 may play a crucial role via hydrogen bonding in guiding the 5'-hydroxyl into place for the religation phase of catalysis. This interaction is likely to be transitory in reactions involving non-damaged DNA. The change caused by the Ara-C base appears to stabilize this interaction, allowing us to visualize it in the structure presented here.
The importance of Asn-722 in human topoisomerase I and the
equivalent Asn-726 in S. cerevisiae topoisomerase I in the
catalytic cycle and camptothecin sensitivity of the enzyme have been
established by several careful biochemical studies. For example,
mutation of Asn-722 to histidine in human topoisomerase I increases the rate of DNA cleavage, while mutation to aspartic acid decreases the DNA
binding affinity of the enzyme (38). An N722S mutation in human
topoisomerase I, in contrast, does not impact the catalytic activity of the enzyme but does reduce its sensitivity to camptothecin (40). We provide structural evidence in this and previous work that
sites of DNA damage impact the ability of Asn-722 to align the active
site of human topoisomerase I both before and after single-strand DNA
cleavage by the enzyme (13). This residue may play a similar role with
other DNA lesions that impact human topoisomerase I, including
ethenoadenine adducts, wobble base pairs, and uracil mismatches. In
summary, we show that relatively subtle modifications caused by the
presence of a single 2'-hydroxyl group on the opposite side of the
substrate DNA duplex can alter the structure of the human topoisomerase
I active site and impact the catalytic action of the enzyme.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank S. Bencharit, R. Watkins, and Y. Xue for thoughtful discussions and assistance in creating figures.
![]() |
FOOTNOTES |
---|
* 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.
The atomic coordinates and the structure factors (code 1NH3) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
** To whom correspondence should be addressed: Dept. of Chemistry Campus Box 3290, University of North Carolina, Chapel Hill, NC 27599-3290. Tel.: 919-843-8910; Fax: 919-966-3675; E-mail: redinbo@unc.edu.
Published, JBC Papers in Press, January 17, 2003, DOI 10.1074/jbc.M212930200
2 DINO: Visualizing Structural Biology (2002) www.dino3d.org.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
Ara-C, -D-arabinofuranosylcytosine;
r.m.s.d., root mean square
deviation;
PDB, protein data bank.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Champoux, J. J. (2001) Annu. Rev. Biochem. 70, 369-413[CrossRef][Medline] [Order article via Infotrieve] |
2. | Wang, J. C. (1996) Annu. Rev. Biochem. 65, 635-692[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Stewart, L.,
Redinbo, M. R.,
Qiu, X.,
Hol, W. G.,
and Champoux, J. J.
(1998)
Science
279,
1534-1541 |
4. | Pommier, Y., Pourquier, P., Fan, Y., and Strumberg, D. (1998) Biochim. Biophys. Acta 1400, 83-105[Medline] [Order article via Infotrieve] |
5. | Holden, J. A. (2001) Curr. Med. Chem. Anti-Cancer Agents 1, 1-25 |
6. |
Hsiang, Y. H.,
Hertzberg, R.,
Hecht, S.,
and Liu, L. F.
(1985)
J. Biol. Chem.
260,
14873-14878 |
7. | Hertzberg, R. P., Caranfa, M. J., and Hecht, S. M. (1989) Biochemistry 28, 4629-4638[Medline] [Order article via Infotrieve] |
8. |
Staker, B. L.,
Hjerrild, K.,
Feese, M. D.,
Behnke, C. A.,
Burgin, A. B.,
and Stewart, L.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
15387-15392 |
9. |
Pourquier, P.,
Ueng, L. M.,
Kohlhagen, G.,
Mazumder, A.,
Gupta, M.,
Kohn, K. W.,
and Pommier, Y.
(1997)
J. Biol. Chem.
272,
7792-7796 |
10. |
Pourquier, P.,
Bjornsti, M. A.,
and Pommier, Y.
(1998)
J. Biol. Chem.
273,
27245-27249 |
11. |
Pourquier, P.,
Ueng, L. M.,
Fertala, J.,
Wang, D.,
Park, H. J.,
Essigmann, J. M.,
Bjornsti, M. A.,
and Pommier, Y.
(1999)
J. Biol. Chem.
274,
8516-8523 |
12. |
Pourquier, P.,
Takebayashi, Y.,
Urasaki, Y.,
Gioffre, C.,
Kohlhagen, G.,
and Pommier, Y.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
1885-1890 |
13. |
Lesher, D-T. T.,
Pommier, Y.,
Stewart, L.,
and Redinbo, M. R.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
12102-12107 |
14. | Grant, S. (2002) Adv. Cancer Res. 72, 197-233 |
15. | Mastrianni, D. M., Tung, N. M., and Tenen, D. G. (1992) Am. J. Med. 92, 286-295[Medline] [Order article via Infotrieve] |
16. | Collins, A. R. S. (1977) Biochim. Biophys. Acta 478, 461-473[Medline] [Order article via Infotrieve] |
17. | Schweitzer, B. I., Mikita, T., Kellogg, G. W., Gardner, K. H., and Beardsley, G. P. (1994) Biochemistry 33, 11460-11475[Medline] [Order article via Infotrieve] |
18. | Gao, Y.-G., van der Marel, G. A., van Boom, J. H., and Wang, A. H. J. (1991) Biochemistry 30, 9922-9931[Medline] [Order article via Infotrieve] |
19. |
Redinbo, M. R.,
Stewart, L.,
Kuhn, P.,
Champoux, J. J.,
and Hol, W. G.
(1998)
Science
279,
1504-1513 |
20. | Redinbo, M. R., Stewart, L., Champoux, J. J., and Hol, W. G. (1999) J. Mol. Biol. 292, 685-696[CrossRef][Medline] [Order article via Infotrieve] |
21. | Redinbo, M. R., Champoux, J. J., and Hol, W. G. (2000) Biochemistry 39, 6832-6840[CrossRef][Medline] [Order article via Infotrieve] |
22. | Burgin, A. B. (2001) Methods Mol. Biol. 95, 119-128[Medline] [Order article via Infotrieve] |
23. | Burgin, A., Huizenga, B., and Nash, H. (1995) Nucleic Acids Res. 23, 2973-2979[Abstract] |
24. | Krogh, B. O., Cheng, C., Burgin, A., and Shuman, S. (1999) Virology 264, 441-451[CrossRef][Medline] [Order article via Infotrieve] |
25. | Burgin, A., and Nash, H. (1995) Curr. Biol. 5, 1312-1321[Medline] [Order article via Infotrieve] |
26. | Hwang, Y., Park, M., Fischer, W. H., Burgin, A., and Bushman, F. (1999) Virology 262, 479-491[CrossRef][Medline] [Order article via Infotrieve] |
27. | Krogh, B. O., and Shuman, S. (2000) Mol. Cell 5, 1035-1041[Medline] [Order article via Infotrieve] |
28. |
Kazmierczak, R. A.,
Swalla, B.,
Burgin, A.,
Gumport, R. I.,
and Gardner, J. F.
(2002)
Nucleic Acids Res.
30,
5193-5204 |
29. | Stewart, L., Ireton, G. C., and Champoux, J. J. (1997) J. Mol. Biol. 269, 355-372[CrossRef][Medline] [Order article via Infotrieve] |
30. | Otwinowski, Z., and Minor, W. (1997) Methods Enzymol. 276, 307-326 |
31. | Navaza, J. (2001) Acta Crystallogr. Sect. D. Biol. Crystallogr. 57, 1367-1372[CrossRef][Medline] [Order article via Infotrieve] |
32. | Yeates, T. O. (1988) Acta Crystallogr. Sect. A 44, 142-144[Medline] [Order article via Infotrieve] |
33. | Brunger, A. T. (1993) Acta Crystallogr. Sect. D. Biol. Crystallogr. D49, 24-36[CrossRef] |
34. | Laskowski, R. A., MacArthur, M. W., Moss, D. S., and Thornton, J. M. (1993) J. Appl. Crystallogr. 26, 283-291[CrossRef] |
35. | Esnouf, R. M. (1999) Acta Crystallogr.Sect. D. Biol. Crystallogr. 55, 938-940[CrossRef][Medline] [Order article via Infotrieve] |
36. | Kraulis, P. J (1991) J. Appl. Crystallogr. 24, 946-950[CrossRef] |
37. | Merritt, E. A., and Bacon, D. J. (1997) Methods Enzymol. 277, 505-524 |
38. | Pourquier, P., and Pommier, Y. (2001) Adv. Cancer Res. 80, 189-216[Medline] [Order article via Infotrieve] |
39. | Thibaudeau, C., Plavec J., Watanabe K. A., and Chattopadhyaya, J. (1994) J. Chem. Soc. Chem. Commun. 537-540 |
40. |
Fertala, J.,
Vance, J. R.,
Pourquier, P.,
Pommier, Y.,
and Bjornsti, M. A.
(2000)
J. Biol. Chem.
275,
15246-15253 |
41. | Grant, S. (1997) Front. Biosc. 2, 242-252 |