MINIREVIEW
Replisome Architecture and
Dynamics in Escherichia coli *
Published, JBC Papers in Press, January 18, 2006, DOI 10.1074/jbc.R500028200
Mike O’Donnell1
From the Laboratory of DNA Replication, Howard Hughes Medical
Institute, The Rockefeller University, New York, New York 10021
The Clamp and Clamp Loader
The ␤ sliding clamp is a homodimer in the shape of a ring, which encircles
DNA (Fig. 1A). The ␤ clamp slides on DNA and binds the pol III core, thereby
acting as a mobile tether and converting the normally distributive pol III core
into a highly processive and rapid polymerase capable of incorporating 500 –
1,000 nucleotides per second (Fig. 1C). The crystal structure of the ␤ dimer
reveals a 6-fold pseudo symmetry that arises from a domain that is repeated
three times in the monomer giving the dimer a 6-fold appearance (6). The
eukaryotic PCNA clamp and phage T4 gp45 clamp are also six-domain rings,
but the monomeric unit contains only two domains and trimerizes to form a
six-domain ring (7).
Clamps do not self-assemble onto DNA but require a multiprotein clamp
loader, which harnesses the energy of ATP hydrolysis to open and close the
clamp around DNA (Fig. 1B). The E. coli ␥ complex clamp loader contains five
subunits that are essential for clamp loading activity, three ␥ protomers and
one copy each of ␦ and ␦⬘. The small ␹ and ␺ subunits are not required for
clamp loading, but they stabilize the clamp loader and stimulate its activity at
elevated ionic strength.
The ␥, ␦, and ␦⬘ clamp loading subunits are members of the AAA⫹ family
(ATPases associated with a variety of cellular functions) (8). The subunits
consist of three domains, and a AAA⫹ region of homology is localized to the
first two domains (9). Many AAA⫹ proteins are homohexamers arranged in a
symmetric circle (10, 11). The ␥3␦␦⬘ pentamer forms an asymmetric circle, and
there is a gap in place of the “missing sixth subunit” (Fig. 1B) (12). The C-terminal domains form an uninterrupted collar that ties the pentamer together.
The ␤ clamp docks onto the AAA⫹ domains of ␥3␦␦⬘ (Fig. 1, B and C). This
placement comes from the structure of ␦-␤1 (13). Replacing ␦ in ␥3␦␦⬘ with
␦-␤1 indicates that ␤ interacts with the other subunits, confirmed biochemically (14). When ␦ binds to ␤ it opens the clamp (15, 16). However, ATP
binding is required for the ␥ complex to adopt a conformation allowing it to
* This minireview will be reprinted in the 2006 Minireview Compendium, which will be
available in January, 2007.
1
To whom correspondence should be addressed: Laboratory of DNA Replication, Howard
Hughes Medical Institute, The Rockefeller University, 1230 York Ave., New York, NY 10021.
Tel.: 212-327-7252; Fax: 212-327-7253; E-mail: [email protected].
2
The abbreviations used are: pol, polymerase; PCNA, proliferating cell nuclear antigen;
RFC, replication factor C; ATP␥S, adenosine 5⬘-O-(3-thiotriphosphate); ssDNA, singlestranded DNA.
APRIL 21, 2006 • VOLUME 281 • NUMBER 16
interact with ␤ (17), and thus the unliganded crystal structure is in an inactive
conformation. The ␦-␤1 crystal structure reveals that the ␤ dimer is under
tension, and when the interface is distorted by ␦, the clamp springs open (13).
The AAA⫹ domains of the ␥ complex are arranged in a spiral (7, 12). Structural studies of the Saccharomyces cerevisiae RFC clamp loader-ATP␥S-PCNA
complex also show this spiral arrangement. The RFC-ATP␥S spiral has a
steeper helical pitch than ␥ complex and closely matches the pitch of B-DNA
for DNA interaction (18). DNA fits inside the clamp loader where the AAA⫹
domains form a central chamber and two ␣ helices on each subunit track the
minor groove of DNA modeled inside. Several polar and basic side chains on
and near these helices are conserved in prokaryotic and eukaryotic clamp
loaders. Mutations of these conserved residues in either ␥ or ␦⬘ significantly
reduce ability of ␥ complex to bind DNA (19).
The Open Clamp Is a Spiral Lockwasher
Fluorescent energy transfer studies in the T4 phage replication system indicate that the gp45 clamp protomers open out-of-plane with a left-handed
helical pitch (20). In the E. coli and S. cerevisiae systems, the right-handed
helical arrangement of ␥3␦␦⬘ and RFC subunits is compatible with a righthanded opening of the clamp, thereby allowing it to dock onto the other clamp
loading subunits. Indeed, molecular simulations of PCNA indicate that it
springs open out-of-plane with a strong tendency to form a right-handed helix
(21). In addition, electron microscopic reconstruction of an archael RFCPCNA-ATP␥S-DNA complex provides direct visual evidence for an open
clamp in a right-handed helix (22). DNA is guided from the central chamber of
the clamp loader into the clamp docked below (see Fig. 1C). DNA binding is
followed by hydrolysis of ATP to eject the clamp loader. Clamp loader ejection
is necessary as the pol III core binds the same face of ␤ as the clamp loader, and
only one of these complexes can interact with ␤ at a given time (23, 24).
Clamp loader structures suggest how specificity for a primed site is gained
(7, 12). Duplex DNA enters into the open clamp and the central chamber of the
clamp loader through the gap between ␦ and ␦⬘. However, the C-terminal
domains of the clamp loader form a tight collar with no interruption and thus
act as a cap that DNA cannot penetrate. To fit into the clamp loader, DNA
must make a sharp bend at the cap and exit out the gap in the side of the clamp
loader. A continuous duplex cannot bend abruptly and thus is unable to fit, but
ssDNA at a primed site provides flexibility for bending, and thus this structure
can be accommodated (e.g. Fig. 1C).
pol III Holoenzyme and Architecture of the Replisome
E. coli pol III holoenzyme is organized by the ␶ subunit (see Fig. 2A; reviewed
in Refs. 3–5). ␶ is encoded by the same gene as ␥, but ␥ lacks the C-terminal 24
kDa of ␶ due to truncation by a translational frameshift. The C-terminal region
unique to ␶, referred to as ␶c, binds to the pol III core. The pol III core consists
of three subunits, ␣ (DNA polymerase), ⑀ (3⬘-5⬘-exonuclease), and ␪. Within
the holoenzyme two copies of ␶ replace two ␥ subunits to form a ␥␶2␦␦⬘␹␺
clamp loader, which in turn binds two pol III cores for leading and lagging
strand replication (Fig. 2A). The C termini of the ␥ subunits protrude from the
collar and contain many prolines and polar residues. Hence, the ␶c region is
likely connected to the ␥ clamp loading domains by a flexible linker to form a
pol III holoenzyme containing two cores with one clamp loader suspended
from flexible linkers between them. The single clamp loader can load ␤ clamps
onto the leading and lagging strands for both DNA pol III cores (25).
The ␶c region connects pol III holoenzyme to the replicative helicase, DnaB
(26). DnaB is a circular homohexamer, similar to the homohexameric helicases
of the T4 and T7 phages (27–30). DnaB helicase encircles ssDNA and tracks
along it during unwinding action; its direction of movement corresponds to
the direction of the replisome provided DnaB encircles the lagging strand (Fig.
2B). Unwinding is achieved by steric exclusion in which one strand is excluded
from the inside channel, while the tracking strand is retained on the inside of
the ring, thus forcing the duplex apart as DnaB moves (30 –32).
It is hypothesized that pol III holoenzyme is functionally asymmetric since leading and lagging strand processes are so different, and evidence for asymmetry has
been gleaned from studies using ATP␥S (33–35). pol III holoenzyme is also structurally asymmetric, as defined by the clamp loader, which has odd numbers of
subunits (36, 37). The DnaB helicase adds asymmetry to the replisome due to its
placement on the lagging strand and has been demonstrated to impose asymmetric function on the two polymerases of pol III holoenzyme (25).
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The field of DNA replication was launched upon discovery of the first DNA
polymerase by Kornberg and Lehman (1, 2). DNA polymerase I displayed the
novel and highly exciting property of template-directed enzymatic action, and
like a split personality, DNA polymerase I could also degrade DNA, from either
direction. Escherichia coli is now known to contain five different DNA polymerases. The chromosomal replicase is DNA polymerase III, while the damage-inducible DNA polymerases II, IV, and V play roles in DNA repair. DNA
polymerase I is involved in both replication and repair and remains the most
advanced model for the structure and function of DNA polymerase action.
DNA polymerase III (pol III)2 functions in the context of a multiprotein
apparatus called DNA pol III holoenzyme (reviewed in (3–5)). pol III holoenzyme contains 10 different proteins, which assort into three major functional
units: 1) pol III core, 2) the ␤ sliding clamp, and 3) the ␥/␶ complex clamp
loader. The holoenzyme contains two copies of the pol III core, which are
connected by the attachment to one clamp loader. The holoenzyme functions
within the context of a dynamic replisome containing pol III holoenzyme, a
hexameric DnaB helicase, DnaG primase, and SSB. During replisome function,
contacts between these proteins are in a constant state of change. This review
briefly summarizes the architecture and dynamic behavior of the E. coli
replisome.
This paper is available online at www.jbc.org
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 281, NO. 16, pp. 10653–10656, April 21, 2006
© 2006 by The American Society for Biochemistry and Molecular Biology, Inc. Printed in the U.S.A.
MINIREVIEW: The E. coli Replication Machine
FIGURE 1. The ␤ clamp and ␥ complex clamp
loader. A, protomers of the E. coli ␤ dimer are colored differently, and DNA is modeled into the center. Adapted with permission from Kong et al. (6).
B, E. coli ␥3 ␦␦⬘. The C-terminal domains form the
collar, which mediates tight pentameric contacts.
The AAA⫹ region encompass the two N-terminal
domains of each subunit. DNA passes through the
gap between ␦ and ␦⬘. Adapted with permission
from Jeruzalmi et al. (13). C, DNA binds in the central chamber of the clamp loader (left). ATP hydrolysis ejects the clamp loader (middle). pol III associates with the ␤ clamp for processive synthesis
(right).
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FIGURE 2. pol III holoenzyme and replisome
architecture. A, arrangement of proteins within
pol III holoenzyme is illustrated to the left. Subunit
names and their mass are indicated in the table to
the right. B, replisome architecture; see text under
“Replisome Dynamics” for details.
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MINIREVIEW: The E. coli Replication Machine
Interestingly, DnaB can also encircle both strands of duplex DNA and tracks
along the duplex with force fueled by ATP (38). Although DnaB does not
unwind DNA in this mode, it displaces proteins and catalyzes branch migration of Holliday junctions. It is interesting to note that the eukaryotic helicase,
the MCM heterohexamer (reviewed in Refs. 39 – 41), also tracks on both
ssDNA and double-stranded DNA, like DnaB (42, 43). During bi-directional
replication the two replication forks are thought to be held together in a replication factory in prokaryotes and possibly eukaryotes as well (44, 45). The
SV40 T-antigen helicase and archael MCM form double hexamers: two hexamer rings stacked on top of one another (46, 47). Double hexamers of helicases provide a compelling view of how two forks in one factory may be
attached (45). A recent study indicates that the two replication forks within a
factory are functionally uncoupled (48).
Replisome Dynamics
DNA can be threaded through the replisome as illustrated in Fig. 2B. The
leading strand is excluded from the inside of the DnaB ring. The leading pol III
core-␤ continuously extends DNA in the direction of unwinding. Due to the
antiparallel structure of DNA, the lagging strand is chemically extended in the
opposite direction of the replication fork. Yet the lagging pol III core must
physically move with the fork due to its connection to the leading polymerase and helicase. Hence, the lagging strand must form a loop to accommodate these opposed motions, as hypothesized in the “trombone model”
of replication (49).
Discontinuous lagging strand fragments are 1–3 kb in length, and primase
initiates each fragment by forming a short RNA primer. Primase must interact
with DnaB for activity, which ensures that priming occurs at the replication
fork junction (45). Primase acts in a fully distributive fashion (50). As Okazaki
fragments are produced every few seconds, primase action is highly dynamic,
coming on and off DNA for each priming event.
pol III core held to DNA by a protein ring fits nicely for continuous replication of the leading strand. However, discontinuous synthesis on the lagging
strand requires the pol III core to hop from a finished fragment to start the next
Okazaki fragment every few seconds. How can pol III core rapidly dissociate
APRIL 21, 2006 • VOLUME 281 • NUMBER 16
from a completed fragment when it is held tight to DNA by a clamp? This
dilemma is solved by a processivity switch (51, 52). When the lagging pol III
core finishes an Okazaki fragment, it rapidly dissociates from the clamp (and
DNA). The clamp loader repeatedly loads ␤ clamps onto RNA primers, which
allows the lagging pol III core to reassociate with a new clamp for extension of
the next fragment (Fig. 3A). The need for polymerase to be processive, yet
rapidly recycle, may underlie one reason that replicative polymerases have
evolved to function with clamps.
Separation of pol III from ␤ is achieved by the ␶c portion of ␶ subunit (53). pol
III-␤ separation is regulated by DNA; primed DNA turns ␶c off and blocks it
from separating pol III from ␤. But when synthesis is complete, ␶c separates pol
III from ␤, leaving the clamp on DNA (53). This stoichiometric use of one ␤
clamp for each Okazaki fragment results in a build-up of used clamps on the
lagging strand, consistent with the high copy number of ␤ (⬃300/cell) relative
to pol III (10 –20/cell) (25). The ␤ clamp also interacts with pol I and ligase (54),
which are needed to replace the RNA with DNA and seal the finished fragments. Hence, leftover ␤ clamps may mark spots on DNA where these
enzymes are needed.
Okazaki fragments outnumber ␤ by about 10-fold, and therefore ␤ clamps
must be recycled. The high stability of ␤ on DNA (t1⁄2 ⬃1 h at 37 °C) implies that
clamps are actively removed from DNA. The ␥ complex clamp loader is capable of unloading clamps, but the more likely clamp unloader is ␦, which is in
excess over other clamp loading subunits in the cell (55). ␦ cannot load ␤ onto
DNA but can open ␤ and is highly active in unloading ␤ from DNA.
Physically Coupled but Mechanically Uncoupled Polymerases
Do the two DNA polymerases “communicate” their replication status to one
another? For example, if the lagging polymerase becomes blocked (i.e. by a
lesion), does the leading polymerase also stop? Studies in the T4 and T7 phage
replication systems, which also utilize twin DNA polymerases, demonstrate
strict polymerase coupling (56, 57). Shut-down of one DNA polymerase results
in cessation of the other. However, in the E. coli system leading strand synthesis continues unabated upon blocking the lagging polymerase (58, 59). In fact,
continued fork progression frees the lagging polymerase from the stall site and
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FIGURE 3. Protein trafficking on sliding clamps.
A, lagging pol III core hops among ␤ clamps. Left
diagram, lagging pol III core is extending an Okazaki fragment. Right diagram, the lagging pol III
core finishes the fragment and releases from ␤.
Also, the ␥/␶ clamp loader places ␤ on a new
primed site. Bottom diagram, lagging pol III core
binds a new clamp for the next fragment. B, pol IV
and pol III bind the same ␤ toolbelt (left). pol III
stalls at a lesion (middle), allowing pol IV to take
control of the primed site (right). pol III regains the
DNA when moving conditions are reestablished.
MINIREVIEW: The E. coli Replication Machine
allows replication of the lagging strand to continue. Hence, the two cores are
functionally uncoupled. Blocks on the leading strand also lead to uncoupling,
but the replication fork halts its advance within 1 kb (59).
Why do the phage replicases strictly couple synthesis of the two strands,
while the E. coli replicase does not? One may speculate that rapid duplication
of the cellular chromosome is too important to wait for repair of damaged
nucleotides on the lagging strand; these can be left behind and repaired later.
Due to the smaller size of phage genomes, blocking lesions may only be
encountered in a fraction of replicating molecules, allowing chromosomes
containing stalled forks to simply be discarded. In this scenario, tight polymerase coupling may act as a fidelity mechanism rather than a strategy for efficient replication.
Protein Trafficking on Clamps
Concluding Remarks
Structural and biochemical studies provide detailed insight into the dynamic
workings of the E. coli replisome. Typical of scientific inquiry, these advances
leave us with yet more questions. For example, how do moving replication
forks handle the myriad array of lesions and DNA bound proteins encountered
during synthesis? How do the many proteins that bind ␤ coordinate their
action on the clamp? What role does DnaB play while it encircles doublestrand DNA? How are the forks in a replication factory held together, and what
other proteins are present? How do replication enzymes meld their action with
repair and recombination processes, such as must occur during replication
restart after DNA damage? How do replication fork proteins coordinate with
topoisomerases during synthesis and upon termination of the chromosome?
These are only some of the many exciting questions that remain for the future.
REFERENCES
1. Lehman, I. R., Bessman, M. J., Simms, E. S., and Kornberg, A. (1958) J. Biol. Chem. 233,
163–170
2. Kornberg, A., and Baker, T. (1992) DNA Replication, Second Ed., W. H. Freeman and
Co., New York
3. Johnson, A., and O’Donnell, M. (2005) Annu. Rev. Biochem. 74, 283–315
4. McHenry, C. S. (2003) Mol. Microbiol. 49, 1157–1165
5. O’Donnell, M., Jeruzalmi, D., and Kuriyan, J. (2001) Curr. Biol. 11, R935–R946
6. Kong, X. P., Onrust, R., O’Donnell, M., and Kuriyan, J. (1992) Cell 69, 425– 437
7. Bowman, G. D., Goedken, E. R., Kazmirski, S. L., O’Donnell, M., and Kuriyan, J. (2005)
FEBS Lett. 579, 863– 867
8. Neuwald, A. F., Aravind, L., Spouge, J. L., and Koonin, E. V. (1999) Genome Res. 9,
27– 43
9. Guenther, B., Onrust, R., Sali, A., O’Donnell, M., and Kuriyan, J. (1997) Cell 91,
335–345
10. Yu, R. C., Hanson, P. I., Jahn, R., and Brunger, A. T. (1998) Nat. Struct. Biol. 5,
803– 811
11. Lenzen, C. U., Steinmann, D., Whiteheart, S. W., and Weis, W. I. (1998) Cell 94,
525–536
12. Jeruzalmi, D., O’Donnell, M., and Kuriyan, J. (2001) Cell 106, 429 – 441
13. Jeruzalmi, D., Yurieva, O., Zhao, Y., Young, M., Stewart, J., Hingorani, M., O’Donnell,
M., and Kuriyan, J. (2001) Cell 106, 417– 428
14. Leu, F. P., and O’Donnell, M. (2001) J. Biol. Chem. 276, 47185– 47194
15. Stewart, J., Hingorani, M. M., Kelman, Z., and O’Donnell, M. (2001) J. Biol. Chem.
276, 19182–19189
10656 JOURNAL OF BIOLOGICAL CHEMISTRY
VOLUME 281 • NUMBER 16 • APRIL 21, 2006
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The ␤ clamp interacts with many different proteins, including all five E. coli
DNA polymerases, MutS, ligase, and UvrB. These various proteins bind to the
hydrophobic site in ␤ to which ␦ binds (54, 60 – 62). E. coli ␣-polymerase
subunit contains two regions that bind ␤ (62). The extreme C-terminal
sequence binds the hydrophobic site on ␤, and the second region is about 200
residues internal to the C terminus (53, 62). Mutations in either region alter the
affinity of ␣ for ␤. The structure of the C-terminal domain of pol IV bound to
␤ also reveals two different sites of interaction with ␤ (63). The extreme C-terminal residues of pol IV bind to one of the hydrophobic pockets in the ␤ dimer,
while an internal region of pol IV binds at an edge of the ␤ ring.
The fact that sliding clamps bind numerous proteins has led to speculation that
multiple proteins may bind the clamp at the same time (64, 65). In this “toolbelt”
hypothesis, the clamp brings together factors that are needed sequentially on
DNA. Toolbelt function has been demonstrated for E. coli ␤ in which pol IV and
pol III form a ternary complex with ␤ (66). pol IV is a low fidelity Y-family polymerase enabling it to bypass lesions that block the replicase (67– 69). When pol III
stalls, pol IV rapidly gains control of ␤ and the primed template. When the block is
relieved, pol III regains the primed template. These protein dynamics limit the
action of the low fidelity Pol IV to the locale of the block.
16. Turner, J., Hingorani, M. M., Kelman, Z., and O’Donnell, M. (1999) EMBO J. 18,
771–783
17. Naktinis, V., Onrust, R., Fang, L., and O’Donnell, M. (1995) J. Biol. Chem. 270,
13358 –13365
18. Bowman, G. D., O’Donnell, M., and Kuriyan, J. (2004) Nature 429, 724 –730
19. Goedken, E. R., Kazmirski, S. L., Bowman, G. D., O’Donnell, M., and Kuriyan, J. (2005)
Nat. Struct. Mol. Biol. 12, 183–190
20. Trakselis, M. A., Alley, S. C., Abel-Santos, E., and Benkovic, S. J. (2001) Proc. Natl.
Acad. Sci. U. S. A. 98, 8368 – 8375
21. Kazmirski, S. L., Zhao, Y., Bowman, G. D., O’Donnell, M., and Kuriyan, J. (2005) Proc.
Natl. Acad. Sci. U. S. A. 102, 13801–13806
22. Miyata, T., Suzuki, H., Oyama, T., Mayanagi, K., Ishino, Y., and Morikawa, K. (2005)
Proc. Natl. Acad. Sci. U. S. A. 102, 13795–13800
23. Ason, B., Handayani, R., Williams, C. R., Bertram, J. G., Hingorani, M. M., O’Donnell,
M., Goodman, M. F., and Bloom, L. B. (2003) J. Biol. Chem. 278, 10033–10040
24. Naktinis, V., Turner, J., and O’Donnell, M. (1996) Cell 84, 137–145
25. Yuzhakov, A., Turner, J., and O’Donnell, M. (1996) Cell 86, 877– 886
26. Kim, S., Dallmann, H. G., McHenry, C. S., and Marians, K. J. (1996) Cell 84, 643– 650
27. Patel, S. S., and Picha, K. M. (2000) Annu. Rev. Biochem. 69, 651– 697
28. Sawaya, M. R., Guo, S., Tabor, S., Richardson, C. C., and Ellenberger, T. (1999) Cell 99,
167–177
29. Singleton, M. R., Sawaya, M. R., Ellenberger, T., and Wigley, D. B. (2000) Cell 101,
589 – 600
30. Delagoutte, E., and von Hippel, P. H. (2003) Q. Rev. Biophys. 36, 1– 69
31. Jezewska, M. J., Rajendran, S., Bujalowska, D., and Bujalowski, W. (1998) J. Biol. Chem.
273, 10515–10529
32. Kaplan, D. L. (2000) J. Mol. Biol. 301, 285–299
33. Johanson, K. O., and McHenry, C. S. (1984) J. Biol. Chem. 259, 4589 – 4595
34. Glover, B. P., and McHenry, C. S. (2001) Cell 105, 925–934
35. Wu, C. A., Zechner, E. L., Hughes, A. J., Jr., Franden, M. A., McHenry, C. S., and
Marians, K. J. (1992) J. Biol. Chem. 267, 4064 – 4073
36. Maki, H., Maki, S., and Kornberg, A. (1988) J. Biol. Chem. 263, 6570 – 6578
37. Onrust, R., Finkelstein, J., Naktinis, V., Turner, J., Fang, L., and O’Donnell, M. (1995)
J. Biol. Chem. 270, 13348 –13357
38. Kaplan, D. L., and O’Donnell, M. (2002) Mol. Cell 10, 647– 657
39. Takahashi, T. S., Wigley, D. B., and Walter, J. C. (2005) Trends Biochem. Sci. 30,
437– 444
40. Tye, B. K. (1999) Annu. Rev. Biochem. 68, 649 – 686
41. Mendez, J., and Stillman, B. (2003) BioEssays 25, 1158 –1167
42. Kaplan, D. L., Davey, M. J., and O’Donnell, M. (2003) J. Biol. Chem. 278, 49171– 49182
43. Lee, J. K., and Hurwitz, J. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 54 –59
44. Webb, C. D., Teleman, A., Gordon, S., Straight, A., Belmont, A., Lin, D. C.-H., Grossman, A. D., Wright, A., and Losick, R. (1997) Cell 88, 667– 674
45. Lemon, K. P., and Grossman, A. D. (2000) Mol. Cell 6, 1321–1330
46. Smelkova, N. V., and Borowiec, J. A. (1997) J. Virol. 71, 8766 – 8773
47. Chong, J. P., Hayashi, M. K., Simon, M. N., Xu, R. M., and Stillman, B. (2000) Proc.
Natl. Acad. Sci. U. S. A. 97, 1530 –1535
48. Breier, A. M., Weier, H. U., and Cozzarelli, N. R. (2005) Proc. Natl. Acad. Sci. U. S. A.
102, 3942–3947
49. Alberts, B. M., Barry, J., Bedinger, P., Formosa, T., Jongeneel, C. V., and Kreuzer, K. N.
(1983) Cold Spring Harbor Symp. Quant. Biol. 47, 655– 668
50. Tougu, K., Peng, H., and Marians, K. J. (1994) J. Biol. Chem. 269, 4675– 4682
51. O’Donnell, M. E. (1987) J. Biol. Chem. 262, 16558 –16565
52. Stukenberg, P. T., Turner, J., and O’Donnell, M. (1994) Cell 78, 877– 887
53. Lopez de Saro, F. J., Georgescu, R. E., and O’Donnell, M. (2003) Proc. Natl. Acad. Sci.
U. S. A. 100, 14689 –14694
54. Lopez de Saro, F. J., and O’Donnell, M. (2001) Proc. Natl. Acad. Sci. U. S. A. 98,
8376 – 8380
55. Leu, F. P., Hingorani, M. M., Turner, J., and O’Donnell, M. (2000) J. Biol. Chem. 275,
34609 –34618
56. Lee, J., Chastain, P. D., II, Kusakabe, T., Griffith, J. D., and Richardson, C. C. (1998)
Mol. Cell 1, 1001–1010
57. Salinas, F., and Benkovic, S. J. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 7196 –7201
58. McInerney, P., and O’Donnell, M. (2004) J. Biol. Chem. 279, 21543–21551
59. Higuchi, K., Katayama, T., Iwai, S., Hidaka, M., Horiuchi, T., and Maki, H. (2003)
Genes Cells 8, 437– 449
60. Lopez de Saro, F. J., Georgescu, R. E., Goodman, M. F., and O’Donnell, M. (2003)
EMBO J. 22, 6408 – 6418
61. Dalrymple, B. P., Kongsuwan, K., Wijffels, G., Dixon, N. E., and Jennings, P. A. (2001)
Proc. Natl. Acad. Sci. U. S. A. 98, 11627–11632
62. Dohrmann, P., and McHenry, C. S. (2005) J. Mol. Biol. 350, 228 –239
63. Bunting, K. A., Roe, S. M., and Pearl, L. H. (2003) EMBO J. 22, 5883–5892
64. Fujii, S., and Fuchs, R. P. (2004) EMBO J. 23, 4342– 4352
65. Warbrick, E. (2000) BioEssays 22, 997–1006
66. Indiani, C., McInerney, P., Georgescu, R., Goodman, M. F., and O’Donnell, M. (2005)
Mol. Cell 19, 805– 815
67. Lenne-Samuel, N., Wagner, J., Etienne, H., and Fuchs, R. P. (2002) EMBO Rep. 3,
45– 49
68. Goodman, M. F. (2002) Annu. Rev. Biochem. 71, 17–50
69. Napolitano, R., Janel-Bintz, R., Wagner, J., and Fuchs, R. P. (2000) EMBO J. 19,
6259 – 6265