Toward understanding the conformational dynamics of RNA ligation.

Abstract

Members of the genus Trypanosoma, which include the pathogenic species Trypanosoma brucei and Trypanosoma cruzi, edit their post-transcriptional mitochondrial RNA via a multiprotein complex called the editosome. In T. brucei, the RNA is nicked prior to uridylate insertion and deletion. Following editing, nicked RNA is religated by one of two RNA-editing ligases (TbREL). This study describes a recent 70 ns molecular dynamics simulation of TbREL1, an ATP-dependent RNA-editing ligase of the nucleotidyltransferase superfamily that is required for the survival of T. brucei insect and bloodstream forms. In this work, a model of TbREL1 in complex with its full double-stranded RNA (dsRNA) substrate is created on the basis of the homologous relation between TbREL1 and T4 Rnl2. The simulation captures TbREL1 dynamics in the state immediately preceding RNA ligation, providing insights into the functional dynamics and catalytic mechanism of the kinetoplastid ligation reaction. Important features of RNA binding and specificity are revealed for kinetoplastid ligases and the broader nucleotidyltransferase superfamily.

Full text

Page 1

Toward Understanding the Conformational Dynamics of RNA Ligation†
Robert V. Swift,‡,§Jacob Durrant,§,|Rommie E. Amaro,*,‡and J. Andrew McCammon‡,⊥
Department of Chemistry and Biochemistry, Department of Pharmacology, and NSF Center for Theoretical Biological Physics,
UniVersity of California at San Diego, La Jolla, California 92093-0365, Biomedical Sciences Graduate Program, UniVersity of
California at San Diego, La Jolla, California 92093-0365, and Howard Hughes Medical Institute, UniVersity of California at
San Diego, La Jolla, California 92093-0365
ReceiVed September 23, 2008; ReVised Manuscript ReceiVed December 4, 2008
ABSTRACT: Members of the genus Trypanosoma, which include the pathogenic species Trypanosoma brucei
and Trypanosoma cruzi, edit their post-transcriptional mitochondrial RNA via a multiprotein complex
called the editosome. In T. brucei, the RNA is nicked prior to uridylate insertion and deletion. Following
editing, nicked RNA is religated by one of two RNA-editing ligases (TbREL). This study describes a
recent 70 ns molecular dynamics simulation of TbREL1, an ATP-dependent RNA-editing ligase of the
nucleotidyltransferase superfamily that is required for the survival of T. brucei insect and bloodstream
forms. In this work, a model of TbREL1 in complex with its full double-stranded RNA (dsRNA) substrate
is created on the basis of the homologous relation between TbREL1 and T4 Rnl2. The simulation captures
TbREL1 dynamics in the state immediately preceding RNA ligation, providing insights into the functional
dynamics and catalytic mechanism of the kinetoplastid ligation reaction. Important features of RNA binding
and specificity are revealed for kinetoplastid ligases and the broader nucleotidyltransferase superfamily.
The intricate processing of genetic information is central
to biological complexity and to the replication processes that
sustain life. The discovery of post-transcriptional RNA
editing was a rich addition to our appreciation of the central
dogma of biology and is still a vibrant area of research with
many unanswered questions. Within the mitochondria of
kinetoplastids, a unique form of RNA editing exists whereby
uridylates are inserted or deleted (1). To carry out the editing
process, kinetoplastids utilize a multiprotein complex known
as the editosome. Though many details remain unresolved,
the editosome is known to include a core 20S complex of
1.6 MDa containing 16-20 different proteins. Additionally,
at least three distinct 20S complexes, distinguished by their
constituent proteins and specificity for insertion or deletion
editing, have been detected (2), suggesting that an inherent
dynamism in the 20S core may accommodate the varied
specificity requirements of different gene products.
The editosome is localized in the mitochondria, where
mitochondrial DNA has a topologically linked network of
thousands of minicircles, which encode guide RNAs (gR-
NAs),1and dozens of maxicircles, which encode components
of respiratory complexes and energy transduction systems.
After transcription, gRNAs base pair with pre-mRNA from
maxicircles via conserved “anchor sequences” (3, 4). Fol-
lowing gRNA base pairing, an endonucleolytic component
of the editosome cleaves the pre-mRNA at points of
nucleotide mismatch between the pre-mRNA and the gRNA.
Depending on the type of mismatch, uridylates are then
inserted or removed from the nicked site of the pre-mRNA
by a terminal uridydyl transferase or a U-specific 3′ exonu-
clease, respectively (5, 6). One of two RNA-editing ligases,
TbREL1 or TbREL2, then religates the nicked RNA, gener-
ally depending on whether the preceding mRNA processing
was insertion or deletion, respectively. The religated, “ma-
ture” mRNA is then ready for translation. TbREL1 is relevant
to structure-based rational drug design and associated
structure-function studies both because its catalytically
competent N-terminal adenyltransferase domain has been
crystallized in complex with ATP at 1.2 Å resolution (7)
and because it is required for the survival of Trypanosoma
brucei insect and bloodstream forms (8, 9).
An ATP-dependent RNA ligase, TbREL1 belongs to the
nucleotidyltransferase superfamily, whose members include
ATP and NAD+-dependent DNA ligases as well as eukary-
otic mRNA capping enzymes (10). Each superfamily member
shares five conserved active site structural motifs, along with
a conserved catalytic mechanism in which a nucleotide is
transferred to the 5′ end of a polynucleotide via a covalent
enzyme-NMP intermediate. Catalysis of dsRNA by TbREL1
†This work was supported by the National Institutes of Health (Grant
F32-GM077729 to R.E.A. and Grant GM31749 to J.A.M.), the National
Science Foundation (Grant MRAC CHE060073N to R.E.A. and Grants
MCB-0506593 and MCA93S013 to J.A.M.), and the Howard Hughes
Medical Institute (to J.A.M.).
* To whom correspondence should be addressed: Department of
Chemistry and Biochemistry, University of California at San Diego,
9500 Gilman Dr., Mail Code 0365, La Jolla, CA 92093-0365. Phone:
(858)822-0169. Fax:(858)
mccammon.ucsd.edu.
‡Department of Chemistry and Biochemistry, Department of
Pharmacology, and NSF Center for Theoretical Biological Physics.
§These authors contributed equally to this work.
|Biomedical Sciences Graduate Program.
⊥Howard Hughes Medical Institute.
534-4974. E-mail:ramaro@
1Abbreviations: TbREL, T. brucei RNA-editing ligase; MD, mo-
lecular dynamics; gRNA, guide RNA; LIG1, DNA ligase 1; AppN,
AMP attached to the nick 5′-PO4 moiety; rmsd, root-mean-square
deviation; rmsf, root-mean-square fluctuation; dsRNA, double-stranded
RNA.
Biochemistry 2009, 48, 709–719709
10.1021/bi8018114 CCC: $40.75
 2009 American Chemical Society
Published on Web 01/09/2009

Page 2

is initiated when a conserved catalytic lysine nucleophile
attacks an ATP R-phosphate, displacing pyrophosphate and
forming an enzyme-AMP intermediate. In the second step,
nicked dsRNA binds and AMP is transferred from the
enzyme to the 5′-PO4 end of the nick, forming a 5′-5′
phosphoanhydride linkage. In the third step, the 3′-OH end,
which apposes the 5′-PO4, attacks the phosphoanhydride
linkage, displacing AMP and forming the religated dsRNA
product.
Though the three religation steps of TbREL1 are widely
acknowledged, the exact dependence on divalent metal
cofactors remains in doubt. For example, though the electron
density map of the ATP-bound TbREL1 N-terminal domain
clearly shows a Mg2+coordinated between the nonbridging
oxygen atoms of the ATP ?- and γ-phosphates (7), nucleotide
transfer has not yet occurred, suggesting that a single Mg2+
may be insufficient to trigger the first chemical step, though
the orthogonal position of the lysine nucleophile relative to
the R-phosphate electrophile, which hinders nucleophilic
attack, is an alternative explanation (11). Additionally, kinetic
evidence for related superfamily members suggests that the
nucleotidyltransferase step proceeds via a two-Mg2+mech-
anism (12), while crystal structure data from both the
PBCV-1 capping enzyme and T4 RNA ligase 2 (Rnl2) show
that a divalent metal cofactor in the vicinity of the R-phos-
phate remains after step 1 of catalysis (11, 13) and imply
that a two-metal mechanism may optimize the nucleotide
transfer turnover rate. These contrasting results suggest that
an allosteric signal, possibly from other protein members of
the editosome, may lead to the uptake and coordination of a
second Mg2+, thereby triggering nucleotide transfer (7). The
plausibility of ATP-dependent allosteric communication
between TbREL1 and other editosome members was dem-
onstrated in molecular dynamics (MD) simulations of apo
and ATP-bound TbREL1 (14). These simulations suggested
that ATP binding induces increased mobility of a putative
allosteric loop; similar phenomena may provide the impetus
for the uptake of a second Mg2+.
Both sequence and structural phylogenetic analyses of
superfamily members indicates that the closest known
TbREL1 homologue is the T4 phage Rnl2 (7, 14). Recently,
the structure of full-length Rnl2, in complex with its nucleic
acid substrate, was captured in two different conformations
(13). These two unique conformations illustrate the active
site conformational rearrangements that occur as the complex
proceeds from the step 2 product, immediately following step
2 chemistry, to the step 3 substrate, immediately preceding
step 3 chemistry. Additionally, comparing the crystal struc-
ture of human DNA ligase 1 (LIG1), in complex with its
DNA substrate, to the Rnl2 nucleic acid complex reveals
the salient features of substrate conformational requirements
and substrate specificity. In both LIG1 and Rnl2, the
nucleotides local to the nick are found in a RNA-like
A-conformation. This conformation aligns the 3′-OH and
apposing 5′-PO4at the nick site and is likely required for
step 3 chemistry. Normally in the canonical B-form, the DNA
substrate of LIG1 is distorted into the required A-conforma-
tion via two phenylalanines that intercalate the minor grove.
These phenylalanines, conserved in DNA ligases, are absent
in RNA ligases like Rnl2 and TbREL1. Further details are
presented in the original crystal structure paper (13) and are
FIGURE 1: Nicked dsRNA bound to TbREL1, poised to catalyze religation. A unique kinetoplastid insert is colored yellow, and a cluster of
residues thought to be important in RNA recognition on the 5′-PO4side of the nick is colored orange. The magnified inset shows protein
residues that make stabilizing contacts with the RNA.
FIGURE 2: Equilibrated structure illustrating the close contacts
between key active site residues (shown in licorice), the AppRNA
(shown in ball and stick), and the 3′-OH RNA strand.
710
Biochemistry, Vol. 48, No. 4, 2009
Swift et al.

Page 3

reviewed elsewhere in the context of other DNA and RNA
ligases (15).
Motivated by the close homology between Rnl2 and
TbREL1 and the desire to understand the functional dynamics
of the kinetoplastid ligation reaction, we investigate the
conformational dynamics of TbREL1 in complex with a
nicked dsRNA substrate in its canonical A-form. On the basis
of the homologous structural and functional relationships
between TbREL1 and T4 Rnl2, we develop a model of a
nicked dsRNA-TbREL1 complex (Figure 1) and perform
an all-atom explicitly solvated molecular dynamics simula-
tion of the system. The integration of RNA into MD
simulations has benefited from recent improvements in both
force field development and algorithms (16, 17), allowing
us to gain insights into the dynamics and catalytic mecha-
nism. The simulated system contains AMP attached to the
nick 5′-PO4 moiety (AppN), characteristic of the step 2
intermediate (13). The non-nicked strand represents the
template, or gRNA, and the nicked strand represents the post-
transcriptionally modified pre-mRNA. Fluctuations of con-
served interactions between the enzyme and RNA substrate,
as well as the dynamics of conserved active site residues,
are examined and discussed in the context of step 3 of the
religation mechanism. Furthermore, we explore the effect
of a unique kinetoplastid insert region on dsRNA binding.
MATERIALS AND METHODS
System Preparation. A model of the step 2 intermediate
of TbREL1 in complex with A-form dsRNA was built in
homology to chain B of the T4 Rnl2 crystal structure [Protein
Data Bank (PDB) entry 2HVR], which captures T4 Rnl2
immediately preceding step 3 chemistry (13). A 12-mer of
A-form dsRNA, with a sequence identical to that of the
nicked DNA-RNA hybrid cocrystallized with T4 Rnl2, was
generated using the NUCGEN module of Amber 9 (18),
which creates accurate nucleotide models based on fiber
diffraction studies. The dsRNA was then aligned with the
nicked DNA-RNA hybrid by minimizing the root-mean-
square deviation (rmsd) between the two nitrogenous bases
of the DNA-RNA hybrid that flanked the nick and the
commensurate nitrogenous bases of the model dsRNA. Next,
the dsRNA was translated onto the crystal structure of the
TbREL1 N-terminus (PDB entry 1XDN) by minimizing the
rmsd between the AppN adenine moiety of the DNA-RNA
hybrid and the adenine moiety of the ATP bound in the
TbREL1 active site. A nick was then created in the model
dsRNA identical to the nick of the DNA-RNA hybrid bound
to T4 Rnl2. The ?- and γ-phosphates of the TbREL1-bound
ATP were subsequently deleted, and the O-PAMP-O-P5′PO4
dihedral angle was brought into agreement with the corre-
sponding dihedral angle of the T4 Rnl2 crystal structure. The
resulting AMP is linked through its phosphate to the 5′-PO4
nicked terminal nucleotide; we designate this AMP-phosphate-
phosphate-nucleotide residue AppN according to the no-
menclature of Nandakumar and co-workers (13).
To ensure that the active site of our model reflected the
conformation of Rnl2 immediately preceding step 3 chem-
istry, TbREL1 active site residue R111 was given a confor-
mation commensurate with its Rnl2 homologue, R55.
Specifically, R111 was rotated away from the nonbridging
oxygen of the AppN R-phosphate and placed within hydro-
gen bonding distance of the Y58 backbone carbonyl. The
conformations of the other TbREL1 active site residues were
in close agreement with the conformations of the homologous
active site residues of T4 Rnl2 immediately preceding step
3 chemistry and so required no further manipulation. A Mg2+
cation was manually positioned approximately 2 Å from a
nonbridging AMP R-phosphate on the basis of the assump-
tions that ion dissociation occurs neither during RNA binding
nor during subsequent active site rearrangements and that
the TbREL1 catalytic mechanism proceeds via a two-Mg2+
mechanism. This Mg2+position was chosen on the basis of
the crystal structure data of both the PBCV-1 capping
enzyme (11) and the T4 Rnl2 step 1 product (13), as well as
on a similarly positioned Ca2+cation in the crystal structure
of T4 Rnl1 bound to the unreactive analogue AMPCPP (19).
Five additional water molecules, each approximately 2 Å
from the Mg2+cation, were manually positioned to duplicate
the expected octahedral coordination complex.
All cocrystallized water molecules in the TbREL1 structure
were retained in our model. Selenomethionines used for
crystal structure refinement were replaced with methionines,
and WHATIF (20) was used to assign histidine protonation
states. Protonation state assignments were manually verified.
With the exception of the nonstandard AppN residue, all
RNA and protein hydrogens and force field parameters were
assigned according to the Amber ff99SB force field (21)
using the xleap module in Amber 9 (18). The nonstandard
residue AppN is composed of an adenosine linked to a
cytidine via a phosphoanhydride linkage; consequently, we
assigned AppN charges using the RA5 and RC5 residues of
the Amber ff99SB force field as a template. The charge of
the oxygen bridging the phosphoanhydride linkage was
assumed to be identical to that of the oxygen bridging the
R- and ?-phosphates in the Carlson ADP parameter set (22).
All missing AppN force field parameters came from the
GAFF force field (23). Following parameterization, the
complex was immersed in a rectangular box of TIP3P waters
(24) with a 10 Å buffer from the protein to the edge of the
water box in each direction. Sodium and chloride ions were
added to the system, bringing it to electric neutrality and
yielding a 100 mM salt solution.
MD Simulations. Spurious contacts were removed through
a four-tier set of 45000 energy minimization steps. The first
10000 steps were performed in two 5000-step cycles.
Hydrogen atoms were relaxed during the first 5000 steps,
all other atoms being held fixed. Hydrogen atoms, water
molecules, and ions were relaxed during the next 5000 steps,
the protein and nucleic acid being held fixed. Ten thousand
steps of minimization followed in which only the protein
backbone was fixed. No constraints were applied during the
last 25000 steps of minimization.
To equilibrate our model, we performed a 1 ns restrained
simulation using the minimized system as a starting point.
The temperature was introduced at 310 K with harmonic
restraints on the protein backbone. Through the gradual
relaxation of the backbone restraints, the protein structure
was preserved as kinetic energy was introduced. For the first
250 ps, a harmonic restraining force of 4.0 kcal mol-1Å-2
was applied to the backbone atoms. Similarly, for the second,
third, and fourth 250 ps segments, the harmonic restraints
were consistently weakened to 3.0, 2.0, and 1.0 kcal mol-1
Å-2, respectively.
Conformational Dynamics of RNA Ligation
Biochemistry, Vol. 48, No. 4, 2009 711

Page 4

A 70 ns MD simulation followed in which unrestrained
dynamics were performed with a 1 fs time step. The duration
of the simulation was sufficient to equilibrate the system on
the basis of rmsd values [overall R-carbon (Figure S1 of the
Supporting Information) and the 3′-terminal sugar pucker
phase angle (Figure 3)], while allowing us to sample
functionally relevant conformations local to the step 2
intermediate. The temperature bath was maintained by
Langevin dynamics at 310 K, and pressure was maintained
with the hybrid Nose Hoover-Langevin piston method at 1
atm (25) using period and decay times of 100 and 50 fs,
respectively. The particle mesh Ewald algorithm was used
to treat long-range electrostatics (26). A multiple-time step
algorithm was employed in which bonded interactions were
computed at every time step, short-range nonbonded interac-
tions were computed every two time steps, and full electro-
statics were computed every two time steps. The water
hydrogen-oxygen and hydrogen-hydrogen distances were
constrained using the SHAKEH algorithm (27) to be within
0.0005 Å of the nominal lengths. All minimization and MD
simulations were carried out with NAMD 2.6 (28) using the
Amber ff99SB force field (21). Simulations were performed
on the TeraGrid ABE cluster. A typical benchmark for the
57575-atom system on 128 processors was 0.21 days per
nanosecond of simulation. System configurations were
sampled every 50 ps, generating 1400 coordinate snapshots
for subsequent analysis.
Pucker Analysis. The pseudorotational formalism proposed
by Rao, Westhof, and Sundaralingam (29) was used to
characterize the puckering of the nick 3′-ribose ring. We first
defined the following Fourier sums:
A)2
5∑θjcos(
B)-2
4πj
5)
4πj
5)
5∑θjsin(
where j ranges from 0 to 4 and θ0, θ1, θ2, θ3, and θ4are the
torsion angles about the C2′-C3′, C3′-C4′, C4′-O4′,
O4′-C1′, and C1′-C2′ bonds, respectively. The pucker
phase angle, P, identifies which part of the ring is furthest
from planarity. Ribose configurations generally take on the
2′-endo pucker typical of DNA or the 3′-endo pucker typical
of RNA. The possible values of P, which range from 0 to
360, correspond to the stages along the so-called pseudoro-
tation pathway. The ribose ring has five atoms, each of which
can be above or below the plane, yielding 10 distinct stages.
The envelope conformations correspond to P values of 0°,
36°, 72°, etc., and alternate with the twist conformations at
P values of 18°, 54°, 90°, etc. In the Rao-Westhof-Sundar-
alingam pseudorotational formalism
P)tan-1(
B
A)
The maximal torsion angle or puckering amplitude, θm,
measures the magnitude of the deviation from planarity (29).
θm)√A2+B2
Polar plots are generally used to represent sugar pucker,
with the puckering amplitude and phase angle assigned as
the radial and angular coordinate, respectively. Analysis of
experimental structures shows that the 2′-endo ribose con-
figuration typical of DNA has a P between 137° and 194°,
and the 3′-endo ribose configuration typical of RNA has a
P between -1° and 34° (30).
Principal Component Analysis. Principal component analy-
sis, or PCA, is a common statistical data analysis technique
whose application to the field of biomolecular simulation
was pioneered by Berendsen (31) and Garcia (32). By
diagonalizing a covariance matrix constructed of molecular
coordinates sampled over the duration of a MD trajectory
and listing the resulting eigenvectors in order of descending
eigenvalue magnitude, PCA separates large, low-frequency
conformational changes from small, high-frequency motions.
By considering only the conformational fluctuations of the
first few modes, researchers frequently use PCA to filter out
large isomerization events and to reduce the complexity of
trajectory analysis. In this work, PCA was used to explore
the large, low-frequency motions of the dsRNA substrate.
Following trajectory equilibration, principal component
analysis was carried out on the phosphate atoms of the
FIGURE 3: Equilibrated ribofuranose pseudorotation angle for the 3′-OH nucleotide as depicted within the active site (left panel) and in
polar coordinates (right panel; equilibration data colored gray and dynamics data black).
712
Biochemistry, Vol. 48, No. 4, 2009
Swift et al.

Page 5

dsRNA substrate, using the ptraj module in Amber 9 (18).
Prior to the performance of PCA, translational and rotational
motions were removed by fitting the sampled coordinates to
the first equilibrated frame of the trajectory.
RESULTS AND DISCUSSION
Equilibration. A 70 ns constraint-free simulation was
performed on the dsRNA-bound TbREL1 system to examine
TbREL1 dynamics. A plot of the time evolution of the rmsd
of the protein R-carbons (Figure S1 of the Supporting
Information), in which each trajectory frame was aligned with
the initial frame to remove rotational and translational
motion, suggested that our preliminary efforts to equilibrate
the system prior to our 70 ns simulation were successful.
However, subsequent examination of the distance between
the nick 3′-OH and the phosphorus atom of the 5′-phospho-
anhydride linkage suggested that equilibration was not
complete until after 38.75 ns. During the initial phase
(0-38.75 ns), these two atoms, between which a bond forms
following step 3, were distant (7.16 ( 1.20 Å). During the
second phase (38.75-70 ns), however, the nick 3′-OH and
the nick 5′-phosphorus moved closer together (3.95 ( 0.29
Å), as required for catalysis. Correspondingly, the trajectory
time from 0 to 38.75 ns is called the “equilibration phase”
and from 38.75 to 70 ns the “dynamics phase”. Unless
otherwise specified, all analysis was performed on the
dynamics phase.
ActiVe Site Interactions. The atomic interactions within
the active site can be divided into three categories: interac-
tions between TbREL1 and the adenine moiety of AppN,
interactions between TbREL1 and the ribofuranose of AppN,
and interactions between TbREL1 and the pyrophosphate of
AppN. Throughout our discussion, we adopt the naming
scheme of Nandakumar and co-workers, naming the nicked
strands on the basis of their active site ends (i.e., the 5′-PO4
strand and the 3′-OH strand) (13).
Deep within the active site, F209, E86, and V88 interact
with the adenine moiety of AppN and stabilize its motion
(Figure 2). F209 acts as a conserved hydrophobic platform
that forms π-π interactions with the aromatic adenine
moiety of AppN, preventing large lateral fluctuations of this
base within the binding pocket. Hydrophobic stabilization
in this position is essential for step 3 catalysis in other type
II RNA ligases, as demonstrated by the partial recovery of
religation activity when a leucine residue was substituted
for the F209 homologue in an alanine scan of T4 Rnl2 (33).
Our simulation is consistent with this mutagenesis study;
F209 exhibits minor fluctuations, with an average distance
of 4.14 Å between the centers of mass of its phenyl group
and the AppN adenine moiety (Table 1). The movement of
AppN is also restrained by E86, which accepts a hydrogen
bond from the adenine N6 amine that holds O1 (or O2) of
E86 within 2.97 Å, on average (Table 1). Conserved among
type II RNA ligases, E86 is essential in mediating step 3
catalysis in T4 Rnl2 (33). The observed distances and
fluctuations of both F209 and E86 are similar to those
observed in a previous simulation of the ATP-TbREL1
complex (14). These results, along with the T4 Rnl2
mutational analysis (33), reflect the importance of F209 and
E86 in mediating the religation of the adenylated dsRNA
substrate and the formation of the adenylated enzyme
intermediate.
Stabilizing interactions between the protein and the AppN
ribofuranose complement the contacts formed between the
adenine moiety and residues deep within the binding pocket.
In particular, E159, conserved across all members of the
superfamily, and N92, conserved across all type II RNA
editing ligases (33), make critical hydrogen bonds with the
ribofuranose O2′ hydroxyl group (Figure 2 and Table 1).
The close, stable contact made by E159 was observed in a
previous simulation of the ATP-bound complex (14) and is
consistent with the conservative mutational analysis, which
identified E159 as an essential mediator of steps 1 and 3 of
catalysis (33). Similarly, an alanine scan of T4 Rnl2 revealed
that the N92 homologue plays a critical role in the religation
of the preadenylated dsRNA substrate (33). No isomerization
of the AppN ribofuranose moiety was observed during the
course of catalysis in the T4 Rnl2 homologue (13); conse-
quently, the mutational analysis results, coupled with the
short, stable contacts observed during our simulation, suggest
that the N92-ribofuranose interaction helps maintain the
ribofuranose conformation throughout the catalytic cycle.
Charged residues at the periphery of the binding site, K87,
K307, and R111, each interact with the AppN pyrophosphate
and stabilize the step 3 substrate conformation (Figure 2).
The orthogonal orientation of the K87 nucleophile to the
R-phosphate is typical of the step 3 substrate conformation
and persists throughout the 31.25 ns dynamics phase. The
system does not undergo isomerization away from the step
3 substrate conformation, indicating that the network of
interactions captured in the homologous T4 Rnl2 crystal
structure is a reasonable model of the TbREL1 active site
architecture prior to step 3 of catalysis. Interestingly, K87
remains within hydrogen bonding distance of a nonbridging
pyrophosphate oxygen during the dynamics phase of the
simulation (Table 1). This result suggests that the K87
nucleophile helps maintain the protein in the step 3 confor-
mation once isomerization into that conformation has oc-
curred. Consequently, K87 seems to perform a dual role
during catalysis: during step 1, it plays the central chemical
role as the nucleophile, and during step 3, it plays a peripheral
stabilizing role as a hydrogen bond donor. To the best of
our knowledge, no experimental evidence has been published
supporting this hypothesis, but the putative dual role of K87
Table 1: Active Site Interactions, Including Protein-AppN Interactions,
Protein-Nick Interactions, and Interactions with the Mg2+-Coordinated
Water Molecules
length (Å)
[standard deviation (Å)]
AppN adenine-F209 phenyl group
E86 O-AppN adenine N6
E159 OE1-AppN ribose O2′
N92 ND2-AppN ribose O2′
K87 NZ-AppN O1P
K307 NZ-AppN O2P
R111 NH1-AppN O1P
nick 3′-HO-nearest nonbridging
nick 5′-PO4oxygen
R111 CZ-nick O3′
H89 ND1-WAT O
E159 CD-WAT O
E283 CD-WAT O
E283 CD-WAT O
nick O2′-WAT O
4.14 (0.22)
2.97 (0.20)
2.55 (0.08)
3.30 (0.41)
3.39 (0.42)
3.87 (0.56)
2.80 (0.11)
1.90 (0.64)
3.95 (0.34)
2.85 (0.21)
3.49 (0.27)
3.45 (0.24)
3.44 (0.21)
3.30 (0.53)
Conformational Dynamics of RNA Ligation
Biochemistry, Vol. 48, No. 4, 2009 713