FRET analysis reveals distinct conformations of
IN tetramers in the presence of viral DNA or
Jacques J. Kessl1, Min Li2, Michael Ignatov3, Nikolozi Shkriabai1, Jocelyn O. Eidahl1,
Lei Feng1, Karin Musier-Forsyth3, Robert Craigie2and Mamuka Kvaratskhelia1,*
1Center for Retrovirus Research and Comprehensive Cancer Center, College of Pharmacy, The Ohio State
University, Columbus, OH 43210,2Laboratory of Molecular Biology, National Institute of Diabetes and Digestive
and Kidney Diseases, National Institutes of Health, Bethesda, MD, 20892,3Departments of Chemistry and
Biochemistry, Center for Retrovirus Research and Center for RNA Biology, The Ohio State University,
Columbus, OH 43210, USA
Received October 28, 2010; Revised June 29, 2011; Accepted June 30, 2011
A tetramer of HIV-1 integrase (IN) stably associates
with the viral DNA ends to form a fully functional
concerted integration intermediate.
a key cellular binding partner of the lentiviral
enzyme, also stabilizes a tetrameric form of IN.
However, functional assays have indicated the
importance of the order of viral DNA and LEDGF/
p75addition toIN for
integration. Here, we employed Fo ¨rster Resonance
Energy Transfer (FRET) to monitor assembly of indi-
vidual IN subunits into tetramers in the presence
of viral DNA and LEDGF/p75. The IN–viral DNA
and IN–LEDGF/p75 complexes yielded significantly
different FRET values suggesting two distinct IN
conformations in these complexes. Furthermore,
the order of addition experiments indicated that
FRET for the preformed IN–viral DNA complex
remained unchanged upon its subsequent binding
to LEDGF/p75, whereas pre-incubation of LEDGF/
p75 and IN followed by addition of viral DNA
yielded FRET very similar to the IN–LEDGF/p75
complex. These findings provide new insights into
the structural organization of IN subunits in func-
suggest that differential multimerization of IN in
the presence of various ligands could be exploited
as a plausible therapeutic target for development of
HIV-1 integrase (IN) functions as a multimer to catalyze
integration of the reverse transcribed DNA copy of the
viral genome into a host chromosome [reviewed in(1)].
The enzyme stably associates with two viral DNA ends
to form a large nucleoprotein complex termed pre-
number of viral and cellular proteins contributing to the
retroviral integration (2–16). Quantities of PICs extracted
from the infected cells are not sufficient to perform
detailed structural or even lower resolution biophysical
analyses. Therefore, purified recombinant IN and model
DNA substrates have been employed to better understand
mechanistic and structural foundations for the retroviral
Notably, recent in vitro studies (17,18) defined key con-
certed integration intermediates and provided a powerful
model system closely mimicking IN interactions with viral
DNA within PICs in the infected cells. The step-wise
interactions between IN, viral and target DNAs proceed
through formationof highly
complexes. First, a tetramer of IN associates with a pair
of viral DNA ends to form stable synaptic complexes
(SSC). In common with PICs isolated from infected
cells, the SSCs assembled in vitro are resistant to treat-
ments with high ionic strength buffers containing 1M
NaCl. The 30-processing reaction takes place within the
SSC. IN remains stably associated with the pair of viral
DNA ends after capture of a target DNA and DNA
strand transfer. This second stable complex is termed the
strand transfer complex.
*To whom correspondence should be addressed. Tel: +1 614 292 6091; Fax: +1 614 292 7766; Email: email@example.com
Published online 19 July 2011 Nucleic Acids Research, 2011, Vol. 39, No. 20 9009–9022
? The Author(s) 2011. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/3.0), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
HIV-1 IN is comprised of three structurally and mech-
anistically distinct domains including the N-terminal
domain (NTD) which coordinates a Zn2+ion, the catalytic
core domain (CCD),which
DDE motif and the highly basic C-terminal domain
(CTD). Each of these domains contribute to functional
multimerization of IN (19–22). In the absence of cognate
DNA, the full-length protein can form monomers, dimers,
tetramers or higher order oligomers and the relative
abundance of these species depends on the protein con-
centration and solution conditions (20,23–25). Structural
studies with full-length HIV-1 IN or its complex with
cognate DNA have not been successful presumably
due to limited protein solubility and inherent flexibility
of the three-domain protein. Instead, atomic structures
for individual protein domains have been determined
(21,22,26–30), which paved the way for molecular
modeling of IN–DNA interactions [see (31) for recent
review]. Most recently, the co-crystal structure of proto-
type foamy virus (PFV) IN with cognate DNA (32) has
been exploited to build a model for a tetramer of HIV-1
IN interacting with two viral DNA ends (33). In this
model, two of the four IN subunits directly bind DNA.
The other two protomers seem to play a supporting
role and contribute to IN multimerization.
The main cellular binding partner of HIV-1 IN is the
protein known as lens epithelium-derived growth factor
(LEDGF/p75). LEDGF/p75 knockdown and knockout
experiments revealed the importance of this cellular
cofactor for effective HIV-1 integration and viral replica-
tion (3,34–36). LEDGF/p75 primarily functions during
HIV-1 infection to tether PICs to active genes during
integration (15,37). The cellular protein directly interacts
with HIV-1 IN via its C-terminal domain, which is
termed the integrase binding domain (IBD) (38,39). The
N-terminal part of LEDGF/p75, which contains a PWWP
domain, nuclear localization signal and dual copy of
the AT-hook DNA binding motif [reviewed in (40)]
tightly associates with the chromatin.
In vitro functional assays have indicated that LEDGF/
p75 differentially modulates HIV-1 IN activities (41–43).
The cellular protein markedly enhances integration of
single viral DNA into the target DNA regardless of the
order of addition of proteins and DNA substrates in
the reaction mixture. In contrast, the efficiency of the bio-
logically relevant concerted integration of two viral
DNA ends strongly depends on temporal interactions
of viral DNA and LEDGF/p75 with HIV-1 IN (41–43).
Addition of substoichiometric concentrations of LEDGF/
p75 to the preassembled IN–viral DNA complex stimu-
lates the pair-wise integration of viral DNA substrates.
The preformed IN–LEDGF/p75 complex is impaired
for the biologically essential concerted integration, while
still stimulating single-end integration reactions (41–43).
with LEDGF/p75 indicated that the cellular cofactor
promotes IN tetramerization (41). While IN in the func-
tional SSC is also a tetramer, mass spectrometry
(MS)-based protein footprinting of IN–viral DNA and
IN–LEDGF/p75 complexes uncovered the following im-
portant differences. HIV-1 IN undergoes significant
HIV-1 IN interactions
conformational change upon binding with cognate DNA
involving the flexible connecting segment between the
CCD and the CTD (44). In contrast, such changes have
not been observed upon LEDGF/p75 binding to IN (41).
Taken together, these observations led us to hypothesize
that tetrameric forms of IN in complex with viral DNA
and LEDGF/p75 are not identical, and that LEDGF/p75
binding modulates the structures of free IN and the IN–
viral DNA complex. To test this premise, we monitored
how individual IN subunits are organized in their
complexes with viral DNA and LEDGF/p75 using
Fo ¨ rster ResonanceEnergy
findings reveal two distinct IN tetramers formed in the
presence of viral DNA or LEDGF/p75.
Transfer (FRET). Our
MATERIALS AND METHODS
Preparation of recombinant proteins and DNA substrates
Full-length wild-type and mutated (C56S/C65S) integrase
proteins were expressed in Escherichia coli. The point
mutations were introduced in the wild-type IN sequence
usinga QuikChange Site-Directed
(Stratagene, CA, USA). Wild-type and mutant full-length
IN proteins were purified as described previously (41,45).
The isolated CTD (213–288) was prepared similarly to the
reported protocol (30). Purified recombinant LEDGF/p75
and LEDGF/IBD were obtained as described previously
(14,38,46). The blunt-end viral DNA substrate (?1kb)
was obtained by PCR using Phusion polymerase (New
England Biolabs, MA, USA), pU3U5 plasmid (47) and
the following primers: dra3 (5-GATGGTTCACGTAGT
GGGCC-3) and u5r (5-ACTGCTAGAGATTTTCCACA
CTG-3). Viral DNA was further purified by agarose gel
prior to its use in our assays. Double-stranded DNA
(3kb) derived from pGEM (Promega, WI, USA) was
used as a non-specific DNA for control experiments.
Scaling-up of SSC preparations
The reported protocol (48) for the assembly of SSCs was
modified as follows to allow 100-fold scale-up. To trap the
SSC and prevent formation of the strand transfer complex
purified wild-type IN or the mutant (C56/65S) protein
labeled with fluorophore were incubated with 80nM
1-kb blunt-ended viral DNA and 10mM IN strand
transfer inhibitor 118-D-24 (49) obtained from the the
NIH AIDS Research and Reference Reagent Program.
The reaction buffer contained 20mM HEPES, pH 7.5,
12% DMSO, 5mM DTT, 10% PEG 6000, 10mM
MgCl2, 20mM ZnCl2 and 100mM NaCl. The reaction
mixture was pre-incubated on ice for 0.5h and then
transferred to 37?C for 2h. The mixture was centrifuged
at 10000g for 40min, the supernatant was removed and
the pellet was resuspended in the suspension buffer (20ml
of 20mM HEPES, pH 7.5, containing 1M NaCl). The
solution was then loaded onto a Quantum Prep PCR
Kleen Spin Column (Bio-Rad, Hercules, CA, USA)
pre-equilibrated with the suspension buffer. SSCs were
eluted by centrifugation at 735g for 2min. Freshly
prepared SSCs were used immediately for their biophysic-
al characterization. Purities of SSCs were examined with
9010Nucleic Acids Research, 2011,Vol.39, No. 20
gel electrophoresis through a 0.8% agarose–TBE-1M
urea gel in a TBE buffer containing 1M urea as described
previously (18). This concentration of urea in the gel and
running buffer reduces interaction of SSCs with agarose
and improves resolution
nucleoprotein complexes (18). SSCs and free DNA were
visualized by ethidium bromide staining. IN and LEDGF/
p75 proteins associated with SSCs were examined by
SDS–PAGE and visualized by western blot using mono-
clonal antibodies against IN (8G4) (50) from the NIH
AIDS Research and Reference Reagent Program and
against human LEDGF from BD Biosciences.
Size exclusion chromatography
Superdex 200 prep grade column (GE Healthcare) at
1ml/min in buffer containing 50mM HEPES (pH 7.4),
750mM NaCl and 10% glycerol. The column was
calibrated with the following proteins: bovine thyroglobu-
lin (670000 Da), bovine a-globulin (158000 Da), chicken
ovalbumin (44000 Da), horse myoglobin (17000 Da),
vitamin B12 (1350 Da). These proteins were detected by
absorbance at 280nm.
were performedusing HiLoad16/60
Site-selective labeling of HIV-1 IN with fluorophores
We used the commercially available Alexa Fluor 488 and
568 (Invitrogen, Carlsbad, CA, USA) as a donor (D) and
acceptor (A) pair with a Fo ¨ rster distance (R0) of 62A˚.
In parallel preparations, 2mM of Alexa 488 or Alexa 568
maleimide were incubated with 8mM mutant (C56/65S)
protein for 60min at room temperature in a buffer
containing 50mM HEPES, pH 7.4, 1M NaCl. The
fluorophore/protein molar ratio of 1:4 was optimal to
effectively label IN and avoid protein precipitation.
b-mercaptoethanol. Labeled proteins were then extensive-
ly dialyzed at 4?C against a buffer containing 50mM
HEPES, pH 7.4, 1M NaCl, 7.5mM CHAPS and 2mM
b-mercaptoethanol. A degree of labeling of 25±2% was
achieved in all reactions as determined by measuring
absorbance of the dye (Alexa 488 at 493nm and Alexa
568 at 575nm) and the protein at 280nm. The following
extinction coefficients were used: e493=72000 for Alexa
488, e575=92000 for Alexa 568 and e280=50800 for
Steady-state FRET measurement
Fluorescence was recorded at 25?C using a Cary Eclipse
Fluorescence Spectrophotometer (Agilent Technologies,
CA, USA). The assay buffer contained 20mM HEPES,
pH 7.5, 1M NaCl and 1mM DTT. The donor fluores-
cence was excited at 488nm and fluorescence intensities
were monitored in the range of 500–650nm. Two fluores-
cence spectra were obtained for each set of experiments:
(i) the D alone, where IN(C56/C65S) labeled with Alexa
488 was mixed with the unlabeled protein and (ii) the D–A
pair, where two preparations of IN(C56/C65S), one
labeled with Alexa 488 and another with Alexa 568 were
Time-resolved FRET measurement
Fluorescence decays were measured by time-correlated
single-photon counting using laser excitation (LifeSpec-
red, Edinburgh Instruments). The donor fluorescence
was excited at 467nm and the emission was collected at
520nm with a polarizer at the magic angle (55?). All meas-
urements were performed at 25?C using Versafluo cuvettes
(BioRad, Hercules, CA, USA) with a 3?3mm path
length. Two fluorescence decay curves were obtained for
each set of energy transfer experiments: (i) the D alone,
where IN(C56/C65S) labeled with Alexa 488 was mixed
with the unlabeled protein and (ii) the D–A pair, where
two preparations of IN(C56/C65S) one labeled with Alexa
488 and another with Alexa 568 were mixed. The fluores-
cence intensity decays were deconvolved assuming a
sum of exponentials:
where tiand ?iare lifetime components and their relative
amplitudes, respectively. The goodness of the fit was
judged byreduced chi-square
function of weighted residuals. The amplitude-weighted
lifetime <t> for the donor only or the donor and
acceptor pair was calculated by:
The average energy transfer efficiency E was calculated
? h i ¼
E ¼ 1 ??DA
where <tD> and <tDA> are the amplitude-averaged
excited state lifetimes of the donor in the absence and
presence of an acceptor, respectively.
The average distance r between the donor and acceptor
was calculated by:
where R0is the constant Fo ¨ rster distance (62A˚ for the
Alexa 488–Alexa 568 pair). Time-resolved (tr) fluorescence
complexed with LEDGF/p75 or viral DNA yielded fast
rotational correlation times (see ‘Results’ section and
Supplementary data) suggesting that the fluorophore
tethered to IN exhibited significant degree of free
motion. Therefore, in our calculations a dipole orientation
factor of 2/3 was assumed.
r ¼ R0
i ? ?DA
in its freeform and
Structures of individual IN domains were obtained from
the two domain structures of HIV-1 IN (21,22). The
CCDs were used as the common alignment feature and
the CCD–CCD interfaces were maintained throughout
Nucleic Acids Research, 2011,Vol.39, No. 20 9011
our modeling studies due to their essential role in IN
generate a molecular model for the HIV-1 IN–viral
DNA complex, we used the structure of the PFV
intasome (32) as the scaffold. We then employed our
FRET results to position CTDs in the supporting
subunits, which are absent in the reported crystal structure
(32). Next, LEDGF/IBD was docked onto the SSC by
establishing its interactions with the CCDs of one dimer
and the NTD of another dimer similar to the published
structures (51,52). For these studies, we used the online
homology modeling server ‘SWISSModel’ (53) and the
software ‘Modeller’ (54).
To build the full-length HIV-1 IN complex with
LEDGF/IBD we analyzed available crystal structures of
the two domain fragments of HIV-2 and maedi-visna virus
(MVV) INs in the complex with LEDGF/IBD (52,55).
The cellular protein interacted with the HIV-2 NTD–
CCD dimer (55), while it stabilized a tetrameric form of
the MVV NTD–CCD (52). Our published biochemical
data (41) have shown that full-length LEDGF/p75
promotes HIV-1 IN tetramerization. Therefore, in our
model LEDGF/IBD (Figure 7B) bridges between the
two dimers in agreement with our MS footpriniting data
(41) and the MVV structure (52). To extend these studies
by including CTDs, which are missing in the published
two domain structures (52,55), we considered our FRET
data for modeling of the full-length IN tetramer. The
molecular shape and global dimensions for the full-length
IN tetramer in the complex with LEDGF/IBD obtained
by small angle X-ray scattering (SAXS) experiments (56)
were also employed as an additional constraint for our
modeling experiments. The loops connecting IN domains
were refined with the ‘loop refinement’ function in
‘Modeller’ software. The ‘COOT’ software (57) was used
for building the model, resolving local clashes and
optimizing the side chain conformations of the protein.
The ‘minimization’ function of an Insight II software
package (Accelrys Inc., CA, USA) was used to refine
Interactions between IN and LEDGF/p75 can
monitored at sufficiently high concentrations of these
proteins for their biochemical and biophysical analysis.
In contrast, SSCs have previously been productively
assembled only at very dilute concentrations of donor
DNA and IN, which allowed biochemical characterization
of concerted integration intermediates (17,18), but were
not sufficient for biophysical studies. Therefore, our
initial experiments focused on scaling-up purification of
SSCs. We estimated that ?100-fold scale-up of the previ-
ously described preparations was necessary to enable
Our experimental strategy for preparation of SSCs is
depicted in Figure 1A. Reported reaction conditions
for assembly of SSCs (17,18,48) served as a starting
point. Recombinant IN and a long viral donor DNA in
combination with IN strand transfer inhibitor 118-D-24
(49) allowed us to effectively trap the SSC. The inhibitor
effectively impairs binding of the target DNA to the SSC
and prevents formation of the strand transfer complex
(18,58). To increase the yield of SSCs, reaction volumes
were increased 20-fold from 25ml used in previous studies
(17,18,48) to 500ml in our assays. Using the same buffer
conditions as previously reported (17,18,48) IN concentra-
tions in the reaction mixture were optimized as shown
in Figure 1B.
Under these conditions, IN forms both specific and
non-specific complexes with donor DNAs. To delineate
between these, we exploited the intrinsic property of
SSCs, which unlike non-specific IN–DNA interactions,
are resistant to high ionic strength conditions. The
mixture was subjected to treatments with 1M NaCl
followed by size exclusion spin column chromatography.
SSCs and free DNAs were readily eluted from the column
due to their large molecular weights, while free IN, which
dissociated from non-specific DNA sites under high ionic
strength conditions, was retained by the column. The
obtained fractions were analyzed by SDS–PAGE to
monitor relative quantities of IN in the complexes and
by non-denaturing agarose gel electrophoresis to deter-
mine the purity of the final products.
Figure 1B shows that the optimal concentration range
of IN for the assembly of SSCs under these conditions is
200–400nM. At these concentrations, free IN is predom-
inantly a dimer (59,60) (see also Supplementary Figure S1
and Supplementary Table S1). At higher protein concen-
trations, IN forms tetramers (?2mM) with subsequent
concentration increments leading to formation of higher
order oligomers and protein precipitation. To delineate
the role of IN tetramers in SSC formation, we compared
the data in Figure 1B with IN 30-processing activities
(Supplementary Figure S2) as pre-assembled IN tetramers
are active in this reaction (41). These experiments revealed
a sharp contrast between the 30-processing activities and
the formation of SSCs. The highest 30-processing activities
were detected with 800–1600nM IN, whereas these
protein concentrations were very ineffective for the SSC
assembly. These results are consistent with our earlier
observations that the highly dynamic interplay between
individualIN subunits is
concerted integration, and that a preformed IN tetramer
lacks sufficient flexibility to form the fully functional
nucleoprotein complex (41).
Figure 1C compares IN interactions with cognate donor
and non-specific DNAs. In line with the previous report
(17) IN formed SSCs resistant to 1M NaCl treatment only
with cognate DNA substrate and not with non-specific
DNA (compare lanes 6 and 7 with lanes 4 and 5 in
Non-denaturing agarose gel electrophoresis results
(Figure 1D) demonstrate successful scale-up of SSCs
(compare lanes 3–5). In the 100-fold scale-up (lane 5),
the band corresponding to the SSC was readily detectable
with ethidium bromide staining with only residual
amounts of dimerized SSCs being observed. These
optimized preparations of SSCs were employed for
further biophysical analysis.
9012 Nucleic Acids Research, 2011,Vol.39, No. 20
Previous reports (42,43) indicated the importance of the
order of viral DNA and LEDGF/p75 addition to IN for
effective concerted integration. Particularly puzzling has
been the observation that the preformed IN–LEDGF/
p75 complex is selectively defective for concerted integra-
tion (41–43). Moreover, the mechanism behind these
exhibits dual activities, with its N-terminal domain
tightly binding DNA and its C-terminal IBD directly
interacting with HIV-1 IN. Each of these properties of
the full-length protein could potentially affect SSC forma-
tion by different mechanisms. For example, we previously
demonstrated that increasing concentrations of LEDGF/
p75 effectively competed with HIV-1 IN for viral DNA
binding and inhibited the 30-processing reaction (41). In
contrast, LEDGF/IBD strongly modulated dynamic inter-
play between individual IN subunits and stimulated
the 30-processing reaction but potently impaired concerted
LEDGF/p75 and to examine how its direct interaction
with IN could affect the formation of the SSC, we
employed both, the full-length protein and LEDGF/IBD
in our studies. Addition of LEDGF/p75 to free IN with
subsequent exposure of protein–protein complexes to
donor DNA effectively impaired formation of SSCs
(Figure 2A, lane 5). Agarose gel electrophoresis results
(Figure 2B) corroborated with the western blot data.
No SSCs were observed when viral DNA was exposed
to preformedIN complexes
(Figure 2B, lane 4). Very similar results were obtained
when the above experiments were conducted with
LEDGF/IBD instead of the full-length protein (data not
shown). Therefore, we conclude that direct interactions of
LEDGF/p75 with HIV-1 IN modulate the structure of the
retroviral enzyme in a way that impairs formation of
We next examined whether under our reaction condi-
tions, LEDGF/p75 associated with the SSC (Figure 2C
and D). The SSCs prepared according to Figure 1A
were incubated with LEDGF/p75 in a binding buffer con-
taining 750mM NaCl to prevent non-specific association
of the full-length cellular protein with viral DNA.
While LEDGF/p75 potently binds DNA in low ionic
strength buffers, these interactions are inhibited at NaCl
concentrations >200mM (61). Indeed, no binding of
Figure 1. Scaled-up preparations of the SSC. (A) Experimental design. (B) Optimization of IN concentrations for the SSC assembly. Purified SSCs
were subjected to SDS–PAGE and the IN band was visualized by western blot. Lane 1, IN load; lane 2, protein markers: MagicMark XP Western
Protein Standard (Invitrogen, Carlsbad, CA, USA); lanes 3–9, SSCs assembled with increasing IN concentrations. At the optimal protein concen-
trations (200–400nM), ?20% of total IN was assembled in the SSC. (C) Comparison of HIV-1 IN interactions with specific and non-specific DNA
(nsDNA). Lane 1, protein markers: MagicMark XP Western Protein Standard (Invitrogen, Carlsbad, CA, USA); lane 2, IN load; lane 3, no DNA
was included in the reaction mixture; lanes 4 and 5, SSC assembly with nsDNA; lanes 6 and 7, SSC assembly with viral DNA. (D) Agarose gel
electrophoresis: Lane 1, viral DNA alone; lane 2, DNA markers: GeneMate Quanti-Marker 1kb (BioExpress, Kaysville, UT, USA); lane 3, the initial
(1?) scale for SSC preparations as reported previously (17); lane 4, 10-fold scale-up; lane 5, 100-fold scale-up.
Nucleic Acids Research, 2011,Vol.39, No. 209013
LEDGF/p75 with viral DNA was detected in our experi-
ments (Figure 2D, lane 6). In contrast, 750mM NaCl did
not significantly interfere with LEDGF/p75 binding to IN
(see Supplementary Figure S1) and the cellular protein
effectively interacted with the SSC (Figure 2C, lane 4).
reaction conditions, LEDGF/p75 associated with the
SSC through its biologically relevant interactions with IN.
To gain structural insight into how LEDGF/p75 affects
IN conformations, we employed protein–protein FRET.
The experimental strategy for FRET studies is outlined in
Figure 3. Two separate preparations of IN were used: one
labeled with the D and the other with the A fluorophores.
employed, which is optimal for assembly of the SSC
(Figure 1B). At these concentrations, unliganded IN is
predominantly a dimer (59,60) (see also Supplementary
Figure S1). Upon binding to viral DNA ends, two
separate dimers of IN assemble into a tetramer (17).
LEDGF/p75 also promotes IN tetramerization (41,62)
indicatethat under our
(see also Supplementary Figure S1). Assembly of two
dimers labeled with D and A fluorophores into tetramers
in the presence of viral DNA or LEDGF/p75 is expected
to yield a FRET signal. The goal of our experiments was
to compare average FRET values for IN–viral DNA and
whether IN tetramers formed in these complexes differed
from one another.
A crucial step for effective FRET experiments is to
site-selectively tether D and A dyes to IN preparations.
The Alexa fluorophores chosen for these studies contain
reactive maleimide groups enabling covalent attachment
to surface Cys residues. Wild-type HIV-1 IN contains
6 Cys residues that present a challenge for site-specific
labeling. Of these, C40 and C43 coordinate the structural
Zn2+ion and C130 is partially buried in the structure and
not surface accessible. Therefore, these residues were
expected to be chemically inert. In contrast, C56, C65
and C280 are surface exposed and could readily react
and thereby determine
Figure 2. Effects of the order of viral DNA and LEDGF/p75 additions to HIV-1 IN on the SSC assembly. (A) SDS–PAGE analysis of SSCs. Lane
1: 1/10 of IN load, lane 2: protein markers: MagicMark XP Western Protein Standard (Invitrogen, Carlsbad, CA, USA), lane 3: no DNA was
included in the reaction mixture, lane 4: the SSC assembly with IN and viral DNA, lane 5: LEDGF/p75 was pre-incubated with IN and then viral
DNA was added to the reaction. IN was visualized by western blot using the respective antibody as described in ‘Materials and Methods’ section. (B)
Non-denaturing agarose gel electrophoresis. Lane 1, DNA markers: GeneMate Quanti-Marker 1kb (BioExpress, Kaysville, UT, USA); lane 2, no IN
was included in the reaction mixture; lane 3, the SSC assembly with IN and viral DNA; lane 4, LEDGF/p75 was pre-incubated with IN and then
viral DNA was added to the reaction. Free DNA and the SSC were visualized by ethidium bromide staining. (C) Experimental design to probe
LEDGF/p75 interactions with the SSC. (D) SDS–PAGE analysis of LEDGF/p75 interactions with the SSC. Lane 1, protein markers: MagicMark
XP Western Protein Standard (Invitrogen, Carlsbad, CA, USA); lanes 2–6, the following samples were incubated in the buffer containing 750mM
NaCl and then subjected to size exclusion chromatography as shown in (C): IN alone (lane 2), the purified SSC (lane 3), LEDGF/p75 plus the SSC
(lane 4), LEDGF/p75 alone (lane 5), LEDGF/p75 plus viral DNA (lane 6). IN and LEDGF/p75 were visualized by western blot using respective
antibodies as described in ‘Materials and Methods’ section.
9014 Nucleic Acids Research, 2011,Vol.39, No. 20
mutagenesis studies (63,64) showed that each of the three
surface cysteines could individually be mutated to Ser
without significantly compromising IN catalytic activities
in vitro or viral replication in infected cells. In addition, we
also noted that C56 and C65 are proximal to the viral
DNA binding channel (33), and the placement of
fluorophores at these locations could potentially interfere
with protein–DNA interactions. In contrast, C280 is sig-
nificantly removed from both viral DNA and LEDGF/
p75 binding sites. Therefore, to accomplish the selective
tethering of fluorophores, we mutated C56 and C65 to Ser
and exploited the reactivity of the native C280 residue
sulfhydryl. The data presented in Figure 4A demonstrate
that C280 was indeed specifically targeted by the dye. The
C56/C65S variant was readily labeled by both dyes, while
no reactivity was observed in the case of the triple C56/65/
280S variant. Importantly, the fluorophore labeled
mutant protein used in our FRET studies readily inter-
integration activity (Figure 4C).
Prior to proceeding with FRET measurements, we
monitored time-resolved anisotropies of the D labeled
IN variants, both free and complexed with viral DNA
or LEDGF/p75 using time-correlated single photon
counting. Time-resolved anisotropy curves were well-fit
by a single exponential function. Rotational correlation
times of 2.2, 2.7 and 2.6ns were measured for IN alone,
IN complexed with viral DNA and IN bound to LEDGF/
p75, respectively. These fast rotational correlation times
suggest that the probe retained a significant degree of free
motion upon its tethering to C280. Importantly, very
similar values were observed for IN–LEDGF/p75 and
IN–viral DNA complexes (Supplementary Figure S3),
indicating that neither LEDGF/p75 nor viral DNA sig-
nificantly altered the conformational freedom of the probe
on IN. These control experiments assured us that FRET
values obtained for unliganded IN and its complexes with
LEDGF/p75 and viral DNA can reliably be compared
with one another.
To measure FRET between individual IN subunits in
the context of various complexes, we initially conducted
steady-state (ss) measurements, which reveal average
FRET intensities. In the absence of a binding partner,
the IN dimers mixed together did not exhibit any detect-
able FRET signal (Figure 5A). This was anticipated as at
200nM concentrations of IN(C280-A) and IN(C280-D) in
the reaction mixture, the protein was predominantly a
dimer (Supplementary Figure S1). Furthermore, under
such conditions we did not expect to observe significant
Figure 3. Scheme illustrating design of protein–protein FRET experiments. Two IN proteins are prepared in parallel: one labeled with the D probe
and another with the A probe. The protein concentration range in the reaction mixture is 200–400nM, where free IN is predominantly a dimer. Two
sites (A1/A2 and D1/D2) are labeled in each dimer. Two IN preparations are mixed in ice-cold buffer to minimize the subunit exchange between free
dimeric IN proteins. Subsequent addition of viral DNA or LEDGF/p75 promotes IN tetramerization. Three different populations of IN–viral DNA
or IN–LEDGF/p75 complexes are formed. Of these, only the complex containing D1–D2 and A1–A2 pairs yields FRET. For the cartoon, a
molecular model of full-length HIV-1 IN in complex with LEDGF/IBD was employed.
Nucleic Acids Research, 2011,Vol.39, No. 209015
subunit exchange between two IN preparations for the
following reasons. The dissociation constant for IN
dimers has been reported to be in the subnanomolar
range (59,60). Therefore, effective exchange of individual
subunits between dimeric forms of IN can only be
observed at subnanomolar to lownM protein concentra-
tions (60,65). In contrast, IN forms stable dimers at the
(Supplementary Figure S1). At the same time, this concen-
tration range is low enough to avoid tetramer formation.
IN tetramers and effective exchange of the stable dimers
between tetrameric forms of IN can be detected at ?2mM
protein (41). Therefore, under our assay conditions, the
background FRET due to subunit exchange was very
minimal (Figure 5A).
Addition of LEDGF/p75 promoted formation of a
tetramer by bridging two IN dimers (41) (see also
Supplementary Figure S1) and resulted in significant
energy transfer (Figure 5B). In parallel reactions, viral
DNA was added to the IN(C280-A) and IN(C280-D)
mixture to form the SSC (Figure 5C). While IN in
the context of the SSC is also a tetramer, the IN–viral
DNA complex displayed a significantly higher FRET
than IN–LEDGF/p75 (see overlay of the spectra in
Figure 5D). These results suggest that tetrameric forms
of IN in IN–LEDGF/p75 and IN–viral DNA complexes
We also examined the samples using tr-FRET (Figure 6
and Supplementary Figure S4). We noted that donor only
controls, where IN(C280-D) was mixed with the unlabeled
protein (Figure 6A) and then incubated with LEDGF/p75
(Figure 6B) or viral DNA (Figure 6D), yielded complex
decay curves (Supplementary Figure S5) suggesting that
the fluorophore tethered to IN adopts multiple conform-
ations. Potential asymmetric arrangements of individual
subunits within multimeric IN could contribute to this.
Alternatively, local environment at C280 could allow the
fluorophore to adopt multiple conformations. To delin-
eate between these possibilities, we used the isolated
CTD as a reliable control. Consistent with previous
reports (29,30) isolated CTD formed dimers in our experi-
ments as judged by size exclusion chromatography (data
not shown). Each symmetrical subunit of this protein
fragment contains a single Cys residue at the position
corresponding to C280 in the full-length protein (29,30).
Similarly to full-length IN, the isolated CTD also yielded
three exponential decay curves. These findings suggest that
the local environment at the tethering site contributes to
multiple conformations of the fluorophore. Our results are
reminiscent of the published (66) tr-FRET analysis of
Trp residues in proteins indicating that surface trypto-
phans typicallyadopt different
yield multi-exponential decays curves. In common with
the donor alone, experiments analysis of D–A pairs
(Figure 6 and Supplementary Figure S4) yielded the
decay curves that were best fit to a three exponential
decay (Supplementery Figure
detected when IN(C280-A) was mixed with IN(C280-D)
(Figure 6A).In contrast,
FRET when incubated with LEDGF/p75 or viral DNA
(Figure 6B and D). Average distances calculated from
tr-FRET results were ?81A˚
complex and ?69A˚
distinct IN conformations in these complexes.
these proteins yielded
for the IN–LEDGF/p75
for the SSC (Table 1) indicating
Figure 4. Site-selective labeling of HIV-1 IN with a fluorophore. (A) In parallel experiments, C56/65S and C56/65/280S mutants were subjected to
treatment with Alexa 488 maleimide. The reactions were quenched with DTT and subjected to SDS–PAGE. Images of the same gel following
coomassie staining (upper panel) and UV-light exposure (lower panel) are shown. No fluorescence signal was detected for the C56/65/280S protein
(lane 2), while the C56/65S mutant (lane 1) was effectively labeled with the dye. (B) Assembly of SSCs with the labeled mutant IN. Lane 1, wild-type
IN load; lane 2, load of the IN (C56/65S) mutant; lane 3, protein markers; lane 4, no DNA control; lane 5, the SSC assembly with wild-type IN and
viral DNA; lane 6, the IN (C56/65S) mutant without DNA; lane 7, the SSC assembly with the IN (C56/65S) mutant. (C) Concerted integration
assays of wild-type and mutant (C56/65S) IN proteins: Lane 1: DNA markers, lanes 2 and 3: increasing concentration of the IN (C56/65S) mutant,
lanes 4 and 5: wild-type IN activities.
9016 Nucleic Acids Research, 2011,Vol.39, No. 20
We then extended the tr-FRET experiments to test
more complex interactions involving IN, viral DNA and
LEDGF/p75. The following two pathways for the
assemblyof large nucleoprotein
considered. First, the IN–LEDGF/p75 complex was pre-
formed and viral DNA was then added. The fluorescence
decay profile for this complex was virtually identical to
that for the IN–LEDGF/p75 complex (Figure 6 and
Table 1). In the second set of experiments, we first
obtained the SSC and then exposed it to LEDGF/p75.
The tr-FRET data for this large nucleoprotein complex
was very similar to that of the SSC (Figure 6 and
Table 1). The above FRET experiments were also
obtained with full-length LEDGF/p75 (Figure 6). Taken
together, these data show that the conformation of IN
tetramer depends on the order of ligand addition.
Our FRET results together with available crystallo-
graphic data were employed to generate molecular
models for HIV-1 IN interactions with viral DNA and
LEDGF/IBD (Figure 7 and Supplementary Figure S6).
Figure 7A and B depict two separate conformations of
IN tetramers. The model in Figure 7A was generated
stepwise by first modeling HIV-1 IN interactions with
viral DNA and then docking LEDGF/IBD into the nu-
cleoprotein complex. Interactions between two functional
HIV-1 IN subunits with viral DNA (Figure 7A) were
modeled based on the crystal structure of PFV IN in the
complex with cognate DNA (32) and the resulting nucleo-
protein interactions were similar to those proposed
recently (33). However, published studies (32,33) did not
define NTDs and CTDs in the supporting two subunits.
Our FRET data provided complementary information
and enabled us to model interactions of the full-length
IN tetramer with viral DNA ends (Figure 7A and
Supplementary Figure S6). As shown in Supplementary
Figure S6 (left panels), there are four possible D–A dis-
tances. Of these, distances between D2–A2 and D1–A1
pairs are identical due to the 2-fold symmetry between
two IN dimers, while the distance between D2–A1 pairs
exceeds an effective FRET range (>2?R0) and would not
affect our FRET measurements. Therefore, average
FRET distances are likely to be derived from the following
three distances D1–A2, D2–A2 and D1–A1. In fact,
FRET measurements and molecular modeling results
suggest that these three distances are very similar. For
example, the D1–A2 distance of ?68A˚
agreement with the structure-based modeling using the
PFV intasome as a template. This distance together with
the average FRET distance measurement of ?69A˚for this
complex (Table 1) and the limited length of the loop con-
necting the CCD and CTD, provided us with significant
constraints to position CTDs in supporting subunits as
shown in Figure 7A and Supplementary Figure S6 (left
It should be noted that PFV IN does not interact with
LEDGF/IBD and the published model of HIV-1 intasome
(33) did not address its interactions with the key cellular
cofactor. Our data show that LEDGF/p75 potently binds
the SSC through its biologically relevant site on HIV-1 IN
is in excellent
Figure 5. Ss-FRET plots for IN–viral DNA and IN–LEDGF/p75
complexes. (A) IN alone, (B) IN–LEDGF/p75 complex, (C) IN–viral
DNA complex, (D) overlay of the spectra from experiments shown in
panels A–C. For control experiments, Alexa 488-labeled IN(C56/65S)
was mixed with the unlabeled protein (blue circles in A) and then
incubated with LEDGF/p75 (blue circles in B) or viral DNA (blue
circles in C). The fluorescence intensities for the D–A pairs are
depicted with diamonds and color coded as follows: IN alone,
orange; IN–LEDGF/p75 complex, cyan; IN–viral DNA complex,
magenta. Fluorescence quenching at 520nm and concomitant increase
of emission intensities at 610nm demonstrate FRET. The spectra was
normalizedby defining the maximum
fluorophore in each donor alone experiment as 100%.
intensityof the donor
Nucleic Acids Research, 2011,Vol.39, No. 209017
(Figure 2D) and that the cellular protein does not alter the
pre-formed architecture of the nucleoprotein complex
(Figure 6 and Table 1). In complete agreement with
these experimental data, we were able to dock two
LEDGF/IBD molecules onto the SSC without altering
the pre-existing IN–viral DNA interactions (Figure 7A).
In particular, each LEDGF/IBD engages the CCDs of one
dimer and establishes additional contacts with the NTD
from another dimer, which ensures high-affinity binding
of the cellular cofactor to the complex (41). Taken
together, the experimental results presented here have
allowed us to extend previous modeling of HIV-1 IN–
viral DNA interactions (33) by building a ternary
complex between viral DNA, full-length IN tetramer
and LEDGF/IBD (Figure 7A).
Figure 7B shows an alternative conformation of the IN
tetramer in complex with LEDGF/IBD. To create this
model, we considered the following
Our published biochemical studies (41) have shown that
interactions between the NTD and the CCD are important
for IN tetramerization and high-affinity LEDGF/p75
interactions. In agreement with these findings, subsequent
crystallographic studies (52) with the two domain con-
struct of MVV IN have demonstrated that LEDGF/IBD
bridges between the NTD of one dimer and the CTD of
another dimer. Therefore, these interactions were included
Figure 6. Tr-fluorescence decay plots for IN–viral DNA and IN–LEDGF/p75 complexes. (A) IN alone; (B) IN–LEDGF/p75 complex, (C) IN–
LEDGF/p75 complex was preformed and then exposed to viral DNA; (D) IN–viral DNA complex; (E) IN–viral DNA complex was preformed and
then LEDGF/p75 was added to the nucleoprotein complex. Blue plots show fluorescence decays for donor only control, where Alexa 488-labeled
IN(C56/65S) was mixed with the unlabeled protein. Magenta plots show data for the D–A pairs.
Table 1. Tr-FRET measurements
?DA, nsEr, A˚
?Dis the average excited state lifetime of the donor, ?DAis the average
excited state lifetime of the donor–acceptor pair, E is the average
energy transfer efficiency, r is the average calculated distance. The
number in parenthesis is standard deviation of three independent
9018 Nucleic Acids Research, 2011,Vol.39, No. 20
in our model. Published studies (41,52), however, did not
address the positioning of the CTDs in the protein–protein
complex. Therefore, our FRET data (Table 1) were
employed to create the model between full-length IN
and LEDGF/IBD (Figure 7 and Supplementary Figure
S6). Additional constraints were provided by SAXS data
(56), which revealed the molecular shape and global
dimensions for the full-length HIV-1 IN complex with
distances for D–A pairs are given in Figure 7A and
Supplementary Figure S6 (right panel).
Our FRET studies show that IN tetramers formed in
IN–viral DNA and IN–LEDGF/p75 complexes are
distinct. The order of addition experiments (Figure 6)
further underscore the conclusion that there are different
(Figure 7). The most noticeable difference between these
tetramers is differential positioning of CTDs. In the IN–
viral DNA complex (Figure 7A), the two functional CTDs
tightly interact with the ends of viral DNA and are pos-
itioned immediately adjacent to the CCDs. Our earlier
mass spectrometry-based protein footprinting studies
demonstrated that the CTD undergoes a significant
conformational change upon formation of the IN–viral
DNA complex, while such changes are not observed in
the IN–LEDGF/p75 complex (44). Our FRET results
show that the CTDs are positioned closer to each other
in the presence of viral DNA than LEDGF/p75 (Figures 5
and 6; Table 1). We propose that such conformational
flexibility of the CTDs is crucial for the effective
assembly of the SSC. In contrast, the CTDs in the IN
tetramer formed upon binding to LEDGF/IBD may not
be sufficiently flexible to fully engage viral DNA in a
manner that would ensure formation of the SSC.
For example, bound LEDGF/IBD could indirectly limit
repositioning of the CTD upon subsequent interactions of
the protein–protein complex with viral DNA. At the same
time, it should be noted that preformed IN–LEDGF/IBD
can bind viral DNA to form a less stable nucleoprotein
complex, which effectively catalyzes the 30-processing
reaction but fails to carry out the concerted integration
(41). In line with these observations, both viral and
target DNA can be modeled in the IN–LEDGF/IBD
complex (data not shown). However, the DNA binding
cleft in such a complex would differ from that observed
with the SSC (Figure 7A). In other words, we propose that
there are different modes of viral DNA binding to HIV-1
IN, and only the SSC is capable of productive concerted
The biological relevance of our findings is corroborated
by the following data. PICs isolated from LEDGF/p75
knockout cells exhibit normal levels of DNA strand
transfer activity in vitro, suggesting that IN and viral
DNA can effectively assemble in the cytoplasm of
infected cells in the absence of the cellular cofactor (15).
Furthermore, LEDGF/p75 knockout did not affect
nuclear import of PICs (15). Instead, the cellular
cofactor has been shown to bind PICs at a later stage in
the nucleus and navigate them to active genes on the
chromatin (15,37). While LEDGF/p75 is a predominantly
nuclear protein, it has also been suggested that low
endogenous amounts of the cellular cofactor in the cyto-
plasm could engage PICs (2). Our findings do not preclude
this possibility but rather outline a necessary order of
pre-integration events in which IN engages viral DNA
ends prior to recruitment of LEDGF/p75 into the PIC.
An alternative sequence of events could be detrimental
for HIV-1 integration. For example, overexpression of
LEDGF/IBD that lacked a nuclear localization signal
and therefore interacted with HIV-1 IN in the cytoplasm,
effectively impaired retroviral integration (3). One possible
explanation for this is a potential competition between
LEDGF/IBD and endogenous LEDGF/p75. However,
this mechanism alone cannot explain the findings that
LEDGF/IBD was significantly more effective at suppress-
cells compared with cells containing normal levels of the
cellular cofactor (3). Instead, our FRET results suggest
Figure 7. Molecular modeling of IN, viral DNA and LEDGF/IBD interactions. (A) The assembly of the fully functional nucleoprotein complex.
First, IN interacts with viral DNA to form the SSC. Then, LEDGF/IBD tightly binds the SSC by bridging between the two dimers. Four individual
subunits of IN are colored orange, cyan, magenta and green. The cyan and magenta protomers directly interact with viral DNA, while green and
orange subunits play supporting roles. LEDGF/IBD is depicted in gray. (B) IN interactions with LEDGF/IBD. Colors for IN subunits and LEDGF/
IBD are the same as in A. Locations of C280 in each subunit are shown by red spheres.
Nucleic Acids Research, 2011,Vol.39, No. 209019
that LEDGF/IBD can stabilize an alternative tetrameric
conformation of IN, which is defective in concerted inte-
gration. This notion is further supported by the observa-
tion that overexpressed LEDGF/IBD stabilized the IN
structure in infected cells and protected the retroviral
protein from proteasomal degradation (67). It should
also be noted that IN bound to LEDGF/IBD can still
interact with viral DNA and carry out 30-processing reac-
tions (41). Moreover, LEDGF/IBD is not expected to
interfere with the nuclear import of PICs as LEDGF/
p75 is not involved in this process (15). In contrast, in
line with in vitro observations (41), cell-based assays
have shown that the IN complex with LEDGF/IBD is
defective in concerted integration (3). Taken together,
these studies indicate that modulation of IN structure
prior to its binding to viral DNA is detrimental for
multimerization is highly important for the development
of allosteric inhibitors. We recently proposed (65) a
mechanism for inhibiting HIV-1 IN that would mimic
LEDGF/IBD effects, and reported the discovery of a
small molecule that interacted with K173 at the IN
dimer interface and stabilized an inactive multimeric
conformation of the protein in vitro. More recently,
allosteric inhibitors have been described that impair IN
interactions with LEDGF/p75 in infected cells (68).
These compounds bind at the IN dimer interface, occupy-
ing the LEDGF/IBD binding pocket. In general, detailed
analysis of available structures of the HIV-1 IN CCD
dimer revealedtwo separate
Moreover, several compounds have been reported to
target these sites (69,70). Further studies in this direction
are warranted as the HIV-1 IN multimer is an unexploited
therapeutic target and allosteric drugs binding these sites
are likely to be effective against HIV-1 strains resistant
to current therapies.
Supplementary Data are available at NAR Online.
We are grateful to Dr Christopher McKee for critical
reading of the manuscript and helpful comments.
The National Institutes of Health (grants AI062520,
K.M.-F.); the intramural research program of the
National Institute of Diabetes and Digestive and Kidney
Diseases of the National Institutes of Health and by the
NIH AIDS Targeted Antiviral Program (to R.C.).
Funding for open access charge: National Institutes of
Health (grant AI062520).
Conflict of interest statement. None declared.
1. Brown,P.O. (1997) Integration. In Coffin,J.M., Hughes,S.H. and
Varmus,H.E. (eds), Retroviruses. Cold Spring Harbor Laboratory,
Plainview, NY, pp. 161–204.
2. Llano,M., Vanegas,M., Fregoso,O., Saenz,D., Chung,S., Peretz,M.
and Poeschla,E.M. (2004) LEDGF/p75 determines cellular
trafficking of diverse lentiviral but not murine oncoretroviral
integrase proteins and is a component of functional lentiviral
preintegration complexes. J. Virol., 78, 9524–9537.
3. Llano,M., Saenz,D.T., Meehan,A., Wongthida,P., Peretz,M.,
Walker,W.H., Teo,W. and Poeschla,E.M. (2006) An essential role
for LEDGF/p75 in HIV integration. Science, 314, 461–464.
4. Lewinski,M.K., Yamashita,M., Emerman,M., Ciuffi,A.,
Marshall,H., Crawford,G., Collins,F., Shinn,P., Leipzig,J.,
Hannenhalli,S. et al. (2006) Retroviral DNA integration: viral and
cellular determinants of target-site selection. PLoS Pathog., 2,
5. Farnet,C.M. and Haseltine,W.A. (1990) Integration of human
immunodeficiency virus type 1 DNA in vitro. Proc. Natl Acad.
Sci. USA, 87, 4164–4168.
6. Bukrinsky,M.I., Sharova,N., McDonald,T.L., Pushkarskaya,T.,
Tarpley,W.G. and Stevenson,M. (1993) Association of integrase,
matrix, and reverse transcriptase antigens of human
immunodeficiency virus type 1 with viral nucleic acids following
acute infection. Proc. Natl Acad. Sci. USA, 90, 6125–6129.
7. Buckman,J.S., Bosche,W.J. and Gorelick,R.J. (2003) Human
immunodeficiency virus type 1 nucleocapsid zn(2+) fingers are
required for efficient reverse transcription, initial integration
processes, and protection of newly synthesized viral DNA.
J. Virol., 77, 1469–1480.
8. Carteau,S., Batson,S.C., Poljak,L., Mouscadet,J.F., de
Rocquigny,H., Darlix,J.L., Roques,B.P., Kas,E. and Auclair,C.
(1997) Human immunodeficiency virus type 1 nucleocapsid
protein specifically stimulates Mg2+-dependent DNA integration
in vitro. J. Virol., 71, 6225–6229.
9. Carteau,S., Gorelick,R.J. and Bushman,F.D. (1999) Coupled
integration of human immunodeficiency virus type 1 cDNA ends
by purified integrase in vitro: stimulation by the viral nucleocapsid
protein. J. Virol., 73, 6670–6679.
10. Chen,H. and Engelman,A. (1998) The barrier-to-autointegration
protein is a host factor for HIV type 1 integration.
Proc. Natl Acad. Sci. USA, 95, 15270–15274.
11. Lee,M.S. and Craigie,R. (1994) Protection of retroviral
DNA from autointegration: involvement of a cellular factor.
Proc. Natl Acad. Sci. USA, 91, 9823–9827.
12. Lee,M.S. and Craigie,R. (1998) A previously unidentified
host protein protects retroviral DNA from autointegration.
Proc. Natl Acad. Sci. USA, 95, 1528–1533.
13. Cherepanov,P., Maertens,G., Proost,P., Devreese,B., Van
Beeumen,J., Engelborghs,Y., De Clercq,E. and Debyser,Z. (2003)
HIV-1 integrase forms stable tetramers and associates with
LEDGF/p75 protein in human cells. J. Biol. Chem, 278, 372–381.
14. Cherepanov,P., Sun,Z.Y., Rahman,S., Maertens,G., Wagner,G.
and Engelman,A. (2005) Solution structure of the HIV-1
integrase-binding domain in LEDGF/p75. Nat. Struct. Mol. Biol.,
15. Shun,M.C., Raghavendra,N.K., Vandegraaff,N., Daigle,J.E.,
Hughes,S., Kellam,P., Cherepanov,P. and Engelman,A. (2007)
LEDGF/p75 functions downstream from preintegration complex
formation to effect gene-specific HIV-1 integration. Genes Dev,
16. Engelman,A. (2006) Host cell factors and HIV-1 integration.
Future HIV Ther, 1, 415–426.
17. Li,M., Mizuuchi,M., Burke,T.R. Jr and Craigie,R. (2006)
Retroviral DNA integration: reaction pathway and critical
intermediates. EMBO J, 25, 1295–1304.
18. Li,M. and Craigie,R. (2009) Nucleoprotein complex intermediates
in HIV-1 integration. Methods, 47, 237–242.
19. Andrake,M.D. and Skalka,A.M. (1995) Multimerization
determinants reside in both the catalytic core and C terminus of
avian sarcoma virus integrase. J. Biol. Chem, 270, 29299–29306.
20. Jenkins,T.M., Engelman,A., Ghirlando,R. and Craigie,R. (1996)
A soluble active mutant of HIV-1 integrase: involvement of both
9020 Nucleic Acids Research, 2011,Vol.39, No. 20
the core and carboxyl-terminal domains in multimerization.
J. Biol. Chem., 271, 7712–7718.
21. Chen,J.C., Krucinski,J., Miercke,L.J., Finer-Moore,J.S.,
Tang,A.H., Leavitt,A.D. and Stroud,R.M. (2000) Crystal
structure of the HIV-1 integrase catalytic core and C-terminal
domains: a model for viral DNA binding. Proc. Natl Acad. Sci.
USA, 97, 8233–8238.
22. Wang,J.Y., Ling,H., Yang,W. and Craigie,R. (2001) Structure of
a two-domain fragment of HIV-1 integrase: implications for
domain organization in the intact protein. EMBO J, 20,
23. van Gent,D.C., Elgersma,Y., Bolk,M.W., Vink,C. and
Plasterk,R.H. (1991) DNA binding properties of the integrase
proteins of human immunodeficiency viruses types 1 and 2.
Nucleic Acids Res, 19, 3821–3827.
24. Vincent,K.A., Ellison,V., Chow,S.A. and Brown,P.O. (1993)
Characterization of human immunodeficiency virus type 1
integrase expressed in Escherichia coli and analysis of variants
with amino-terminal mutations. J. Virol., 67, 425–437.
25. Deprez,E., Tauc,P., Leh,H., Mouscadet,J.F., Auclair,C. and
Brochon,J.C. (2000) Oligomeric states of the HIV-1 integrase as
measured by time-resolved fluorescence anisotropy. Biochemistry,
26. Cai,M., Zheng,R., Caffrey,M., Craigie,R., Clore,G.M. and
Gronenborn,A.M. (1997) Solution structure of the N-terminal
zinc binding domain of HIV-1 integrase. Nat. Struct. Biol., 4,
27. Dyda,F., Hickman,A.B., Jenkins,T.M., Engelman,A., Craigie,R.
and Davies,D.R. (1994) Crystal structure of the catalytic domain
of HIV-1 integrase: similarity to other polynucleotidyl
transferases. Science, 266, 1981–1986.
28. Goldgur,Y., Dyda,F., Hickman,A.B., Jenkins,T.M., Craigie,R.
and Davies,D.R. (1998) Three new structures of the core domain
of HIV-1 integrase: an active site that binds magnesium. Proc.
Natl Acad. Sci. USA, 95, 9150–9154.
29. Eijkelenboom,A.P., Lutzke,R.A., Boelens,R., Plasterk,R.H.,
Kaptein,R. and Hard,K. (1995) The DNA-binding domain of
HIV-1 integrase has an SH3-like fold. Nat. Struct. Biol., 2,
30. Lodi,P.J., Ernst,J.A., Kuszewski,J., Hickman,A.B., Engelman,A.,
Craigie,R., Clore,G.M. and Gronenborn,A.M. (1995) Solution
structure of the DNA binding domain of HIV-1 integrase.
Biochemistry, 34, 9826–9833.
31. Kessl,J.J., McKee,C.J., Eidahl,J.O., Shkriabai,N., Katz,A. and
Kvaratskhelia,M. (2009) HIV-1 Integrase-DNA Recognition
Mechanisms. Viruses, 1, 713–736.
32. Hare,S., Gupta,S.S., Valkov,E., Engelman,A. and Cherepanov,P.
(2010) Retroviral intasome assembly and inhibition of DNA
strand transfer. Nature, 464, 232–236.
33. Krishnan,L., Li,X., Naraharisetty,H.L., Hare,S., Cherepanov,P.
and Engelman,A. (2010) Structure-based modeling of the
functional HIV-1 intasome and its inhibition. Proc. Natl Acad.
Sci. USA, 107, 15910–15915.
34. De Rijck,J., Vandekerckhove,L., Gijsbers,R., Hombrouck,A.,
Hendrix,J., Vercammen,J., Engelborghs,Y., Christ,F. and
Debyser,Z. (2006) Overexpression of the lens epithelium-derived
growth factor/p75 integrase binding domain inhibits human
immunodeficiency virus replication. J. Virol, 80, 11498–11509.
35. Vandekerckhove,L., Christ,F., Van Maele,B., De Rijck,J.,
Gijsbers,R., Van den Haute,C., Witvrouw,M. and Debyser,Z.
(2006) Transient and stable knockdown of the integrase cofactor
LEDGF/p75 reveals its role in the replication cycle of human
immunodeficiency virus. J. Virol, 80, 1886–1896.
36. Hombrouck,A., De Rijck,J., Hendrix,J., Vandekerckhove,L.,
Voet,A., De Maeyer,M., Witvrouw,M., Engelborghs,Y., Christ,F.,
Gijsbers,R. et al. (2007) Virus evolution reveals an exclusive role
for LEDGF/p75 in chromosomal tethering of HIV. PLoS Pathog,
37. Ciuffi,A., Llano,M., Poeschla,E., Hoffmann,C., Leipzig,J.,
Shinn,P., Ecker,J.R. and Bushman,F. (2005) A role for LEDGF/
p75 in targeting HIV DNA integration. Nat. Med, 11, 1287–1289.
38. Cherepanov,P., Devroe,E., Silver,P.A. and Engelman,A. (2004)
Identification of an evolutionarily conserved domain in human
lens epithelium-derived growth factor/transcriptional co-activator
p75 (LEDGF/p75) that binds HIV-1 integrase. J. Biol. Chem.,
39. Vanegas,M., Llano,M., Delgado,S., Thompson,D., Peretz,M. and
Poeschla,E. (2005) Identification of the LEDGF/p75 HIV-1
integrase-interaction domain and NLS reveals NLS-independent
chromatin tethering. J. Cell Sci., 118(Pt 8), 1733–1743.
40. Engelman,A. and Cherepanov,P. (2008) The lentiviral integrase
binding protein LEDGF/p75 and HIV-1 replication. PLoS
Pathog, 4, e1000046.
41. McKee,C.J., Kessl,J.J., Shkriabai,N., Dar,M.J., Engelman,A. and
Kvaratskhelia,M. (2008) Dynamic modulation of HIV-1 integrase
structure and function by cellular lens epithelium-derived growth
factor (LEDGF) protein. J. Biol. Chem., 283, 31802–31812.
42. Pandey,K.K., Sinha,S. and Grandgenett,D.P. (2007)
Transcriptional coactivator LEDGF/p75 modulates human
immunodeficiency virus type 1 integrase-mediated concerted
integration. J. Virol., 81, 3969–3979.
43. Raghavendra,N.K. and Engelman,A. (2007) LEDGF/p75
interferes with the formation of synaptic nucleoprotein complexes
that catalyze full-site HIV-1 DNA integration in vitro:
implications for the mechanism of viral cDNA integration.
Virology, 360, 1–5.
44. Zhao,Z., McKee,C.J., Kessl,J.J., Santos,W.L., Daigle,J.E.,
Engelman,A., Verdine,G. and Kvaratskhelia,M. (2008)
Subunit-specific protein footprinting reveals significant structural
rearrangements and a role for N-terminal Lys-14 of HIV-1
Integrase during viral DNA binding. J. Biol. Chem, 283,
45. Cherepanov,P. (2007) LEDGF/p75 interacts with divergent
lentiviral integrases and modulates their enzymatic activity
in vitro. Nucleic Acids Res, 35, 113–124.
46. Maertens,G., Cherepanov,P., Pluymers,W., Busschots,K., De
Clercq,E., Debyser,Z. and Engelborghs,Y. (2003) LEDGF/p75 is
essential for nuclear and chromosomal targeting of HIV-1
integrase in human cells. J. Biol. Chem, 278, 33528–33539.
47. Cherepanov,P., Surratt,D., Toelen,J., Pluymers,W., Griffith,J., De
Clercq,E. and Debyser,Z. (1999) Activity of recombinant HIV-1
integrase on mini-HIV DNA. Nucleic Acids Res, 27, 2202–2210.
48. Kotova,S., Li,M., Dimitriadis,E.K. and Craigie,R. (2010)
Nucleoprotein intermediates in HIV-1 DNA integration visualized
by atomic force microscopy. J. Mol. Biol, 399, 491–500.
49. Zhang,X., Pais,G.C., Svarovskaia,E.S., Marchand,C.,
Johnson,A.A., Karki,R.G., Nicklaus,M.C., Pathak,V.K.,
Pommier,Y. and Burke,T.R. (2003) Azido-containing aryl
beta-diketo acid HIV-1 integrase inhibitors. Bioorg. Med. Chem.
Lett, 13, 1215–1219.
50. Nilsen,B.M., Haugan,I.R., Berg,K., Olsen,L., Brown,P.O. and
Helland,D.E. (1996) Monoclonal antibodies against human
immunodeficiency virus type 1 integrase: epitope mapping and
differential effects on integrase activities in vitro. J. Virol., 70,
51. Cherepanov,P., Ambrosio,A.L., Rahman,S., Ellenberger,T. and
Engelman,A. (2005) Structural basis for the recognition between
HIV-1 integrase and transcriptional coactivator p75. Proc. Natl
Acad. Sci. USA, 102, 17308–17313.
52. Hare,S., Di Nunzio,F., Labeja,A., Wang,J., Engelman,A. and
Cherepanov,P. (2009) Structural basis for functional
tetramerization of lentiviral integrase. PLoS Pathog, 5, e1000515.
53. Arnold,K., Bordoli,L., Kopp,J. and Schwede,T. (2006) The
SWISS-MODEL workspace: a web-based environment for protein
structure homology modelling. Bioinformatics, 22, 195–201.
54. Eswar,N., Webb,B., Marti-Renom,M.A., Madhusudhan,M.S.,
Eramian,D., Shen,M.Y., Pieper,U. and Sali,A. (2006)
Comparative protein structure modeling using Modeller.
Curr. zProtoc. Bioinformatics, Chapter 5, Unit 5 6.
55. Hare,S., Shun,M.C., Gupta,S.S., Valkov,E., Engelman,A. and
Cherepanov,P. (2009) A novel co-crystal structure affords the
design of gain-of-function lentiviral integrase mutants in the
presence of modified PSIP1/LEDGF/p75. PLoS Pathog., 5,
56. Gupta,K., Diamond,T., Hwang,Y., Bushman,F. and Van
Duyne,G.D. (2010) Structural properties of HIV integrase. Lens
epithelium-derived growth factor oligomers. J. Biol. Chem, 285,
Nucleic Acids Research, 2011,Vol.39, No. 20 9021
57. Emsley,P. and Cowtan,K. (2004) Coot: model-building tools for
molecular graphics. Acta Crystallogr. D Biol. Crystallogr., 60,
58. Grobler,J.A., Grobler,J.A., Stillmock,K., Hu,B., Witmer,M.,
Felock,P., Espeseth,A.S., Wolfe,A., Egbertson,M. et al. (2002)
Diketo acid inhibitor mechanism and HIV-1 integrase:
implications for metal binding in the active site of
phosphotransferase enzymes. Proc. Natl Acad. Sci. USA, 99,
59. Tsiang,M., Jones,G.S., Hung,M., Samuel,D., Novikov,N.,
Mukund,S., Brendza,K.M., Niedziela-Majka,A., Jin,D., Liu,X.
et al. (2010) Dithiothreitol causes HIV-1 integrase dimer
dissociation while agents interacting with the integrase dimer
interface promote dimer formation. Biochemistry, 50, 1567–1581.
60. Tsiang,M., Jones,G.S., Hung,M., Mukund,S., Han,B., Liu,X.,
Babaoglu,K., Lansdon,E., Chen,X., Todd,J. et al. (2009) Affinities
between the binding partners of the HIV-1 integrase dimer-lens
epithelium-derived growth factor (IN dimer-LEDGF) complex.
J. Biol. Chem., 284, 33580–33599.
61. Botbol,Y., Raghavendra,N.K., Rahman,S., Engelman,A. and
Lavigne,M. (2008) Chromatinized templates reveal the
requirement for the LEDGF/p75 PWWP domain during HIV-1
integration in vitro. Nucleic Acids Res., 36, 1237–1246.
62. Michel,F., Crucifix,C., Granger,F., Eiler,S., Mouscadet,J.F.,
Korolev,S., Agapkina,J., Ziganshin,R., Gottikh,M., Nazabal,A.
et al. (2009) Structural basis for HIV-1 DNA integration in the
human genome, role of the LEDGF/P75 cofactor. EMBO J, 28,
63. Bischerour,J., Leh,H., Deprez,E., Brochon,J.C. and
Mouscadet,J.F. (2003) Disulfide-linked integrase oligomers
involving C280 residues are formed in vitro and in vivo but are
not essential for human immunodeficiency virus replication.
J. Virol., 77, 135–141.
64. Zhu,K., Dobard,C. and Chow,S.A. (2004) Requirement for
integrase during reverse transcription of human immunodeficiency
virus type 1 and the effect of cysteine mutations of integrase on
its interactions with reverse transcriptase. J. Virol., 78, 5045–5055.
65. Kessl,J.J., Eidahl,J.O., Shkriabai,N., Zhao,Z., McKee,C.J.,
Hess,S., Burke,T.R. Jr, Kvaratskhelia,M. et al. (2009) An
allosteric mechanism for inhibiting HIV-1 integrase with a small
molecule. Mol. Pharmacol., 76, 824–832.
66. Huang,F., Lerner,E., Sato,S., Amir,D., Haas,E. and Fersht,A.
(2009) Time-resolved FRET study shows a compact denatured
state of the B domain of Protein A. Biochemistry, 48, 3468–3476.
67. Llano,M., Delgado,S., Vanegas,M. and Poeschla,E.M. (2004)
Lens epithelium-derived growth factor/p75 prevents
proteasomal degradation of HIV-1 integrase. J. Biol. Chem., 279,
68. Christ,F., Voet,A., Marchand,A., Nicolet,S., Desimmie,B.A.,
Marchand,D., Bardiot,D., Van der Veken,N.J., Van
Remoortel,B., Strelkov,S.V. et al. (2010) Rational design of
small-molecule inhibitors of the LEDGF/p75-integrase interaction
and HIV replication. Nat. Chem. Biol., 6, 442–448.
69. Molteni,V., Greenwald,J., Rhodes,D., Hwang,Y.,
Kwiatkowski,W., Bushman,F.D., Siegel,J.S. and Choe,S. (2001)
Identification of a small-molecule binding site at the dimer
interface of the HIV integrase catalytic domain. Acta Crystallogr.
D Biol. Crystallogr., 57(Pt 4), 536–544.
70. Al-Mawsawi,L.Q., Fikkert,V., Dayam,R., Witvrouw,M.,
Burke,T.R. Jr, Borchers,C.H. and Neamati,N. (2006) Discovery
of a small-molecule HIV-1 integrase inhibitor-binding site. Proc.
Natl Acad. Sci. USA, 103, 10080–10085.
9022 Nucleic Acids Research, 2011,Vol.39, No. 20