impact of template overhang-binding region of hiv-1 rt on the binding and orientation of the duplex...

This article was published in the above mentioned Springer issue.The material, including all portions thereof, is protected by copyright;all rights are held exclusively by Springer Science + Business Media.

The material is for personal use only;commercial use is not permitted.

Unauthorized reproduction, transfer and/or usemay be a violation of criminal as well as civil law.

ISSN 0300-8177, Volume 338, Combined 1-2

Impact of template overhang-binding region of HIV-1 RTon the binding and orientation of the duplex regionof the template-primer

Alok K. Upadhyay • Tanaji T. Talele •

Virendra N. Pandey

Received: 20 August 2009 / Accepted: 29 October 2009 / Published online: 17 November 2009

� Springer Science+Business Media, LLC. 2009

Abstract Fingers domain of HIV-1 RT is one of the

constituents of the dNTP-binding pocket that is involved in

binding of both dNTP and the template-primer. In the

ternary complex of HIV-1 RT, two residues Trp-24 and

Phe-61 located on the b1 and b3, respectively, are seen

interacting with N ? 1 to N ? 3 nucleotides in the tem-

plate overhang. We generated nonconservative and con-

servative mutant derivatives of these residues and

examined their impact on the template-primer binding and

polymerase function of the enzyme. We noted that W24A,

F61A, and F61Y and the double mutant (W24A/F61A)

were significantly affected in their ability to bind template-

primer and also to catalyze the polymerase reaction while

W24F remained unaffected. Using a specially designed

template-primer with photoactivatable bromo-dU base in

the duplex region at the penultimate position to the primer

terminus, we demonstrated that F61A, W24A, F61Y as

well as the double mutant were also affected in their cross-

linking ability with the duplex region of the template-pri-

mer. We also isolated the E–TP covalent complexes of

these mutants and examined their ability to catalyze single

dNTP incorporation onto the immobilized primer terminus.

The E–TP covalent complexes from W24F mutant dis-

played wild-type activity while those from W24A, F61A,

F61Y, and the double mutant (W24A/F61A) were

significantly impaired in their ability to catalyze dNTP

incorporation onto the immobilized primer terminus. This

unusual observation indicated that amino acid residues

involved in the positioning of the template overhang may

also influence the binding and orientation of the duplex

region of the template-primer. Molecular modeling studies

based on our biochemical results suggested that confor-

mation of both W24 and F61 are interdependent on their

interactions with each other, which together are required

for proper positioning of the ?1 template nucleotide in the

binary and ternary complexes.

Keywords HIV-1 reverse transcriptase �Template overhang � HIV-1 RT-DNA binary complex �HIV-1 RT-DNA-dNTP ternary complex �Duplex DNA binding track � HIV-1 RT dimer

Abbreviations

HIV-1 Human immunodeficiency virus

type 1

RT Reverse transcriptase

BSA Bovine serum albumin

DTT Dithiothreitol

dNTP Deoxyribonucleoside

triphosphate

ddNTP Dideoxy-nucleoside

triphosphate

TP Template-primer

U5-PBS RNA template HIV-1 genomic RNA template

corresponding to the primer

binding sequence region

U5-PBS DNA template HIV-1 genomic DNA template

corresponding to the primer

binding sequence region

A. K. Upadhyay � V. N. Pandey (&)

Department of Biochemistry and Molecular Biology,

UMDNJ-New Jersey Medical School, 185 South Orange

Avenue, Newark, NJ 07103, USA

e-mail: [email protected]

T. T. Talele

Department of Pharmaceutical Sciences, College of Pharmacy

and Allied Health Professions, St. John’s University,

8000 Utopia Parkway, Jamaica, NY 11439, USA

123

Mol Cell Biochem (2010) 338:19–33

DOI 10.1007/s11010-009-0316-x Author's personal copy

E–DNA Enzyme bound with DNA

template-primer in the binary

complex

E–DNA–dNTP Enzyme bound with DNA

template-primer and dNTP in

the ternary complex

Introduction

Retroviruses containing a single-stranded (?) RNA genome

cause a variety of diseases, including leukemia, sarcoma,

anemia, and arthritis, as well as immunodeficiency states.

Among these, the human immunodeficiency virus (HIV),

which causes acquired immunodeficiency syndrome

(AIDS) in humans, has been the target of a massive crusade.

A unique feature of their replication is reverse transcription

of their single-stranded RNA genome and its integration as

duplex DNA into the host chromosome [1, 2]. Reverse

transcription is carried out at an early stage after infection

by the virion-encapsidated reverse transcriptase enzyme

(HIV-1 RT). The enzyme is multifunctional, exhibiting

both RNA- and DNA-dependent DNA polymerase activi-

ties, as well as an RNase-H activity that is both exo- and

endonucleolytic [2–5]. These features, combined with the

apparent lack of reverse transcription in the normal

metabolism of eukaryotic cells, have made reverse trans-

criptase an attractive target in the search for chemothera-

peutic agents to combat the spread of retroviral infections.

Human immunodeficiency virus-1 reverse transcriptase

(HIV-1 RT) is a 117-kDa heterodimeric protein (p66/p51);

the smaller p51 subunit is probably generated from p66 of

p66/66 homodimeric enzyme by proteolytic cleavage in

one of the subunits and removal of the COOH-terminal

15-kDa RNase-H domain [2]. In vitro, the enzyme is

functional in both homodimeric (p66/66) and hetero-

dimeric (p66/51) forms with identical kinetic constants [6];

in vivo, however, it exists only as a heterodimeric enzyme

[2]. The catalytic activity of the enzyme resides in the

larger subunit, which is folded into an open structure

containing the polymerase-active site cleft, while the cat-

alytically inactive smaller subunit is closed and compact

[7]. The polymerase cleft is further folded into three dis-

tinct subdomains that resemble the palm, finger, and thumb

of a right hand [8]. Each subdomain has been implicated in

facilitating different functions of HIV-1 RT. The catalysis

of DNA synthesis is performed by highly conserved

aspartate residues in the palm subdomain [8, 9]. The fingers

have been implicated in the binding of incoming dNTP

substrate [10]. The thumb subdomain has been postulated

to participate in translocation of the enzyme along the

template-primer (TP) [11–14]. In the crystal structures of

TP bound HIV-1 RT binary complex [15–17], and TP-

dNTP bound HIV-1 RT ternary complex [18], the position

of the duplex region of the TP appears to be well estab-

lished, while the position and orientation of the single-

stranded template overhang is not well defined.

In the crystal structures of binary and ternary complexes

of different polymerases, the single-stranded region of the

template bends various degrees ‘‘away’’ from the active

site. In the DNA pol I family of enzymes, the first nucleo-

tide in the single-stranded region of the TP is flipped out of

the active site by *90�–180� in both binary and ternary

complexes [19, 20]. In DNA polymerase b, the downstream

duplex region of the TP has an *90� bend at the single-

stranded template overhang of the gapped DNA TP [21]. In

HIV-1 RT, in contrast, the only nucleotide in the overhang

in the binary complex of HIV-1 RT is protruded partially

into the dNTP-binding pocket [16, 17] while, in the ternary

complex, the first unpaired nucleotide is displaced away

from the active site [18]. Also, the bend at the junction of

the duplex region and template overhang is not as sharp as

that in the DNA pol I and DNA pol b family of enzymes.

In the crystal structure of the ternary complex of HIV-1

RT, a number of amino acid residues are seen interacting

with the template overhang [18]. The side chains of two

aromatic amino acids, W24 and F61, are seen interacting

directly with the template overhang. F61 has been impli-

cated in conferring fidelity and sensitivity to nucleoside

analog RT inhibitors [22] and also in strand displacement

activity of the enzyme [23]. While F61 interacts with the

N ? 1 and N ? 2 template nucleotides, the bulky side

chain of W24 is positioned to interact with the N ? 2 and

N ? 3 template nucleotide overhang. However, the inter-

actions of these residues are conspicuously absent in the

crystal structure of enzyme–TP (E–TP) binary complex

HIV-1 RT. Both the residues are far away from the tem-

plate, which contains only a single N ? 1 template

nucleotide overhang. Based on their strategic positions,

both W24 and F61 are expected to be important in binding

the TP in the binary complex, as well as for productive

positioning of the N ? 1 template nucleotide overhang for

the incoming dNTP substrate. Recently, both these residues

have been shown to be involved in accurate binding of TP

to the enzyme [24]. In this study, we generated conserva-

tive and nonconservative mutant derivatives of residues

W24 and F61 and demonstrated that the functions of both

the residues are interdependent on each other. Using a

specially designed photoactivatable self-annealing tem-

plate-primer (SATP), we demonstrated that both W24 and

F61 residues, in spite of being located outside of the duplex

DNA binding tract, significantly influence the productive

binding to the duplex region of TP and positioning of the

N ? 1 template nucleotide in the binary and ternary

complexes.

20 Mol Cell Biochem (2010) 338:19–33

123

Author's personal copy

Materials and methods

Materials

Restriction endonucleases were procured from Promega or

Boehringer Mannheim; DNA sequencing reagents were

from Stratagene (Texas). The HPLC-purified dNTPs were

obtained from Boehringer Mannheim; [32P]-labeled dNTPs

and ATP were obtained from Perkin Elmer Life Science.

All the synthetic oligonucleotides were synthesized at the

Molecular Resource Facility of the University of Medicine

and Dentistry, New Jersey. All other reagents, of the

highest purity grade, were purchased from Fisher, Milli-

pore Corp, and Bio-Rad.

In vitro mutagenesis and protein purification

We used a recombinant clone of HIV-1 RT (pKK223-

RT66) as the template for site-directed mutagenesis. Two

primers corresponding to the sense and antisense strands

containing the desired mutation were used to amplify the

plasmid by high-fidelity PCR using a QuickChangeTM Site-

Directed Mutagenesis Kit (Stratagene). We confirmed the

mutation by sequencing. The wild-type and mutant

enzymes were expressed in E. coli JM109; the homodi-

meric p66 form of enzyme was purified using ion exchange

chromatography as previously described [25, 26].

Thermolysin digestion of wild-type HIV-1 RT

and its mutant derivatives

Five micrograms of the enzyme was treated with 0.03 lg

of thermolysin in a total volume of 10 ll containing

50 mM Tris–HCl (pH 7.5), 2 mM MgCl2, 2 mM CaCl2,

10% glycerol, and 0.2 mM DTT. Reactions were done at

45�C for 30 min and quenched by addition of 5 ll of

protein gel sample buffer (50 mM Tris–HCl, pH 8.5,

100 mM 2-mercaptoethanol, 0.1% SDS, 20% glycerol,

0.01% bromophenol blue) containing 100 mM EDTA.

Samples were resolved on 12% denaturing SDS-poly-

acrylamide gel and visualized by staining with Coomassie

Brilliant Blue.

Preparation of HIV-1 U5-PBS RNA template

We used an HIV-RNA expression clone (pHIV-PBS) for

preparation of the U5-PBS HIV-1 genomic RNA template

as described before [27–29]. We linearized the plasmid

pHIV-PBS with AccI and in vitro transcribed it using T7

RNA polymerase. The enzyme, buffer, and rNTP solutions

were from Ambion. The transcription reaction was done

according to the manufacturer’s protocol.

Polymerase activity assay using primer extension gel

assay

We assayed the DNA- and RNA-dependent DNA poly-

merase activity of wild-type HIV-1 RT and its mutant

derivatives on U5-PBS HIV-1 RNA and U5-PBS DNA

templates primed with 50[32P]-labeled 21-mer DNA PBS

primer. The U5-PBS HIV-1 RNA template was transcribed

from the plasmid pHIV-PBS, which contains a 947-bp

fragment of the HIV-1 genome (?473 to ?1420) corre-

sponding to the PBS region [27]. We did the reaction in a

5 ll volume containing 50 mM Tris–HCl (pH 7.8),

100 lg/ml bovine serum albumin, 2 mM MgCl2, 10 mM

dithiothreitol, 50 mM KCl, 0.5 pmol [32P]-labeled TP

(10 K Cerenkov cpm), 20 lM dNTP, and 7.5 nM enzyme.

We initiated the reaction by adding the enzyme and, after

incubation for 60 s at room temperature, stopped the

reaction by adding an equal volume of gel loading dye

containing 0.025% bromophenol blue, 0.025% xylene

cyanol, 10 mM EDTA, and 40% formamide. We resolved

the reaction products on 8% denaturing polyacrylamide gel

containing 8 M urea and visualized them by phosphorim-

aging analysis.

Photoaffinity cross-linking of enzyme

with the template-primer

We used 50[32P]-labeled 37-mer or 51-mer self-annealing

TPs (SATP) containing photoactivatable bromo-dU base

pairing with the N - 1 template base at the penultimate

position from the 30 primer terminus (Chart 1). We labeled

the TPs at the 50 position using [c-32P] ATP and T4 poly-

nucleotide kinase according to the standard protocol [30].

The labeled TP was separated from free [c-32P] ATP on an

8% polyacrylamide–urea gel. The radioactive band corre-

sponding to the labeled TP was excised, eluted in 0.5 M

ammonium acetate, and purified on an NAP-10 column

(Pharmacia). The cross-linking reaction mixture contained

50 mM Tris–HCl (pH 7.8), 2 mM MgCl2, 1 mM DTT,

50 nM of labeled TP (40 K Cerenkov CPM/pmol), and

512 nM of enzyme in a final volume of 50 ll. The mixture

was exposed to 312 nm UV for 3 min in a Spectrolinker;

the TP-cross-linked enzyme species were resolved by

electrophoresis on SDS-polyacrylamide gel and detected

by phosphorimaging.

Catalytic activity of the E–TP covalent complex

The ability of the cross-linked E–TP covalent complex to

catalyze incorporation of the incoming dNTP onto the

immobilized primer terminus was assessed as described

previously [31, 32]. We incubated 15 pmol of the enzyme

on ice with 25 pmol of unlabeled self-annealing TP

Mol Cell Biochem (2010) 338:19–33 21

123


containing the photoactivatable bromo-dU base, then irra-

diated the E–TP complex at 312 nM UV for 3 min in a

Spectrolinker.

The cross-linked E–TP covalent complexes were puri-

fied as follows: the irradiated samples were loaded on a

DEAE-Sephadex column (0.5 ml) pre-equilibrated with

50 mM Tris–HCl (pH 8.0), 1 mM DTT, 200 mM NaCl,

and 5% glycerol. After extensive washing of the column

with the same buffer to remove the uncross-linked free

enzyme, the E–TP covalent complexes were recovered

from the column by elution with 1.0 M NaCl in the same

buffer. The eluate was desalted and concentrated using

Centriprep-30. The final preparation was free of the

uncross-linked enzyme, as judged by the lack of activity on

the externally added synthetic TP. Incorporation of a single

incoming dNTP was determined by the addition of 10 lCi

of [a32P]-TTP at 0.5 lM concentrations. The reaction, run

for 25 min at room temperature, was terminated by the

addition of 1% SDS and 20 mM EDTA. An aliquot of the

reaction mixture was subjected to SDS-polyacrylamide gel

electrophoresis, then autoradiography.

Gel retardation analysis of E–TP binary complex

and determination of Kd (DNA) value

For gel retardation analysis of E–TP binary complex, an

aliquot of enzyme was incubated with 1 nM of 50[32P]-

labeled TP in an incubation buffer containing 50 mM Tris–

HCl (pH 7.8), 5 mM MgCl2, and 0.01% BSA in a total

volume of 10 ll. After 10 min incubation on ice, we added

to the reaction mixture an equal volume of 29 gel loading

dye containing 0.25% bromophenol blue and 20% glycerol.

We resolved the E–TP binary complexes by electrophore-

sis on a 6% native polyacrylamide gel at 4�C followed by

phosphorimaging.

Using gel retardation analysis, we determined the dis-

sociation constants (Kd) of E–TP binary complexes of the

wild-type HIV-1 RT and of its mutant derivatives, as pre-

viously described [28, 33]. We used a 50[32P]-labeled

21-mer PBS primer annealed with either a 22-mer DNA

template with N ? 1 template overhang or a 26-mer tem-

plate with N ? 5 template overhang. In a total volume of

10 ll, we incubated a fixed concentration of [32P]-labeled

1. U5-PBS HIV-1 RNA containing the primer binding site: 3’-CAG GGA CAA GCC CGC GGU GAC GAU CUC UAA AAG GUG UGA CUG AUU UUC CCA GAC UCC CUA GAG AUC AAU GGU CUC AGU GUG UUG UCU GCC CGU GUG UGA UGA ACU UCC UGA GUU CCG UUC GAA AUA ACU CCG AAU UCG UCA CCC AAG GGA UCA UCG GUC UCU CGA GGG UCC GAG UCU AGA-5’

2. 21-mer DNA PBS primer: 5’-GTCCCTGTTCGGGCGCCACTG-3’

U5-PBS DNA templates corresponding to U5-PBS HIV-1 RNA sequences: 22-mer-3’-CAG GGA CAA GCC CGC GGT GAC G-5’ 23-mer-3’-CAG GGA CAA GCC CGC GGT GAC GA-5’24-mer-3’-CAG GGA CAA GCC CGC GGT GAC GAT-5’ 25-mer-3’-CAG GGA CAA GCC CGC GGT GAC GATC-5’ 26-mer-3’-CAG GGA CAA GCC CGC GGT GAC GATCT-5’

49-mer-3’-CAG GGA CAA GCC CGC GGT GAC GAT CTC TAA AAG GTG TGA CTG ATT TTC C-5’

3. ddC-terminated 21-mer DNA primer and corresponding templates: 5’-GTCCCTGTTCGGGCGCCACTddC-3’ (Primer)

22-mer-3’-CAG GGA CAA GCC CGC GGT GAG G-5’ 26-mer-3’-CAG GGA CAA GCC CGC GGT GAG GATCT-5’

4. Photoactivatable self-annealing template primers

Chart 1 Sequence of template

primers used

22 Mol Cell Biochem (2010) 338:19–33

123


TP (0.3 nM) with increasing concentrations of enzyme in

the incubation buffer. After 10 min of incubation on ice,

we resolved the E–TP binary complexes by native poly-

acrylamide electrophoresis as described. Using Image-

Quant software, we determined the amount of TP bound to

the enzyme in the binary complex and plotted it against

enzyme concentration using Graph Pad software. We cal-

culated the percent of TP bound to the enzyme in the form

of E–TP complex, then fitted it in a nonlinear regression

equation (b = (Vmax * [E])/(Kd ? [E]) as a function of

enzyme concentration. The Kd (DNA) was defined as the

enzyme concentration at which 50% of DNA was bound.

Stable ternary complex formation

We evaluated the ability of HIV-1 RT and its mutant

derivatives to form stable ternary complexes using the

method previously described [34, 35]. Briefly, in a total

volume of 10 ll, we incubated duplicate samples of each

enzyme with 50-[32P]-labeled ddC-terminated 21-mer pri-

mer annealed with a complementary 22-mer or 26-mer

template (0.3 nM). The incubation buffer and conditions

were similar to those used in the binary complex formation.

We first predetermined the amount of each enzyme that

give rise to a complete shift of the labeled TP in the E–TP

binary complex formation. We further incubated the reac-

tion mixture for 10 min in the presence of 200 lM dGTP, a

nucleotide complementary to the first templating base. We

assessed the stability of the ternary complexes by the

degree of labeled TP that remained bound in E–TP–dNTP

complex forms upon the addition of a 1,000-fold excess of

DNA trap. We resolved these complexes on 6% poly-

acrylamide native gel, then did phosphorimaging using

ImageQuant Software.

Glycerol gradient ultracentrifugation

To analyze monomeric and dimeric conformation of the

mutant enzymes, we used glycerol gradient ultracentrifu-

gation analysis [26, 36]. In brief, 50 lg of the enzyme protein

in 50 mM Tris–HCl (pH 8.0), and 400 mM NaCl was

applied on the top of a 5-ml 10–30% linear glycerol gradient

prepared in the same buffer. Gradients were centrifuged at

48,000 rpm in an SW 50.I rotor for 22 h at 4�C and frac-

tionated from the bottom. Sixty-two fractions (80 ll each)

were collected. Protein peaks in the fractions were deter-

mined by OD280 using a Nanodrop spectrophotometer.

CD spectra analysis

We did CD spectra analysis of the wild-type HIV-1 RT and

its mutant derivatives (0.75 lM) in the wavelength range of

195–260 nm at 0.5-nm intervals in an Aviv spectropolarimeter,

model 400 (Lakewood, NJ), using cylindrical fused quartz

cells with a path length of 0.1 cm.

Molecular modeling

The coordinates of 3D structures of HIV-1 RT in E–TP

binary complex and E–TP–dNTP ternary complex were

taken, respectively, from the RCSB PDB 2HMI and 1RTD

files [16, 18]. Due to the lack of template overhang in the

binary structures, we extracted the modeled template

overhang from the binary structure published by Peletskaya

et al. [37], incorporated it into the binary complex crystal

structure of Ding et al. [16], and subjected it to energy

minimization (Macromodel v9.5) to relieve steric hin-

drance between the modeled template overhang and the

protein. To analyze the interaction of mutant derivatives of

W24 and F61, the crystal structure of RT was altered to

include the respective amino acid substitutions. Mutations

were introduced using Maestro v8.0 (Schrodinger, LLC,

New York). Structure display was done using the Maestro

graphical interface. The wild-type and mutant enzyme

complexes were energy-minimized according to a protein

refinement protocol implemented in Macromodel v9.5

(Schrodinger LLC, New York), using the OPLS-AA force-

field parameters [38].

Results

Construction and purification of the mutant enzymes

Five mutants were generated, including a conservative and

a nonconservative mutant at both positions 24 (W24A,

W24F) and 61 (F61A and F61Y), as well as a double

mutant carrying Ala substitutions at both positions (W24A/

F61A). The mutants were expressed in E. coli and purified

to homogeneity with greater than 95% purity. The levels of

their expression, solubility, and electrophoretic profile

were identical to the wild-type enzyme (data not shown).

We also did thermolysin digestion [39] of the mutant

enzymes at 45�C and found no change in their digestion

pattern as compared to the WT enzyme (data not shown)

suggesting that substitution at these positions did not cause

any perturbation in the enzyme structure.

Effect of side chain substitutions at positions 24 and 61

on the polymerase activity of the enzyme

Using both U5-PBS RNA and 49-mer U5-PBS DNA

templates annealed with 21-mer PBS primer, we examined

the effect of conservative and nonconservative substitu-

tions at these positions on the polymerase activity of the

enzyme. Reactions were done for 30 and 60 s, after which

Mol Cell Biochem (2010) 338:19–33 23

123


the reaction products were analyzed on an 8% polyacryl-

amide–urea gel. Nonconservative substitutions at positions

24 and 61 resulted in significant loss of the primer exten-

sion ability of the enzyme (Fig. 1). The W24A mutant was

nearly inactive on the RNA template, while displaying

approximately 50% of wild-type activity on the DNA

template. In contrast, the conservative W24F mutant

exhibited wild-type activity on both the RNA and DNA

templates. Similar patterns of primer extension activity

were obtained with the nonconservative and conservative

mutants of F61. As expected, the primer extension ability

of the double mutant (W24A/F61A) was negligible on both

RNA and DNA templates.

Effect of Ala substitution at positions 61 and 24

on dimer stability of enzyme

Since Gly substitution at position 61 and 24 destabilizes

dimeric conformation of RT [40], we determined whether

Ala substitution at these positions also affects dimeric

conformation leading to reduced TP binding and poly-

merase activity of the enzyme. We used glycerol gradient

ultracentrifugation analysis of the wild-type and mutant

enzymes to determine their dimeric-monomeric conforma-

tions [26, 36]. A dimerization-defective mutant of HIV-1 RT

(W401L) was used as a positive control [41]. Gradients

were fractionated from the bottom and each fraction was

measured for OD280 absorbance. The results indicated that

the sedimentation profiles (35th fraction) of the wild-type

enzyme and all the mutant derivatives of W24 and F61

were identical, having sedimentation peak in the 35th

fraction (Fig. 2). In contrast, the sedimentation peaks of the

dimerization-defective p66W401L mutant of HIV-1 RT and

the p51 subunits were, respectively, in the 39th and 45th

fractions. These results suggest that, unlike Gly substitu-

tion, Ala substitutions at positions 24 and 61 have no

significant impact on the dimer stability of the enzyme. We

also confirmed that Trp ? Ala and Phe ? Ala substitu-

tions at these positions do not alter the secondary structure

of the enzyme since their CD spectra were similar to that of

the wild-type enzyme (Fig. 3).

Effect of template length on the polymerase activity

of the wild-type and its mutant derivatives

In the ternary complex of HIV-1 RT, W24 interacts with

the phosphate backbone of the N ? 2 template nucleotide

and F61 interacts with the base moiety of N ? 2 and with

the sugar ring of the N ? 1 template nucleotides. We,

therefore, postulate that these interactions are crucial for

appropriate positioning of the N ? 1 template for binding

and incorporation of the incoming dNTP substrate. Any

nonconservative substitutions at positions 24 and 61 may

significantly impair the enzyme’s ability to position the

N ? 1 template, thus affecting the polymerase function of

the enzyme. To ascertain this, we examined the polymerase

activity of these mutants on TPs with varying template

overhangs. We used 50-[32P] labeled 21-mer PBS primer

annealed with U5-PBS DNA templates carrying N ? 1 to

N ? 5 nucleotide overhangs (Chart 1). The gel extension

profile of the polymerase products obtained with these

mutants is shown in Fig. 4.

The primer extension ability of the conservative W24F

mutant was similar to that of the wild-type enzyme on all

the template overhangs. The nonconservative W24A

mutant was significantly affected in its primer extension

ability on the templates with a N ? 1 nucleotide overhang,

but had approximately 50% of wild-type activity on tem-

plates with N ? 2 to N ? 5 template nucleotide overhangs.

In contrast, the primer extension ability of F61A and

double mutant W24A/F61A was severely affected on all

template overhangs. The conservative F61Y mutant was

also impaired with a N ? 1 template overhang and was

unable to incorporate the last nucleotide on the other two

templates (N ? 2 and N ? 3) when it encountered a situ-

ation similar to a N ? 1 template. However, the F61Y

mutant synthesized full-length products on templates with

N ? 4 and N ? 5 nucleotide overhangs, probably due to

additional interactions with the longer template overhangs.

The impaired ability of F61A and F61Y to extend the

primer with a N ? 1 template suggests that the Phe side

chain at the 61 position may be required for proper

Fig. 1 Polymerase activity of the wild-type enzyme and its mutant

derivatives on RNA and DNA templates. The polymerase activity of

the wild-type and mutant enzymes was determined on U5-PBS RNA

and U5-PBS DNA templates primed with 50[32P]-labeled 21-mer PBS

primer; a primer extension gel assay was done. Lanes 1 and 2

represent the products formed during the reaction, which was carried

out for 30 and 60 s, respectively. The DM indicates double mutant

(W24A/F61A) while P indicate the position of the labeled primer

24 Mol Cell Biochem (2010) 338:19–33

123


positioning of a N ? 1 template. However, the impaired

ability of W24A on a template with a N ? 1 template was

unexpected, since this mutant had a wild-type Phe residue at

position 61. It is possible that the side chain conformations

of both F61 and W24 are dependent on their interaction

with each other, which may be lost by Ala substitution at

either of the positions.

Photoaffinity cross-linking of mutant enzymes

with the template-primer

The impaired or reduced ability of mutant derivatives of

F61 and W24 to extend the primer on short template

overhangs may be due to either reduced DNA binding or

improper positioning of the template overhang. We exam-

ined the DNA binding function of these mutant enzymes by

covalent cross-linking of the 50[32P]-labeled self-annealing

37-mer TP containing a photoactivatable BrdU base at the

penultimate position from the primer terminus pairing with

N - 1 template in the duplex region. Since 30-OH of the

primer terminus is positioned in the catalytic cleft, the BrdU

base pairing with a N - 1 template is expected to cross-link

selectively with the interacting residue in the catalytic cleft

upon irradiation of the E–TP complex at 312 nm. At this

Fig. 3 CD spectrum of the wild-type HIV-RT and its mutant

derivatives of W24 and F61. CD spectrum analysis was carried out

in the wavelength range of 195–260 nm at 0.5-nm intervals in an

Aviv spectropolarimeter, model 400 (Lakewood, NJ) using cylindri-

cal fused quartz cells with a path length of 0.1 cm

Fig. 2 Glycerol gradient ultracentrifugation analysis of the WT HIV-1

RT and its mutant derivatives of W24 and F61. The wild-type p66/66

HIV-1 RT and its mutant derivatives were individually resolved on

10–30% linear glycerol gradient ultracentrifugation at 48,000 rpm in

an SW 50.I rotor for 22 h at 4�C. Gradients were fractionated from

the bottom and measured for OD280 absorbance using a Nanodrop

spectrophotometer. We also included p51 monomer and a dimeriza-

tion-defective W401L mutant as a positive control

b

Mol Cell Biochem (2010) 338:19–33 25

123


wavelength, only the BrdU base can be activated to cross-

link with the enzyme. We have earlier shown that this TP

selectively crosslinks to the catalytic subunit upon irradia-

tion at 312 nm UV [31]. The cross-linked covalent complex

was resolved on SDS-PAGE. All the mutant enzymes

except the double mutant were able to form a covalent

complex with the 50-[32P]-labeled self-annealing 37-mer

TP, although the extent of the E–TP complexes formed by

nonconservative W24A and F61A mutants was significantly

lower than that produced by the wild-type enzyme

(Fig. 5A). Among these, the double mutant (W24A/F61A)

was severely affected in its ability to form E–TP binary

complex. Since 37-mer SATP had only a 12 bp duplex

region, we also used a 51-mer self-annealing TP with a

19 bp duplex region to rule out the possibility that the

variations in cross-linking may have been due to a shorter

duplex region. Although the extent of cross-linking with

51-mer SATP was marginally improved, the overall pattern

remained identical to 37-mer SATP (Fig. 5B). The two

bands that are seen in the gel are common with these tem-

plat- primers, possibly existing as a mixture of monomer

and dimer due to the presence of complementary sequences.

Ability of mutant enzymes to incorporate a single

incoming dNTP onto the immobilized primer terminus

We examined whether the N ? 1 template nucleotide in

these E–TP covalent complexes is appropriately positioned

to facilitate incorporation of the first incoming dNTP

substrate onto the primer terminus of the immobilized TP.

We incubated the wild-type and mutant enzymes with

unlabeled 37-mer or 51-mer SATP containing a photoac-

tivatable BrdU base pairing with the N - 1 template base

in the duplex region (Chart 1). The E–TP complexes were

then UV irradiated at 312 nm and the E–TP covalent

complex formed was purified. We supplemented the puri-

fied E–TP covalent complexes with [32P]-labeled TTP and

examined their ability to incorporate the first incoming

dNTP substrate onto the immobilized primer terminus.

The results indicated differences in the ability of the

E–TP covalent complexes of the wild-type and mutant

derivatives to incorporate the first incoming dNTP sub-

strate (Fig. 6). While E–TP covalent complexes from

W24F displayed wild-type activity, the complexes from

F61Y, W24A, and F61A mutants were significantly

Fig. 4 DNA polymerase activity of the wild-type enzyme and its

mutant derivatives on template-primers with varying template over-

hangs. Primer extension gel assay was done using the 50-[32P]-labeled

21-mer PBS primer annealed with DNA templates carrying N ? 1,

N ? 2, N ? 3, N ? 4, or N ? 5 template nucleotide 50 overhangs.

The reaction was done for 60 s in a total volume of 5 ll and the

products resolved on denaturing polyacrylamide-urea gel. The

number with (N?) sign below each lane indicates the number of

nucleotides in the 50 template overhang

Fig. 5 Photoaffinity cross-linking of 50-[32P]-labeled A 37-mer and

B 51-mer self-annealing template-primer to the wild-type enzyme and

its mutant derivatives. The 50[32P]-labeled SATP contained photoac-

tivatable bromo-dU base that base pair with the N - 1 template base

at the penultimate nucleotide from the 30 primer terminus (Chart 1).

The labeled SATP (50 nM, 100K Cerenkov cpm) was incubated with

512 nM of each enzyme in a solution containing 50 mM Tris–HCl

(pH 7.8), 2 mM MgCl2, and 1 mM DTT in a final volume of 50 ll.

The mixture was exposed to 312 nm UV for 3 min in a Spectrolinker.

The TP-cross-linked enzyme species were resolved by electrophoresis

on SDS-polyacrylamide gel and detected by phosphorimaging. The

two bands seen in the gel are common with self-annealing TPs since,

due to complementary sequences, they exist as both monomer and

primer-dimer. The control lane represents 50-[32P] labeled SATP

irradiated without the enzyme which has ran out of the gel upon SDS-

PAGE

26 Mol Cell Biochem (2010) 338:19–33

123


impaired in their ability to incorporate the first incoming

dNTP onto the immobilized primer terminus of the cross-

linked TP. As expected, only a trace of nucleotidyltrans-

ferase activity could be detected with E–TP covalent

complexes from the double mutant (W24A/F61A). This is

consistent with its low level of E–TP binary complex for-

mation (Fig. 6A, B). These results support the contention

that W24 and F61 residues are involved not only in binary

complex formation, but also in appropriate positioning of

the N ? 1 template nucleotide for dNTP-binding and

polymerase function of the enzyme.

Effect of template overhang on the DNA binding

affinity of mutant enzymes

To determine whether TP binding affinity of mutant

enzymes is influenced by template nucleotide overhangs,

we first examined the TP binding pattern of the enzymes,

using gel retardation with 22/21 and 26/21-mer TP

carrying, respectively, N ? 1 and N ? 5 template nucleo-

tide overhangs. We incubated each mutant and WT enzyme

at a 1 nM concentration with 1 nM of [32P]-labeled TP. As

shown in Fig. 7, the extent of E–TP binary complex for-

mation by the nonconservative W24A and F61A mutants

depended on the length of the template overhang. Both

mutants were unable to form the E–TP binary complex

when a template had only a single nucleotide overhang.

However, when a template overhang was increased to

N ? 5 nucleotides, the ability of both W24A and F61A

mutants to form the binary complex was slightly improved,

but remained at a significantly lower level than did for-

mation by the wild-type enzyme. As expected, the double

mutant (W24A/F61A) was unable to form detectable bin-

ary complex with both the TPs. In contrast, the extent of

E–TP binary complex formation with conservative W24F

and F61Y mutants was similar to the wild-type enzyme

with both shorter and longer template overhangs.

After qualitatively evaluating E–TP binary complex

formed by these mutants, we determined the equilibrium

dissociation constant (Kd) of E–TP binary complexes for

the wild-type enzyme and its mutant derivatives on the two

TPs carrying N ? 1 and N ? 5 nucleotide template over-

hangs. The gel retardation analyses of E–TP complexes of

the wild-type and its mutant derivatives are shown in

Fig. 8. The values for Kd determined (Table 1) demonstrate

a strong correlation between the polymerase activity of the

mutants and their Kd values. The mutant enzymes W24A

and F61A, which had reduced enzyme activity on all five

Fig. 6 Catalytic activity of the E–TP covalent complex. We incu-

bated 15 pmol of enzyme with 25 pmol of unlabeled cold A 37-mer

SATP or B 51-mer SATP and irradiated the mixture at 312 nm UV

for 3 min. The ability of E–TP covalent complex to catalyze the

incorporation of nucleotide onto the immobilized primer terminus

was examined by incubating the complex with 10 lCi of the first

incoming [a32P]-TTP at a 0.5 lM concentration. The reaction mixture

was incubated for 25 min at room temperature. The reaction was

terminated by the addition of 1% SDS and 20 mM EDTA. An aliquot

of the reaction mixture was subjected to SDS-polyacrylamide gel

electrophoresis, then autoradiography. The control lane represents

incubation of uncrosslinked wild-type enzyme–SATP complex with

[a32P]-TTP and the extended SATP ran out of the gel upon SDS-

PAGE

Fig. 7 Effect of 50 template nucleotide overhang of the template-

primer on binding with the enzyme. One nanomolar of the wild-type

HIV-1 RT or its mutant derivatives was incubated with 1 nM of

50-[32P]-labeled 22/21-mer TP with an N ? 1 template overhang or

26/21 TP with an N ? 5 template overhang. Following 10 min

incubation on ice in an incubation buffer of 50 mM Tris–HCl

(pH 7.8), 5 mM MgCl2, and 0.01% BSA, the E–TP binary complex

formed was analyzed by nondenaturing polyacrylamide gel

electrophoresis

Mol Cell Biochem (2010) 338:19–33 27

123


TPs, also exhibited 10–15-fold lower DNA binding affinity

than did the wild-type enzyme. The DNA binding affinity

of the double mutant was reduced by approximately 125-

fold. As expected, the conservative W24F mutant with

wild-type polymerase activity displayed no change in its

DNA binding affinity.

Stable ternary complex formation

In the crystal structure of the ternary complex of HIV-1 RT

(E–DNA–dNTP), a significant movement of the fingers

subdomain toward the polymerase cleft has been noted

upon binding of dNTP to the E–TP binary complex [18].

Specifically, the fingers subdomain moves 20 A toward the

palm subdomain (fingers closing) so that, following dNTP

binding, the TP in the E–TP binary complex is locked in a

stable ternary complex poised for catalysis. An in vitro

assay for ternary complex formation using dideoxy termi-

nated primer annealed with the template allows the next

correct dNTP to bind in the ternary complex without

actually being incorporated [34]. We used this assay to

examine the ability of these mutant enzymes to transform

E–TP binary complexes carrying (N ? 1) and (N ? 5)

template nucleotide overhangs into stable E–TP–dNTP

ternary complexes in the presence of first incoming dNTP

substrate. Since the binding of dNTP to the enzyme is an

ordered mechanism that occurs only after the formation of

E–TP binary complex, the extent of labeled TP remaining

bound to the enzyme in the presence of dNTP and a large

excess DNA trap represents the extent of the stable ternary

complex formed.

We determined the amount of DNA trap required for

complete dissociation of labeled E–TP binary complex and

the extent of stable ternary complex formed by the wild-

type enzyme (Fig. 9A, B). We then incubated the preformed

E–TP binary complexes of the WT and mutant enzymes in

the presence of a 1,000-fold molar excess of DNA trap

(Fig. 9C, lane 2) or in the presence the correct incoming

dNTP (dGTP; 200 lM), followed by addition of the DNA

trap (Fig. 9C, lane 4). The E–TP binary complexes of all the

mutants and WT enzymes were completely dissociated in

the presence of a large excess of DNA trap (Fig. 9C, lane 2).

In contrast, a significant amount of E–TP binary complexes

were converted to E–TP–dNTP ternary complexes, which

remained resistant to competition with the DNA trap

(Fig. 9C, lane 4). The nonconservative W24A and F61A

mutants and the conservative F61Y mutant were unable to

form stable ternary complexes when TP carried a single

N ? 1 template nucleotide overhang (22/21-TP, lane 4).

Only a trace amount of the ternary complex could be formed

when the template overhang was increased to N ? 5

Fig. 8 Determination of the equilibrium dissociation constant of

E–TP binary complex (Kd [TP]) with TP carrying ?1 and ?5 template

nucleotide overhangs. The 50-[32P]-labeled 21-mer primer was

annealed with complementary 22-mer and 26-mer templates carrying

50 template overhangs of ?1 and ?5 nucleotides, respectively. The

labeled TP was incubated with varying concentrations of the enzyme.

The E–TP binary complexes were resolved by electrophoresis on 6%

native nondenaturing polyacrylamide gel. The amount of labeled TP

bound to the enzyme in the binary complex was determined and

plotted against the enzyme concentration. The Kd was defined as the

concentration of enzyme at which 50% of DNA was bound

Table 1 Kd of the wild-type enzyme and mutant enzymes on TP with

different template overhangsa

Enzyme Kd (nM)

22/21-mer

(N ? 1 overhang)

26/21-mer

(N ? 5 overhang)

WT 0.24 ± 0.12 0.25 ± 0.03

W24A 3.50 ± 0.14 2.0 ± 0.21

W24F 0.30 ± 0.03 0.30 ± 0.01

F61A 4.0 ± 0.08 2.5 ± 0.08

F61Y 1.60 ± 0.15 0.56 ± 0.12

DM (W24A/F61A) 30 ± 7.5 30 ± 9.5

a The data represent the average ± standard deviation of three

experiments

28 Mol Cell Biochem (2010) 338:19–33

123


nucleotides (26/21-TP; lane 4). In contrast, the conservative

W24F mutant was as efficient as the wild-type enzyme in

the ability to transform its E–TP binary complexes into

stable ternary complexes with TP carrying N ? 1 or N ? 5

template overhang. As expected, double mutant (W24A/

F61A) was unable to form stable ternary complex with

either of the two TPs.

Discussion

The crystal structure of several DNA polymerases in the

form of E–TP binary and E–TP–dNTP ternary complexes

reveals that template overhangs in the binary and ternary

complexes do not assume an ordered structure. In poly-

merase b, the template overhang is essentially bent [21],

while in the pol I family of DNA polymerases the template

overhang is flipped away from the active site in both binary

and ternary complexes [20, 42]. The observed bending or

flipping in the template strand, which shifts the next tem-

plate nucleotide away from the active site, may be an

important mechanism to avoid deletion mutagenesis by

discouraging incorrect template base reading [43]. In HIV-

1 RT, the position of the duplex region of the TP is similar

in both the binary and ternary complexes, while the posi-

tion of the single-stranded overhang is significantly dif-

ferent [15, 16, 18].

The structural data on the E–TP binary complex of HIV-

1 RT do not provide information regarding the position of

the template overhang. All of the three reported crystal

structures of the E–TP binary complexes of HIV-1 RT have

a single (?1) template nucleotide overhang [15–17], which

is distorted and partially protruded into the dNTP-binding

pocket. In the ternary complex, the first unpaired nucleo-

tide is displaced away from the active site [18]. Therefore,

the position of the 50-template overhang in the HIV-1 RT

binary complex is not clearly known. Also, the regions of

HIV-1 RT that interact with the single-stranded 50-over-

hang have not been explored. The position of the template

overhang can be only partially inferred from the ternary

complex crystal structure of HIV-1 RT [18]. Using chem-

ical and photo-cross-linking techniques, it has been shown

that in the binary complex of HIV-1 RT the template

overhang interaction sites lie in the finger subdomain 5–7

nucleotides beyond the polymerase-active site [37].

Analysis of the ternary complex of HIV-1 RT [18]

showed that a number of amino acid residues interact with

the template overhang. The N ? 1 template nucleotide is

lodged underneath the side chain of L74 and alongside the

peptide backbone of G152, which seems to facilitate

trapping of the template strand in the closed ternary com-

plex [18]. The N ? 1 template nucleotide also has multiple

interactions with the surrounding amino acid residues. Its

phosphate backbone and base moiety interact with R78 and

F61, respectively, while sugar moiety is stabilized by side

chain of D76 and peptide backbone of V75 [44]. These

residues significantly influence the fidelity of DNA

Fig. 9 A, B Effect of DNA trap concentration on the formation of

E–TP binary and E–TP–dNTP ternary complexes. The E–TP binary

complex and ternary complex E–TP–dNTP of the WT enzyme were

formed by incubating 5 nM of the wild-type enzyme with 0.3 nM of

radiolabeled ddC-terminated TP (22/dd21-mer) at 4�C for 10 min in

the absence of dNTP (panel A) and in the presence of 200 lM dGTP

(panel B). The individual binary and ternary complexes were then

competed out by the addition of increasing concentrations of

unlabeled 22/dd21-mer as the DNA trap from 1 to 300 nM. The

trap-susceptible binary complex and trap-resistant stable E–TP–dNTP

ternary complexes were analyzed on a nondenaturing polyacrylamide

gel. C Analysis of the E–DNA–dNTP ternary complex formed by

wild-type HIV-1 RT and its mutant derivatives on TP carrying ?1

and ?5 template nucleotide overhangs. The E–TP binary complex

with each of the mutant enzymes was formed by incubating with

0.3 nM of radiolabeled ddC-terminated TP carrying a ?1 (22/dd21-

mer) or ?5 (26/dd21-mer) template nucleotide overhang (lane 1). The

concentration of each mutant enzyme required to achieve complete

shift was determined by titration. Addition of a DNA trap at a 300-nM

concentration completely competed out the labeled template-primer

(lane 2). The ternary complex was formed by supplementing the

E–TP binary complex with 200 lM dGTP (lane 3). The individual

ternary complexes were then competed out by the addition of

increasing concentrations of unlabeled TP (lane 4)

Mol Cell Biochem (2010) 338:19–33 29

123


synthesis of the enzyme [22, 23, 45, 46]. Substitutions of

Arg ? Ala and Asp ? Val at positions 78 and 76,

respectively, were also shown to affect the DNA binding

affinity of the enzyme; while R78A decreased the TP

binding affinity by 8-fold, D76V mutation enhanced it by

7-fold [45]. Another important residue involved in stabiliz-

ing the template overhang in the ternary complex is W24,

which interacts with the phosphate backbone between N ? 2

and N ? 3 nucleotides, as well as their sugar moieties, in

coordination with F61, which interacts with the sugar moi-

eties of N ? 1 and N ? 2 and also with the N ? 1 base.

We noted that Ala substitutions at either position 61 or

24 have adverse effects on the binary complex formation of

the enzyme. The double mutant (W24A/F61A) was most

severely affected in its ability to form the binary complex

(Fig. 5A). For this purpose, we used specifically designed

self-annealing TPs that had bromo-dU at the penultimate

position from the primer terminus pairing with the N - 1

template base. Upon irradiation of the E–TP complex at

312 nm UV, only bromo-dU was excited to crosslink with

the enzyme. This result was intriguing since both W24 and

F61 are in the vicinity of the single-stranded template

overhang, while the cross-linking site (BrdU base) is on the

primer strand in the duplex region. In the crystal structure

of binary complex, the side chain of Y183 is close to the

penultimate nucleotide base from the primer terminus,

which may crosslink with the BrdU base upon 312 nm UV

irradiation. This contention is supported by the fact that

Y183A mutant does not crosslink with the BrdU containing

SATPs when exposed to 312 nm UV (data not shown). The

reduced crosslinking observed with W24A, F61A, and the

double mutant enzymes suggests possible alteration in the

positioning of the duplex region of TP in their binary

complexes. It is possible that misalignment of the N ? 1

template overhang also influences positioning of the primer

terminus in the duplex region of the TP.

This contention was supported by the fact that ability of

E–TP covalent complexes of the mutant enzymes to

incorporate a single incoming [a-32P]-dNTP onto the

immobilized primer terminus was significantly reduced as

compared to the wild-type enzyme (Fig. 6) suggesting

improper positioning of the primer terminus and/or the

N ? 1 templating nucleotide. Involvement of both W24

and F61 in productive positioning of the N ? 1 template

overhang was further supported by the fact that their

mutant derivatives, except W24F, had severely limited

ability to catalyze dNTP incorporation on TP with a single

N ? 1 template nucleotide overhang (Fig. 4). The poly-

merase activity of all but one of these mutants was

improved upon increase in the template overhang. The

exception was the double mutant (W24A/F61A), which did

not use these TPs, thereby suggesting significant destabi-

lization of the template position.

We further noted that binary complex formation with

these mutant derivatives is influenced by the length of the

template overhang. A 14–16-fold increase in the Kd (DNA)

for both the W24A and F61A mutants occurred with a

N ? 1 template overhang, in contrast to the 8–10 fold

increase when the template overhang was increased to ?5

(Table 1). As expected, the decrease in the TP binding

affinity of the double mutant (W24A/F61A) was greater

than 125-fold with either an N ? 1 or N ? 5 template

overhang.

In a study in which both W24 and F61 were substituted

with Gly, the double mutant displayed lowest binding

affinity for the TP and about a reduction of 80% in the

steady-state rate of RT activity [24]. These authors have

suggested that Gly substitution at positions 24 and 61

destabilizes the dimeric conformation of the enzyme,

which may have affected the DNA binding function of

the enzyme [40]. Unlike Gly substitutions, we found that

Ala substitutions at these positions have no effect on

dimerization of the enzyme. The observed reduction in

the DNA binding affinity could be due to loss of inter-

action with the N ? 1 template nucleotide. A similar

decrease in DNA binding affinity was noted for a non-

conservative mutant derivative of R78 (R78A), which

interacts with the phosphate backbone of an N ? 1 tem-

plate nucleotide [45]. These results suggest that amino

acid residues interacting with N ? 1 template nucleotide

also affect binding of the duplex region of the TP.

Interaction of both W24 and F61 with the template

overhang is required for binary complex formation. Loss

of this interaction due to Ala substitution at either or both

positions greatly affected the formation of stable ternary

complexes, suggesting a possible role of these residues in

productive positioning of the N ? 1 template nucleotide

in the dNTP-binding pocket.

Vertical scanning mutagenesis at position 61 has shown

that among the conservative and nonconservative mutants,

F61A displayed the highest fidelity and large reductions in

sensitivity to ddNTPs [22]. A similar increase in fidelity

has been reported for W24G mutant [24]. The enhanced

fidelity of F61A and W24G mutants could be due to

improper positioning of the N ? 1 template nucleotide in

their binary complexes causing restraint on the flexible

conformation of the dNTP-binding pocket. Also, because

Ala substitution at either position affects the function of the

other, it is apparent that the side chains at positions 24 and

61 may be interdependent with regard to the stability of

their side chain conformation. In addition to W24 and F61,

other residues such as R78, D76, and V75 interacting with

N ? 1 template may be involved in facilitating the posi-

tioning of the templating base, since mutations at these

positions significantly affect the DNA binding affinity and

fidelity of DNA synthesis.

30 Mol Cell Biochem (2010) 338:19–33

123


To examine the interaction of these residues at the

structural level, we used the binary complex crystal

structure [16] with modeled extended template overhang

[37] and the ternary complex structure with an N ? 3

template overhang [18]. In the binary complex, the fingers

are in the open conformation, giving the extended template

the opportunity to interact, respectively, with W24 and F61

in the b1 and b3 sheets.

In the binary complex, the solvent-exposed W24 is away

from the template strand but within interacting distance

with F61, establishing an edge-to-face contact via its indole

moiety and the phenyl ring of F61 (Fig. 10A, WT). A

conformational search of the F61 side chain, followed by

energy minimization of the resulting E–TP complex,

showed that in one of the allowed conformations, it inter-

acts with the ?1 template nucleotide and exhibits inter-

action with the side chain of W24. These interactions seem

to be essential for proper conformation of the side chains,

which together facilitate positioning of the N ? 1 template

overhang. Ala substitution at position 24 abolishes these

interactions and destabilizes the side chain conformation of

F61, resulting in loss of its key interaction with the N ? 1

template nucleotide (Fig. 10A, W24A). This contention is

supported by the observed inability of both W24A and

F61A mutants to form binary complex and stable ternary

complex with TP carrying a single N ? 1 template nucle-

otide overhang (Fig. 10A, F61A).

In the ternary complex, both W24 and F61 are solvent-

exposed and close to the template overhang (Fig. 10B,

WT). The W24 interacts with N ? 2 and N ? 3 template

nucleotides, whereas F61 establishes interaction with

N ? 1 and N ? 2 template nucleotides. As in the binary

complex, the edge-to-face interactions between aromatic

side chains of W24 and F61 are maintained in the ternary

complex but lost upon Ala substitution at either of these

positions. Taken together, our biochemical data and

molecular modeling study imply that aromatic amino acid

residues at positions 24 and 61 have important functions in

the formation of binary complex and positioning of the

N ? 1 templating base to facilitate the transition of binary

complex into productive, stable ternary complex.

Acknowledgment This research was partly supported by grants

from the NIAID/NIH (AI074477 and AI42520 to VNP).

References

1. Sarafianos SG, Marchand B, Das K, Himmel DM, Parniak MA,

Hughes SH, Arnold E (2009) Structure and function of HIV-1

reverse transcriptase: molecular mechanisms of polymerization

and inhibition. J Mol Biol 385:693–713

2. Telesnitsky A, Goff SP (1997) Reverse transcriptase and gener-

ation of retroviral DNA. In: Coffin JH, Hughes SH, Varmus HE

(eds) Retroviruses. Cold Spring Harbor Laboratory Press, NY

Fig. 10 Interactions between W24 and F61 and the DNA template

overhang. Molecular models of binary and ternary complexes of wild-

type and mutant derivatives of RT. The figure was created on Maestro

8.0 using PDB coordinates of 2HMI and 1RTD. Color scheme:

template is shown in orange, the primer in cyan, amino acid residues

as green ball-and-stick, the incoming dTTP substrate in color by

element as ball-and-stick, and Mg?2 ions in light pink. The positions

of the templating base and the downstream template nucleotides are

labeled ?1, ?2, ?3, and ?4 in the binary complex; template

overhangs in the ternary complex are labeled ?1, ?2, and ?3

Mol Cell Biochem (2010) 338:19–33 31

123


3. Basavapathruni A, Anderson KS (2007) Reverse transcription of

the HIV-1 pandemic. FASEB J 21:3795–3808

4. Katz RA, Skalka AM (1994) The retroviral enzymes. Annu Rev

Biochem 63:133–173

5. Schatz O, Mous J, Le Grice SF (1990) HIV-1 RT-associated

ribonuclease H displays both endonuclease and 30–50 exonuclease

activity. EMBO J 9:1171–1176

6. Beard WA, Wilson SH (1993) Kinetic analysis of template-pri-

mer interactions with recombinant forms of HIV-1 reverse

transcriptase. Biochemistry 32:9745–9753

7. Wang J, Smerdon SJ, Jager J, Kohlstaedt LA, Rice PA, Friedman

JM, Steitz TA (1994) Structural basis of asymmetry in the human

immunodeficiency virus type 1 reverse transcriptase heterodimer.

Proc Natl Acad Sci USA 91:7242–7246

8. Kohlstaedt LA, Wang J, Friedman JM, Rice PA, Steitz TA (1992)

Crystal structure at 3.5 A resolution of HIV-1 reverse transcrip-

tase complexed with an inhibitor. Science 256:1783–1790

9. Kaushik N, Rege N, Yadav PN, Sarafianos SG, Modak MJ,

Pandey VN (1996) Biochemical analysis of catalytically crucial

aspartate mutants of human immunodeficiency virus type 1

reverse transcriptase. Biochemistry 35:11536–11546

10. Dash C, Fisher TS, Prasad VR, Le Grice SF (2006) Examining

interactions of HIV-1 reverse transcriptase with single-stranded

template nucleotides by nucleoside analog interference. J Biol

Chem 281:27873–27881

11. Powell MD, Beard WA, Bebenek K, Howard KJ, Le Grice SF,

Darden TA, Kunkel TA, Wilson SH, Levin JG (1999) Residues in

the alphaH and alphaI helices of the HIV-1 reverse transcriptase

thumb subdomain required for the specificity of RNase H-cata-

lyzed removal of the polypurine tract primer. J Biol Chem

274:19885–19893

12. Hermann T, Meier T, Gotte M, Heumann H (1994) The ‘helix

clamp’ in HIV-1 reverse transcriptase: a new nucleic acid binding

motif common in nucleic acid polymerases. Nucleic Acids Res

22:4625–4633

13. Hermann T, Heumann H (1996) Strained template under the

thumbs. How reverse transcriptase of human immunodeficiency

virus type 1 moves along its template. Eur J Biochem 242:

98–103

14. Hsiou Y, Ding J, Das K, Clark AD Jr, Hughes SH, Arnold E

(1996) Structure of unliganded HIV-1 reverse transcriptase at

2.7 A resolution: implications of conformational changes for

polymerization and inhibition mechanisms. Structure 4:853–860

15. Jacobo-Molina A, Ding J, Nanni RG, Clark AD Jr, Lu X, Tantillo

C, Williams RL, Kamer G, Ferris AL, Clark P et al (1993) Crystal

structure of human immunodeficiency virus type 1 reverse

transcriptase complexed with double-stranded DNA at 3.0 A

resolution shows bent DNA. Proc Natl Acad Sci USA 90:6320–

6324

16. Ding J, Das K, Hsiou Y, Sarafianos SG, Clark AD Jr, Jacobo-

Molina A, Tantillo C, Hughes SH, Arnold E (1998) Structure and

functional implications of the polymerase active site region in a

complex of HIV-1 RT with a double-stranded DNA template-

primer and an antibody Fab fragment at 2.8 A resolution. J Mol

Biol 284:1095–1111

17. Sarafianos SG, Das K, Tantillo C, Clark AD Jr, Ding J, Whitcomb

JM, Boyer PL, Hughes SH, Arnold E (2001) Crystal structure of

HIV-1 reverse transcriptase in complex with a polypurine tract

RNA:DNA. EMBO J 20:1449–1461

18. Huang H, Chopra R, Verdine GL, Harrison SC (1998) Structure

of a covalently trapped catalytic complex of HIV-1 reverse

transcriptase: implications for drug resistance. Science 282:1669–

1675

19. Kiefer JR, Mao C, Braman JC, Beese LS (1998) Visualizing

DNA replication in a catalytically active Bacillus DNA poly-

merase crystal. Nature 391:304–307

20. Li Y, Korolev S, Waksman G (1998) Crystal structures of open

and closed forms of binary and ternary complexes of the large

fragment of Thermus aquaticus DNA polymerase I: structural

basis for nucleotide incorporation. EMBO J 17:7514–7525

21. Sawaya MR, Prasad R, Wilson SH, Kraut J, Pelletier H (1997)

Crystal structures of human DNA polymerase beta complexed

with gapped and nicked DNA: evidence for an induced fit

mechanism. Biochemistry 36:11205–11215

22. Fisher TS, Prasad VR (2002) Substitutions of Phe61 located in

the vicinity of template 50-overhang influence polymerase fidelity

and nucleoside analog sensitivity of HIV-1 reverse transcriptase.

J Biol Chem 277:22345–22352

23. Fisher TS, Darden T, Prasad VR (2003) Substitutions at Phe61 in

the beta3-beta4 hairpin of HIV-1 reverse transcriptase reveal a

role for the Fingers subdomain in strand displacement DNA

synthesis. J Mol Biol 325:443–459

24. Agopian A, Depollier J, Lionne C, Divita G (2007) p66 Trp24

and Phe61 are essential for accurate association of HIV-1 reverse

transcriptase with primer/template. J Mol Biol 373:127–140

25. Hsieh JC, Zinnen S, Modrich P (1993) Kinetic mechanism of the

DNA-dependent DNA polymerase activity of human immuno-

deficiency virus reverse transcriptase. J Biol Chem 268:24607–

24613

26. Pandey PK, Kaushik N, Singh K, Sharma B, Upadhyay AK,

Kumar S, Harris D, Pandey VN (2002) Insertion of a small

peptide of six amino acids into the beta7-beta8 loop of the p51

subunit of HIV-1 reverse transcriptase perturbs the heterodimer

and affects its activities. BMC Biochem 3:18

27. Arts EJ, Li X, Gu Z, Kleiman L, Parniak MA, Wainberg MA

(1994) Comparison of deoxyoligonucleotide and tRNA(Lys-3) as

primers in an endogenous human immunodeficiency virus-1 in

vitro reverse transcription/template-switching reaction. J Biol

Chem 269:14672–14680

28. Sharma B, Kaushik N, Singh K, Kumar S, Pandey VN (2002)

Substitution of conserved hydrophobic residues in motifs B and C

of HIV-1 RT alters the geometry of its catalytic pocket. Bio-

chemistry 41:15685–15697

29. Lee R, Kaushik N, Modak MJ, Vinayak R, Pandey VN (1998)

Polyamide nucleic acid targeted to the primer binding site of the

HIV-1 RNA genome blocks in vitro HIV-1 reverse transcription.

Biochemistry 37:900–910

30. Ausubel F, Brent R, Kingston RE, Moore DD, Seidman JS, Smith

JA, Struhl K (1987) Current protocols in molecular biology.

Greene Publishing Associates and Wiley-Intersciences, New

York

31. Harris D, Lee R, Misra HS, Pandey PK, Pandey VN (1998) The

p51 subunit of human immunodeficiency virus type 1 reverse

transcriptase is essential in loading the p66 subunit on the tem-

plate primer. Biochemistry 37:5903–5908

32. Pandey VN, Kaushik N, Rege N, Sarafianos SG, Yadav PN,

Modak MJ (1996) Role of methionine 184 of human immuno-

deficiency virus type-1 reverse transcriptase in the polymerase

function and fidelity of DNA synthesis. Biochemistry 35:2168–

2179

33. Astatke M, Grindley ND, Joyce CM (1995) Deoxynucleoside

triphosphate and pyrophosphate binding sites in the catalytically

competent ternary complex for the polymerase reaction catalyzed

by DNA polymerase I (Klenow fragment). J Biol Chem

270:1945–1954

34. Tong W, Lu CD, Sharma SK, Matsuura S, So AG, Scott WA

(1997) Nucleotide-induced stable complex formation by HIV-1

reverse transcriptase. Biochemistry 36:5749–5757

35. Sharma B, Kaushik N, Upadhyay A, Tripathi S, Singh K, Pandey

VN (2003) A positively charged side chain at position 154 on the

beta8-alphaE loop of HIV-1 RT is required for stable ternary

complex formation. Nucleic Acids Res 31:5167–5174

32 Mol Cell Biochem (2010) 338:19–33

123


36. Pandey PK, Kaushik N, Talele TT, Yadav PN, Pandey VN (2001)

The beta7-beta8 loop of the p51 subunit in the heterodimeric

(p66/p51) human immunodeficiency virus type 1 reverse trans-

criptase is essential for the catalytic function of the p66 subunit.

Biochemistry 40:9505–9512

37. Peletskaya EN, Boyer PL, Kogon AA, Clark P, Kroth H, Sayer

JM, Jerina DM, Hughes SH (2001) Cross-linking of the fingers

subdomain of human immunodeficiency virus type 1 reverse

transcriptase to template-primer. J Virol 75:9435–9445

38. Jorgensen W, Maxwell D, Tirado-Rives J (1996) Development

and testing of the OPLS all atom force field on conformational

energetics and properties of organic liquids. J Am Chem Soc

118:11225–11236

39. Polesky AH, Steitz TA, Grindley ND, Joyce CM (1990) Identi-

fication of residues critical for the polymerase activity of the

Klenow fragment of DNA polymerase I from Escherichia coli.J Biol Chem 265:14579–14591

40. Depollier J, Hourdou ML, Aldrian-Herrada G, Rothwell P, Restle

T, Divita G (2005) Insight into the mechanism of a peptide

inhibitor of HIV reverse transcriptase dimerization. Biochemistry

44:1909–1918

41. Tachedjian G, Radzio J, Sluis-Cremer N (2005) Relationship

between enzyme activity and dimeric structure of recombinant

HIV-1 reverse transcriptase. Proteins 60:5–13

42. Doublie S, Tabor S, Long AM, Richardson CC, Ellenberger T

(1998) Crystal structure of a bacteriophage T7 DNA replication

complex at 2.2 A resolution. Nature 391:251–258

43. Ling H, Boudsocq F, Woodgate R, Yang W (2001) Crystal

structure of a Y-family DNA polymerase in action: a mechanism

for error-prone and lesion-bypass replication. Cell 107:91–102

44. Matamoros T, Kim B, Menendez-Arias L (2008) Mechanistic

insights into the role of Val75 of HIV-1 reverse transcriptase in

misinsertion and mispair extension fidelity of DNA synthesis.

J Mol Biol 375:1234–1248

45. Kim B, Ayran JC, Sagar SG, Adman ET, Fuller SM, Tran NH,

Horrigan J (1999) New human immunodeficiency virus, type 1

reverse transcriptase (HIV-1 RT) mutants with increased fidelity

of DNA synthesis. Accuracy, template binding, and processivity.

J Biol Chem 274:27666–27673

46. Kim B, Hathaway TR, Loeb LA (1998) Fidelity of mutant HIV-1

reverse transcriptases: interaction with the single-stranded tem-

plate influences the accuracy of DNA synthesis. Biochemistry

37:5831–5839

Mol Cell Biochem (2010) 338:19–33 33

123


impact of template overhang-binding region of hiv-1 rt on the binding and orientation of the duplex...

Documents