Newly Synthesized APOBEC3G Is Incorporated
into HIV Virions, Inhibited by HIV RNA,
and Subsequently Activated by RNase H
Vanessa B. Soros1, Wes Yonemoto1, Warner C. Greene1,2,3*
1 Gladstone Institute of Virology and Immunology, San Francisco, California, United States of America, 2 Department of Medicine, University of California San Francisco, San
Francisco, California, United States of America, 3 Department of Microbiology and Immunology, University of California San Francisco, San Francisco, California, United States
APOBEC3G (A3G) is a potent antiretroviral deoxycytidine deaminase that, when incorporated into HIV virions,
hypermutates nascent viral DNA formed during reverse transcription. HIV Vif counters the effect of A3G by depleting
intracellular stores of the enzyme, thereby blocking its virion incorporation. Through pulse-chase analyses, we
demonstrate that virion A3G is mainly recruited from the cellular pool of newly synthesized enzyme compared to older
‘‘mature’’ A3G already residing in high-molecular-mass RNA–protein complexes. Virion-incorporated A3G forms a large
complex with viral genomic RNA that is clearly distinct from cellular HMM A3G complexes, as revealed by both gel
filtration and biochemical fractionation. Unexpectedly, the enzymatic activity of virion-incorporated A3G is lost upon
its stable association with HIV RNA. The activity of the latent A3G enzyme is ultimately restored during reverse
transcription by the action of HIV RNase H. Degradation of the viral genomic RNA by RNase H not only generates the
minus-strand DNA substrate targeted by A3G for hypermutation but also removes the inhibitory RNA bound to A3G,
thereby enabling its function as a deoxycytidine deaminase. These findings highlight an unexpected interplay between
host and virus where initiation of antiviral enzymatic activity is dependent on the action of an essential viral enzyme.
Citation: Soros VB, Yonemoto W, Greene WC (2007) Newly synthesized APOBEC3G is incorporated into HIV virions, inhibited by HIV RNA, and subsequently activated by
RNase H. PLoS Pathog 3(2): e15. doi:10.1371/journal.ppat.0030015
APOBEC3G (A3G) is a highly active antiretroviral deoxy-
cytidine deaminase that greatly impairs HIV spread in
cultures of activated CD4 T cells provided the HIV Vif
protein is absent . In these activated cells, the antiviral
action of A3G involves its effective incorporation into
budding virions and subsequent hypermutation of nascent
viral DNA formed during the next round of infection [2–6].
Vif has been proposed to block the incorporation of A3G into
HIV virions by targeting this enzyme for accelerated
degradation in the 26S proteasome [7–12] and partially
blocking its de novo synthesis [7,13]. A different situation
occurs in resting CD4 T cells and likely monocytes, which are
not permissive for HIV infection. In these cells, a low-
molecular-mass (LMM) form of cellular A3G is present, and it
functions as a potent postentry restriction factor for HIV by
blocking late reverse transcription . This antiviral action
of A3G is unchecked by Vif because insufficient quantities of
Vif are present in the incoming virions and the virus has not
progressed far enough into its life cycle to synthesize new Vif.
Thus, the growth of wild-type (WT) HIV is effectively
restricted in these cells by LMM A3G.
Incorporation of A3G into virions budding from HIV-
infected CD4 T cells has been proposed to involve assembly
with the nucleocapsid (NC) component of the Gag poly-
protein and/or viral genomic RNA [15–22]. Recent studies
with highly divergent Gag proteins  or treatment with
RNase A [16,18,19,22] suggest that Gag binding may be
indirect, involving an RNA intermediate. Following the entry
of A3G-containing virions into new target cells, A3G
deoxycytidine deaminase activity targets the minus-strand
DNA product of reverse transcription, leading to the
appearance of deoxyuridines in lieu of deoxycytidines at
canonical sites of deamination (59CC; the residue targeted for
A3G-mediated deamination is italicized) [1–6,24]. The non-
templated action of various DNA repair enzymes, including
uracil N-glycosylase, may mediate DNA strand cleavage ,
although a recent study suggests that uracil-N-glycosylase 2 is
dispensable for the antiviral action of A3G . If plus-strand
synthesis proceeds, dA residues are introduced at sites of dC
deamination, which results in dG-to-dA hypermutation in the
viral coding strand. These mutations may compromise HIV
infectivity by altering various viral open reading frames and
introducing inappropriate translation termination codons.
In contrast to the LMM form of A3G in resting CD4 T cells,
A3G in activated CD4 T cells principally resides in high-
molecular-mass (HMM) RNA–protein complexes . These
complexes include Staufen RNA transporting granules and
Editor: Richard A. Koup, National Institutes of Health, United States of America
Received August 14, 2006; Accepted December 18, 2006; Published February 9,
Copyright: ? 2007 Soros et al. This is an open-access article distributed under the
terms of the Creative Commons Attribution License, which permits unrestricted
use, distribution, and reproduction in any medium, provided the original author
and source are credited.
Abbreviations: A3G, APOBEC3G (apolipoprotein B mRNA-editing enzyme, catalytic
polypeptide-like); CA, Capsid; FPLC, fast protein liquid chromatography; HMM,
high-molecular-mass; IN, integrase; IVAC, intravirion A3G complex; LMM, low-
molecular-mass; NC, nucleocapsid; RNP, ribonucleoprotein; RT, reverse tran-
scriptase; ssDNA, single-stranded DNA; WT, wild-type
* To whom correspondence should be addressed. E-mail: firstname.lastname@example.org.
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150152
Ro/La ribonucleoprotein (RNP) complexes containing Alu
and hY retrotransposon RNA [27,28]. These complexes lack
detectable deoxycytidine deaminase activity in vitro but
interrupt Alu retrotransposition by sequestering the retroele-
ment RNA in the cytoplasm away from the requisite nuclear
LINE machinery. Treatment of these complexes with RNase A
promotes complex disassembly and generates the LMM,
enzymatically active form of A3G. Thus, the cellular forms
of A3G in resting and activated CD4 T cells are remarkably
different. The recruitment of A3G into HMM RNA–protein
complexes during the course of T-cell activation likely
explains why cellular A3G fails to function as a postentry
restriction factor for HIV in these activated cells.
The purposes of this study were to analyze the form of A3G
that is incorporated into HIVDVif virions and to assess its
enzymatic activity. Since virion A3G is readily able to mediate
hypermutation of viral DNA formed during reverse tran-
scription, we anticipated that enzymatically active forms of
A3G would predominate in virions. We have found that newly
synthesized A3G, not preexisting A3G already assembled into
the inactive cellular HMM complexes, is encapsidated into
budding virions. We also found that A3G recruited into
virions assembles with viral RNA to form a large intravirion
A3G complex (IVAC) that is enzymatically inactive. Finally,
we have demonstrated that the action of viral RNase H during
reverse transcription ultimately releases A3G from its state of
inhibition, allowing hypermutation of the minus-strand viral
DNA. Thus, activation of the enzyme-dependent antiviral
action of A3G appears to critically depend on the action of an
HIV enzyme, RNase H.
A3G Recruited into Virions Cofractionates with Virion
Initially, we sought to identify transfection conditions for
the generation of A3G-containing HIV virions that would
recapitulate the virion encapsidation of A3G that naturally
occurs in T cell lines and primary T cells infected with
HIVDVif viruses. Activated blood-derived primary CD4 T
cells or H9 T cells expressing endogenous A3G were
spinoculated with HIVDVif and emergent viruses were
harvested 2 d postinfection. In parallel, 293T cells were
cotransfected with a fixed dose of proviral expression plasmid
DNA and increasing doses of A3G expression plasmid DNA.
Virions were similarly collected from the transfected cells 2 d
later, after purification of virions by ultracentrifugation
through iodixanol cushions. Virion lysates were subjected to
immunoblotting to determine the amount of A3G incorpo-
rated relative to p24-CA content. These virion preparations
were not contaminated with significant cellular material or
microvesicles, as determined by immunoblotting with anti–
14-3-3c and anti-CD45 antibodies (Figure S1). Transfection of
increasing amounts of A3G expression plasmid resulted in
increasing amounts of A3G virion incorporation relative to
p24-Capsid (CA) (Figure 1A and 1B). However, when
compared to virions produced from infected primary CD4
T cells or H9 T cells, the 293T transfection conditions that
best recapitulated the ‘‘natural’’ packaging levels of endoge-
nously expressed A3G were achieved at a plasmid microgram
ratio of 2 (HA-A3G) to 60 (pNL4–3DVif), which equals a
molar ratio of 1 (HA-A3G) to 12.5 (pNL4–3DVif). Thus, this
condition was used for the production of A3G-containing
virions in all the subsequent experiments unless otherwise
To determine if A3G is packaged into the core of HIV
virions and to assess the localization of the additional A3G
packaging that occurred at higher transfection doses, virions
were subjected to biochemical fractionation. The virion
envelope was removed by brief solubilization with Triton X-
100, as previously described , yielding separable virion
cores containing p24-CA, integrase (IN), reverse transcriptase
(RT), and NC (Figure 1C). Fractionation of viruses containing
A3G at levels comparable to those of virions budding from
primary CD4 T cells revealed that A3G is indeed packaged
into virion cores (Figure 1C). However, virions derived from
cells expressing higher levels of A3G (for example, 20 lg of
HA-A3G:60 lg of pNL4–3DVif) packaged a similar amount of
A3G into virion cores as well as additional A3G that
fractionated into the gp41-containing supernatant after
Triton X-100 solubilization (Figure 1A–1C). These findings
suggest that when A3G is overexpressed in virus-producing
cells, considerable amounts of the enzyme reside outside the
viral core, likely in the viral matrix region, between the core
and outer envelope (for example, 1:3 lg ratio). The
appearance of approximately half the virion p24-CA in the
Triton X-100 supernatant after solubilization similarly
reflects excess Gag packaged into virions that does not
contribute to core formation . Thus, A3G may gain access
to HIV virion cores through a specific interaction with viral
RNA and/or Gag (NC). However, under conditions of over-
expression, a lower-affinity interaction, perhaps directly
between A3G and Gag, results in the recruitment of
additional enzyme into the detergent-sensitive matrix space,
into which excess Gag is also packed .
Newly Translated A3G Is Incorporated into HIV Virions
Under steady-state conditions in activated CD4 T cells,
cellular A3G resides in an HMM RNase A–sensitive complex
of 5 to 15 MDa [14,27]. The observation that HA-A3G
PLoS Pathogens | www.plospathogens.orgFebruary 2007 | Volume 3 | Issue 2 | e150153
Biochemical Analysis of Intravirion A3G
APOBEC3G (A3G) is a cellular enzyme that promotes DNA muta-
genesis and can restrict infection by HIV-1. However, HIV counters
the antiviral effects of A3G through the action of its Vif protein. In
the absence of Vif, A3G is effectively incorporated into virions, where
it mutagenizes the first DNA copy (cDNA) generated during reverse
transcription of the viral RNA genome. A3G also appears to be able
to inhibit HIV via nonenzymatic mechanisms. A3G and related
deoxycytidine deaminases can also inhibit the growth of retro-
viruses other than HIV and protect the cellular genome from
endogenous mobile retroelements. In this study, we analyzed the
recruitment and enzymatic activity of A3G incorporated into HIVDVif
virions. Unexpectedly, we found that the binding of A3G to viral
genomic RNA led to inactivation of the enzyme. However, latent
A3G was ultimately activated through the action of HIV RNase H,
which degrades the RNA genome during reverse transcription.
These findings highlight an unexpected interplay between a host
enzyme and HIV, where the antiviral enzymatic activity of the host
factor (A3G) is dependent on the action of an essential HIV enzyme
(RNase H). The strong interaction with viral RNA also suggests a
potential mechanism by which A3G could exert antiviral activity in
the absence of enzymatic activity, by physically impeding reverse
incorporated into virions packages into the virion core
suggested several possible cellular sources of the enzyme. The
first possibility, albeit unlikely, is that entire 5- to 15-MDa
cellular A3G complexes are recruited into the virion core.
Alternatively, HIV RNA and Gag may promote release of A3G
from the cellular HMM A3G complex, allowing its recruit-
ment into the virion, with or without a limited subset of
cellular cofactors, as has been suggested . Finally, newly
synthesized LMM A3G not yet assembled with cellular
cofactors or RNA may be recruited into the virion through
its association with viral RNA and Gag. To determine if newly
synthesized A3G, more ‘‘mature’’ A3G, or both serve as
Figure 1. A3G Is Incorporated into Virion Cores but A3G Overexpression in Cells Results in Additional A3G Packaging Outside of the Core
(A) HA-A3G–containing DVif virions were generated from 293T cells transfected with a fixed amount of proviral plasmid (60 lg) and increasing doses of
HA-A3G (0 to 20 lg). Empty HA vector (0 to 20 lg) was used as balance DNA in the transfections. Sample number 1¼0 (lg of HA-A3G):60 (lg of pNL4-
3DVif), 2 ¼ 1:60, 3 ¼ 2:60, 4 ¼ 5:60, 5 ¼ 10:60, 6 ¼ 20:60. DVif virions were also derived from the H9 T cell line and primary CD4 T cells, which
endogenously express A3G. The virion lysates were subjected to immunoblotting with antibodies specific for p24-CA and A3G. The immunoblot is
representative of several independent analyses used to generate the graph in (B).
(B) Graphical representation of quantification from immunoblots in (A) and unpublished data. Data are averaged from three independent transfections
of 293T cells, five independent spinoculations of activated primary CD4 T cells, and three independent spinoculations of H9 T cells. The error bars
represent standard deviation. The relative ratio of packaged A3G to p24-CA is plotted, with virions derived from CD4 T cells assigned a value of 1.
(C) Virions containing increasing amounts of HA-A3G relative to p24-CA were solubilized by brief Triton X-100 treatment to generate virion cores
containing p24-CA, IN, RT, and NC and supernatants containing gp41 and p24-CA. The triangles represent the increasing dose of A3G and correspond
exactly to the numbered samples presented in Figure 1A. The immunoblots (IB) were also probed for A3G to determine the amount packaged into
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150154
Biochemical Analysis of Intravirion A3G
cellular reservoirs for virion recruitment, we performed
pulse-chase radiolabeling studies. First, the time course for
recruitment of newly synthesized HA-A3G into cellular HMM
complexes in the absence of proviral gene expression was
determined. Cells were pulsed with radiolabel for 10 min,
followed by chases of 30 min to 3 h. Size-fractionation (Figure
S2A) of the pulse and chase lysates identified the presence of
pulse-labeled HA-A3G initially in low and intermediate mass
fractions (Figure 2A, t ¼ 0 fractions 6 and 7) that was chased
into HMM complexes within 30 min (Figure 2A, A3G in
fractions 4 and 5 at 0.5 and 1 h). The radiolabeled A3G
remained stably associated with the cellular HMM complex
during longer chase periods (Figures 2A and S2B). Thus,
newly synthesized A3G is initially LMM and recruited within
30 min into stable cellular HMM RNA–protein complexes.
When HIVDVif was coexpressed, we observed that the
Figure 2. Newly Synthesized HA-A3G Is Recruited into HIV-1 Virions
(A) Cells transfected with HA-A3G were pulse radiolabeled for 10 min, chased with cold medium, and harvested at 0, 0.5, and 1 h. Chase lysate was
subjected to Sepharose CL-6B gel filtration (10 fractions collected per lysate), HA-A3G was immunoprecipitated from each fraction, and the relative
amount of radiolabeled HA-A3G in each fraction was determined by autoradiography. Fraction 4 corresponds to HMM complexes, and fractions 6 to 8
contain proteins similar in size to LMM A3G. See Figure S2A for Sepharose CL-6B fractionation performance.
(B) Cells transfected with HA-A3G and pNL4–3DVif were assessed as in (A).
(C) Cells transfected with HA-A3G were assessed as in (A), except that the pulse was extended to 30 min. Plotted is the t ¼ 0-h pulse sample.
(D) Cells transfected with HA-A3G and pNL4–3DVif were assessed as in (C).
(A–D) Data shown are representative of three independent experiments for each panel.
(E) Virus-producing cells were pulse radiolabeled and chased with cold medium, and both cells and virus-containing supernatants were collected at 0.5,
1, 2, and 4 h. As all of the supernatant was collected at the indicated time points, the virions harvested represent only those that budded during the
intervening time period. Plotted is the percent density of the immunoprecipitated protein for a given time point relative to the total radioactive density
of all time points. Upper panels, immunoprecipitates of HA-A3G from the producer cells. Middle panels, immunoprecipitates of HA-A3G from virions.
Lower panels, immunoprecipitates of p24-CA from virions. Data are from three experiments performed independently.
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150155
Biochemical Analysis of Intravirion A3G
presence of viral RNA and proteins did not alter the ability of
newly synthesized LMM A3G to assemble into cellular HMM
RNA–protein complexes (Figure 2B).
To assess whether newly synthesized or more ‘‘mature’’
preexisting cellular HMM A3G is recruited into virions, we
performed similar pulse-chase radiolabeling studies in cells
producing HIV. To enhance radiolabeling and detection of
intravirion A3G, the length of the pulse was extended from 10
min to 30 min. The longer pulse time did not affect A3G
assembly into cellular HMM complexes, in either the absence
(Figure 2C) or presence (Figure 2D) of HIVDVif expression;
however, it did mask the chase of LMM A3G into HMM A3G.
Expression of HIVDVif also did not affect the turnover of
radiolabeled A3G (Figure S2D). Both producer cells and their
supernatants containing virions were collected and analyzed
simultaneously for radiolabeled HA-A3G and p24-CA con-
tent. Since all of the virus-containing supernatants were
collected at the indicated time points, the radiolabel present
in virion p24-CA or HA-A3G reflects labeling events
occurring during the discrete intervening time points. In
each of the three independent experiments shown, incorpo-
ration of radiolabeled p24-CA into virions increased with
increasing chase time over the first 1 to 2 h and then declined
by 4 h (Figure 2E, lower panels). The incorporation pattern of
p24-CA over time is consistent with previous reports [31,32]
showing increased accumulation of radiolabeled p24 in
virions with long cumulative chase times. In contrast, HA-
A3G incorporation into virions displayed a sharp spike
between 30 and 60 min after the pulse (Figure 2E, middle
panels), even though large cellular pools of radiolabeled A3G
were present both before and after this time point (Figure 2E,
upper panels). Specifically, despite the presence of radio-
labeled cellular HA-A3G at the 2- and 4-h time points, these
pools of A3G were not effectively incorporated into virions
compared to the 1-h time point. The distinct peak of
incorporation of radiolabeled A3G at 1 h after pulse also
was not due to a relative loss of radiolabeled cellular A3G
available for virion incorporation at the later collection
times, since normalization by the available radiolabeled pool
of cellular enzyme did not alter the distinct early kinetic
pattern for A3G incorporation into virions (Figure S2C).
These findings indicate that newly synthesized A3G less than
1.5 h old is incorporated HIV virions and that older, more
‘‘mature’’ A3G in HMM complexes is apparently less available
for virion incorporation during the time course examined
here. Using a modified version of these experiments
(infection with HIVDVif instead of transfection of a proviral
DNA plasmid) and extended chase times, we observed a
similar trend of newly synthesized radiolabeled A3G incor-
poration into virions and low-to-undetectable levels of
radiolabeled A3G in virions up to 9 h after the pulse period
A3G Assembles into a Large RNA-Containing Intravirion
Complex (Intravirion A3G Complex)
The finding that newly synthesized A3G is packaged into
virion cores (Figure 1C) coupled with the observation that
newly synthesized A3G rapidly forms HMM complexes in cells
(Figure 2A–2D) led us to next examine whether intravirion
A3G resolves as monomers/dimers or instead as a larger
complex. We hypothesized that A3G in virions might remain
in an enzymatically active LMM form, because it ultimately
deaminates the viral minus-strand DNA synthesized during
reverse transcription. Lysates derived from virions contain-
ing HA-tagged A3G were size-fractionated by fast protein
liquid chromatography (FPLC). Each fraction was then
analyzed by SDS-PAGE and immunoblotting with anti-HA
monoclonal antibodies (Figure 3A). Surprisingly, virion-
incorporated HA-A3G was detected almost exclusively in
the HMM region, eluting in the void volume of the Superose 6
column (Figure 3A). This result was not due to incomplete
lysis of the virion cores since the p24 viral capsid (p24-CA)
was detected only in the expected LMM fractions.
To determine if the IVAC contains an RNA component,
virion lysates were treated with RNase A (Figure 3B). Under
these conditions, HA-A3G shifted to LMM fractions consis-
tent in size with monomers and/or dimers of the enzyme.
Therefore, reminiscent of the cellular forms of A3G present
in activated CD4 T cells  (and in HIV-producing cells, as
shown below), A3G incorporated into virion assembles into
large RNA–protein complexes that are distinct from the
cellular HMM complexes (see below).
As an alternative approach to examine whether A3G in
virion cores were indeed freely soluble or associated with
other factors, cores purified by solubilization of whole virions
(Figure 1C) were biochemically disassembled by exposure to a
low pH ‘‘STE’’ buffer at 37 8C, as previously described .
This treatment resulted in release of p24-CA into the
supernatant of a pelletable viral RNP complex consisting of
IN, NC, and viral genomic RNA. Under these conditions, RT
is more readily released from the RNP upon biochemical
fractionation of the cores (Figure 3C and as previously
described [34,35]). Analysis of A3G-containing virion cores
revealed that IVAC A3G cofractionated with the viral RNP
proteins (Figure 3C), suggesting a continued association with
the viral genomic RNA and/or NC protein.
Enzymatic Activity of A3G in the IVAC Is Negatively
Regulated by RNA Binding
Since virion-derived A3G ultimately exerts deoxycytidine
deaminase activity during reverse transcription [1–6], we
considered the possibility that A3G might remain enzymati-
cally active even when bound to RNA in the IVAC. However,
in an in vitro deoxycytidine deaminase assay, the IVAC
HAA3G exhibited no detectable enzymatic activity (Figure
4A). Like the cellular HMM A3G complex (Figure 4D and
), deoxycytidine deaminase activity was readily detected
when the virion HA-A3G immunoprecipitates were pre-
treated with RNase A before assessment (Figure 4A). Analysis
of whole-virion lysate similarly showed that RNase A treat-
ment was required for detection of A3G enzymatic activity in
vitro (Figure 4B). These findings indicate that HA-A3G is
incorporated into virions as an enzymatically latent large
RNP complex. Of note, previous reports have observed
readily detectable enzymatic activity from A3G-containing
virions. We believe this may be due to the presence of
additional, noncore LMM A3G packaged into the matrix
space of virions upon A3G overexpression in cells (Figure
1C). When virions containing increasing amounts of A3G
(Figure 1A) were tested for in vitro deaminase activity,
substrate was readily deaminated by those virions which
contained higher proportions of A3G to p24-CA than is
normally packaged by infected CD4 T cells (Figure 4C). In
addition, Yu et al.  reported that virion-derived A3G was
PLoS Pathogens | www.plospathogens.orgFebruary 2007 | Volume 3 | Issue 2 | e15 0156
Biochemical Analysis of Intravirion A3G
enzymatically active in the absence of RNase treatment.
However, in that study, virions were extracted in buffers
containing EDTA , an agent that disrupts some RNA–
protein complexes [36,37]. When we analyzed virion HA-A3G
extracted in EDTA-containing buffers, we also detected
deoxycytidine deaminase activity, suggesting that this treat-
ment likely activated A3G by promoting its dissociation from
inhibitory RNA(s) (Figure S3). Likewise, it has been observed
that the addition of salts like magnesium that promote and
stabilize RNA tertiary structure enhance the activity of
recombinant A3G purified from insect cells while inducing
a shift of A3G from large to intermediate-sized complexes
. Together these findings support the notion that RNA
binding inhibits A3G enzymatic activity, likely by occluding
the catalytic site.
IVAC Contains HIV Genomic RNA
The reported recruitment of A3G into virions through viral
RNA, the cofractionation of intravirion A3G with viral RNP
proteins (Figures 1C and 3C), the assembly of A3G into a large
RNase A–sensitive complex within virions (Figure 3A), and
the RNase A–dependent in vitro enzymatic activity of
intravirion A3G (Figure 4A and 4B) strongly suggested that
HIV genomic RNA may be an important constituent, perhaps
even a nucleating factor for IVAC assembly. To test this
possibility, we immunoprecipitated A3G from IVAC frac-
tions, purified the RNA, and subjected it to RT-PCR with
primers that specifically amplify HIV genomic RNA. Genomic
HIV RNA was readily detected in the IVAC as well as in A3G
immunopre0cipitates prepared from virus-producing cells
Figure 3. Virion-Incorporated HA-A3G Resides in a Large RNase A–Sensitive Complex and Biochemically Fractionates with Viral RNP Proteins
(A) Virions collected from cells expressing HIV-1DVif contain HA-A3G that predominantly fractionates in a large complex (fractions 6 to 8) as assessed by
(B) The IVAC is sensitive to RNase A treatment which shifts HA-A3G into lower fractions (fractions 15 to 19).
(C) Virion cores obtained in Figure 1 were subjected to further biochemical fractionation to generate viral RNPs. Shown are the viral RNPs from virions
either lacking or containing A3G, as indicated, and containing viral RT, IN, and NC but not p24-CA, as detected by immunoblotting (IB). The triangles
represent the increasing dose of A3G relative to provirus and correspond exactly to the sample numbers in Figure 1A.
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150157
Biochemical Analysis of Intravirion A3G
Next, we compared the FPLC fractionation profile of HIV
RNA derived from virions that contained or lacked HA-A3G.
In the presence of HA-A3G, HIV RNA was detected in
fractions that contained the IVAC, indicative of A3G-
dependent assembly of large HIV RNA–protein complexes
in virions (Figure 5B). In the absence of HA-A3G, viral RNA
was detected in lower fractions 11 through 14. The fractions
were determined to contain full-length genome by the
production of PCR products using probes that amplify across
various regions of the genome. However, these gel filtration
experiments were associated with some RNA fragmentation,
particularly in the absence of A3G (Figure 5B, TAR/Gag
amplicons were observed in fractions 11 through 20). We
suspect that such fragmentation also occurred in A3G-
containing virions but that the RNA fragments continued
to resolve in the IVAC fractions through persistent associa-
tion with A3G.
Of note, virion NC, which also binds HIV RNA, was
detected in the FPLC fractions that contained IVAC (Figure
5C). Conversely, RT, which is more readily released from HIV
RNA upon biochemical manipulation/lysis of virions [34,35]
(and Figure 3C), did not display a strong shift into the IVAC
fractions in the presence of HA-A3G upon gel filtration. As
expected, the non–RNP-associated gp41 viral protein re-
solved independently of A3G. In the absence of A3G, NC
resolved in lower FPLC fractions, consistent with pools of
protein that either have dissociated from the viral RNP or
remain associated with RNA fragments (possibly caused by
the gel filtration conditions, as discussed above). In both the
absence  and presence of HIV gene expression (Figure
5D), cellular A3G resolves as HMM upon gel filtration.
Inactive A3G in the IVAC Is Activated by Viral RNase H
during Reverse Transcription
Next our studies focused on how the latent deoxycytidine
deaminase activity of A3G present in the IVAC is ultimately
Figure 4. Intravirion A3G Enzymatic Activity Is Negatively Regulated by Binding to Genomic HIV RNA
(A) HA-A3G was immunoprecipitated from IVAC fraction 7 (F7) of virion lysates (Figure 3A) or from a lower fraction, F17, generated by treatment of the
virion lysates with RNase A (Figure 3B). Immunoprecipitates (IPs) were tested for enzymatic activity in an in vitro deoxycytidine deaminase assay with or
without RNase A addition and contained equivalent amounts of HA-A3G as shown in the corresponding immunoblot. The generation of a shorter
cleavage product from the input ssDNA substrate reveals A3G deoxycytidine deaminase activity. Data shown are representative of multiple
(B) Lysates of virions containing or lacking A3G were assessed in the deaminase assay, with or without RNase A treatment.
(C) Lysates of virions containing increasing amounts of HA-A3G (as shown in the corresponding immunoblot) were assessed in the deaminase assay,
with or without RNase A treatment. The asterisk marks bleed-through of marker loaded to the left of the samples. The triangles represent the increasing
dose of A3G relative to provirus and correspond to the sample numbers presented in Figure 1A.
(A–C) All deaminase reactions were carried out in 50 mM Tris (pH 7.4) with (þ) or without (?) RNase A, as indicated.
(D) IPs of HMM or LMM HA-A3G from producer cell lysates were similarly assessed in the deaminase assay, with (þ) or without (?) added RNase A. The
IPs contained equivalent amounts of HA-A3G as shown in the corresponding immunoblot (IB).
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150158
Biochemical Analysis of Intravirion A3G
activated. As shown in Figure 4A and 4B, the simple addition
of single-stranded DNA (ssDNA) substrate was insufficient for
triggering its activity. In view of the effects of RNase A
treatment in vitro, we were intrigued by the possibility that
viral RNase H enzyme might play a role in the activation of
A3G enzymatic activity. RNase H resides near the C-terminus
of the large subunit of the p66-p51 RT heterodimer [39,40].
The DNA-dependent action of RNase H is required for
commencement of second-strand synthesis and concomi-
tantly generates the free, minus-strand ssDNA substrate that
is targeted by A3G for deamination [2–6]. We hypothesized
that viral RNase H action might not only generate the
substrate for A3G-mediated deamination but also reverse the
RNA-mediated inhibition of A3G deoxycytidine deaminase
activity by degrading the genomic RNA bound to A3G. To
examine this possibility, we established in vitro conditions
Figure 5. Virion-Incorporated HA-A3G Associates with Viral Genomic RNA
(A) Viral genomic RNA, detected by RT-PCR, was detected in virions and virus-producing cells but not in lysates of uninfected cells. Genomic RNA was
also detected in the IVAC derived from virions (fraction 7) and coimmunoprecipitated with HA-A3G from both virions and producer cell lysates. RT was
performed using RNA derived from either whole lysates (L) or anti-HA immunoprecipitates (IP). Control reactions were performed in the absence of RT
(–RT). Control PCRs were performed using proviral plasmid DNA, in the absence or presence of Taq, as indicated.
(B) Viral genomic RNA, detected by RT-PCR, was assessed from size-fractionated virion lysates that lacked (HA) or contained HA-A3G. Amplicons
generated probed across the TAR/Gag region or Pol/Vpu regions, as indicated.
(C) Incorporation of HA-A3G into virions enhances the recruitment of NC into the IVAC.
(D) HA-A3G from virus-producing cells is HMM and is converted to LMM form after RNase A treatment. ‘‘IB’’ indicates immunoblotting with the
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150159
Biochemical Analysis of Intravirion A3G
that lead to RNase H activity and assessed the effects of active
RNase H on A3G deaminase activity. Both recombinant
purified RT and virion-derived RT cleaved an end-labeled
RNA oligonucleotide from an RNA–DNA hybrid substrate
(Figure 6A and 6B). Importantly, RNase H activity was
magnesium dependent , was inhibited by a variety of
small molecules including Compound I [42,43], and was
compromised by specific mutations within its catalytic
domain, for example, E478Q ( and Figure 6A and 6B).
We used these various properties of RNase H and reagents to
probe the potential involvement of RNase H as an activator
of latent A3G deoxycytidine deaminase activity within
First, we tested whether stimulation of endogenous reverse
transcription in virion lysates by the addition of magnesium
and deoxynucleotide triphosphates promoted activation of
Figure 6. Enzymatically Inactive Virion-Incorporated HA-A3G Is Activated by Viral RNase H
(A) Recombinant RTs containing either a WT or mutant (E478Q) RNase H catalytic domain were assessed for RNase H activity in vitro in the absence or
presence of the RNase H inhibitor Compound I (final concentration of 1, 10, or 100 lM). The RNA of an RNA–DNA hybrid remains intact unless RNase H
digests the RNA into a smaller cleavage product that is distinguishable from the more complete cleavage product generated by RNase A. WT RNase H
cannot digest ssDNA or DNA of an RNA–DNA hybrid, or RNA–RNA hybrids (data not shown). RNase H assays were performed in RNase H buffer (50 mM
Tris [pH 8.0], 60 mM KCl) with (þ) or without (?) 5 mM MgCl2or RNase A, as indicated.
(B) Viruses bearing the RNase H E478Q mutation are compromised for in vitro RNase H activity. RNase H assays were performed in RNase H buffer with
(þ) or without (?) 5 mM MgCl2or RNase A, as indicated.
(C) Virion lysates were subjected to endogenous reverse transcription (enRT) conditions with or without Compound I (final concentration of 0.1, 1, 10, or
100 lM), and A3G activity in these samples assessed in the in vitro deoxycytidine deaminase assay. Deaminase assays were performed in RNase H buffer
either supplemented (enRT:þ) or not (enRT:?) with 4 mM MgCl2and 1 mM dNTPs.
(D) Compound I does not inhibit the intrinsic deoxycytidine deaminase activity of A3G. HA-A3G from RNase A–treated virion lysates was assessed for in
vitro deaminase activity in the presence of increasing doses of Compound I (0.1, 1, 10, and 100 lM). Deaminase assay was performed in RNase H buffer
supplemented with RNase A only.
(E) Virions containing WT RNase H or the E478Q mutation in the RNase H catalytic domain were subjected to the enRT reaction followed by assessment
of A3G enzymatic activity. Deaminase assays were performed in RNase H buffer either supplemented (enRT:þ) or not (enRT:?) with 4 mM MgCl2and 1
(F) WT and RNase H–compromised DVif virions containing WT or mutant RNase H displayed equivalent A3G activity when RNase A was added to the
virion lysate. Deaminase assay was performed in RNase H buffer with (þ) or without (?) RNase A, as indicated.
All data are representative of multiple experiments.
PLoS Pathogens | www.plospathogens.orgFebruary 2007 | Volume 3 | Issue 2 | e150160
Biochemical Analysis of Intravirion A3G
A3G enzymatic activity measured in the deaminase assay.
Such treatment effectively induced readily detectable deox-
ycytidine deaminase activity, suggesting a link between
reverse transcription and A3G activation (Figure 6C). Of
note, the appearance of deoxycytidine deaminase activity was
blocked in a dose-dependent manner by Compound I, an
RNase H inhibitor. Importantly, Compound I did not impair
the deoxycytidine deaminase activity of A3G induced by prior
RNase A treatment (Figure 6D), supporting inhibition of
RNase H as the cause of Compound I–mediated inhibition of
A3G enzyme activation. Additionally, the introduction of a
point mutation in the catalytic core of RNase H (E478Q),
which compromised RNase H activity (Figure 6B), also
impaired activation of A3G deoxycytidine deaminase activity
under conditions permissive for endogenous reverse tran-
scription (Figure 6E). HA-A3G was otherwise equally active
upon RNase A treatment of viruses bearing either WT or
compromised RNase H domains (Figure 6F). These findings
demonstrate that, in addition to generating the substrate for
A3G-mediated deamination, HIV-1 RNase H plays a central
role in triggering the activity of the latent virion-associated
Our observation that A3G packages into HIV virion cores
is not unexpected given that RNA and/or the NC region of
Gag recruit the enzyme into the virus [15–22]. Further, A3G
antiviral activity is ultimately manifested during reverse
transcription and therefore proximity to reverse transcrip-
tion complexes would be anticipated. Indeed, Khan et al. 
reported core localization of A3G. However, it is clear from
the virion fractionations that additional enzyme may gain
access to the virion when A3G is overexpressed in virus-
producing cells (Figure 1). This additional A3G is not
specifically recruited into virion cores. It has been previously
demonstrated that roughly twice as much Gag than that
present in the virion core is incorporated into immature
particles . Based on the RNase A sensitivity of the
interaction between Gag (NC) and A3G [16,18,19,22] and
the ability of A3G to interact with highly divergent Gag
proteins [21,23,45–47], it has been suggested that A3G may
recognize an NC–RNA interface that promotes virion
incorporation [23,47]. Alternatively, the conflicting reports
regarding A3G virion recruitment by Gag/NC in an RNA-
dependent [16,18,19,22] or RNA-independent [15,17,20,21]
manner could be a consequence of the relative amounts of
additional, non–core packaging occurring under the con-
ditions of assay. The RNase-sensitivity of Gag interaction may
be observable only at lower (endogenous-like levels) A3G
concentrations where affinity is perhaps governed primarily
by RNA interactions. At higher expression levels of A3G, a
lower affinity but direct interaction with Gag independent of
RNA may occur. Whether virion core–incorporated A3G, like
NC, coats the viral RNA or whether A3G binding is restricted
to certain regions  remains to be determined.
Differences in the absolute amount of A3G packaged into
virions may also contribute to apparent disparities in prior
studies of A3G. Within this study, for example, the additional
packaging of A3G under conditions of A3G overexpression
masked the RNase-dependent ‘‘activation’’ of virion A3G
(Figure 4C), highlighting the importance of establishing
conditions that closely recapitulate physiological levels of
A3G incorporation into virions.
Several observations in this study support the notion that
A3G incorporated into HIV virion cores is assembled into a
large RNA–protein complex that we have termed the IVAC.
First, A3G was incorporated into virion cores (Figure 1C),
which contain viral RNP complexes consisting of viral
genomic RNA, NC, IN, and Vpr. IVAC A3G both coimmu-
noprecipitated viral genomic RNA (Figure 5) and cofractio-
nated with the virion core proteins (Figure 3C). Additionally,
the shift in viral genomic RNA from lower to higher mass
FPLC fractions upon HA-A3G expression supports the
notion that RNA is critical for virion packaging of A3G and
suggests its possible central role in nucleation of the IVAC.
Importantly, although the resolving power of our fractiona-
tion is currently not able to differentiate the sizes of the
cellular HMM A3G complexes and IVAC (all resolve at or
near the void volume of the Superose 6 column), these
complexes are not identical. For instance, the cellular HMM
complexes form in the absence of viral genomic RNA in
activated but uninfected CD4 T cells , while IVAC A3G
interacts with HIV RNA (Figure 5).
Second, our preliminary results indicate that many of the
protein components of the cellular HMM A3G complex
[27,28,48] are not corecruited into HIV virions (unpublished
data and ). Finally, the activation of IVAC A3G by in vitro
endogenous reverse transcription (Figure 6) suggests that
viral RNA inhibits IVAC A3G enzymatic activity unless
removed by RNase H, a virally encoded enzyme that acts on
the RNA component of RNA–DNA hybrids. Of note, the level
of A3G activation obtained when endogenous reverse tran-
scription is stimulated (Figure 6E) was consistently less robust
than the level of enzymatic activity observed when the IVAC
was treated with RNase A (Figure 6F). We suspect this finding
reflects a more complete clearance of RNA from IVAC A3G
by exogenously added RNase A than occurs with RNase H
activation under conditions of endogenous reverse tran-
scription. Alternatively, A3G incorporated into HIV virions
may bind both HIV RNA and non-HIV RNA (for example, the
tRNA-Lys3primer); however, since only the viral RNA genome
is reverse transcribed, thereby forming a substrate for RNase
H activity, only viral RNA-bound A3G may become ‘‘acti-
vated’’ during reverse transcription.
The pulse-chase radiolabeling studies of A3G in cells
revealed that newly synthesized, LMM A3G is rapidly (within
30 min) recruited into cellular HMM complexes (Figure 2A)
and that proviral gene expression has little, if any, effect on
A3G assembly into HMM RNA–protein complexes (Figure
2B). Extension of the pulse radiolabeling time did not impede
assembly but masked detection of the rapid assembly of newly
synthesized LMM A3G into HMM complexes, in both the
absence (Figure 2C) and the presence (Figure 2D) of viral
gene expression. Notably, these complexes appear to be
stable for at least several hours (Figure S2B).
The assembly of intravirion A3G into a large RNP complex
could result from recruitment of any of the cellular HMM
A3G complexes (Staufen RNA transporting granules, Ro/La
RNPs) into virions or by HIV RNA extraction of A3G from
cellular HMM A3G complexes. Alternatively, newly synthe-
sized A3G not yet assembled into fully mature cellular HMM
complexes could bind to HIV RNA, which in turn targets the
enzyme for encapsidation into HIVDVif virions. As noted,
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150161
Biochemical Analysis of Intravirion A3G
A3G assembled into HMM A3G complexes or A3G assembled
with HIV RNA and core proteins sieve near the void volume
of the Superose 6 columns and thus these different types of
complexes cannot be distinguished by FPLC. To investigate
whether newly synthesized A3G or older, ‘‘mature’’ A3G
already assembled into HMM complexes is recruited into HIV
virions, virus-producing cells were subjected to pulse-chase
radiolabeling studies. In each of three experiments, we
observed the appearance of a peak of radiolabeled A3G in
virions at one discrete time point, occurring between 0.5 h
and 1 h after the pulse (Figure 2E, middle panels). Radio-
labeled A3G incorporation into virions decreased dramati-
cally after this peak, despite the persistence of substantial
pools of radiolabeled A3G in the producer cells from which
the virions were derived (Figure 2E, top panel). The loss of
radiolabeled A3G in virions after this peak also could not be
explained by a sharp decline of radiolabeled A3G in the
producer cells (Figure S2C). These findings suggest that once
A3G assembles into the cellular HMM A3G complexes
[27,28,48], it may no longer serve as a major reservoir of
enzyme for virion encapsidation. Pulse radiolabeling of A3G
before the peak of viral Gag expression and extension of the
collection times up to 9 h after the pulse further confirmed
that mature HMM cellular A3G does not form a major pool of
enzyme for incorporation into HIVDVif virions (Figure S2E).
Instead, it appears that newly synthesized A3G is prefer-
entially recruited into HIV virions within 1.5 h after synthesis
(Figure 2E, middle panels). Interestingly, the appearance of
radiolabeled HA-A3G in virions appeared to be slightly
delayed since samples collected (1) immediately after the
pulse and (2) in the first 30-min chase contained relatively
little radiolabeled A3G compared to virions collected over
the second 30-min chase (Figure 2E, 1-h collection time
point). This is clearly within the time required for assembly of
newly synthesized A3G into cellular HMM complexes (within
30 min). Although virions were budded during the 30-min
pulse, newly synthesized Gag did not yet contribute to these
virions until the first 30-min chase period (Figure 2E, lower
panels). Thus, the recruitment of newly synthesized A3G into
virions may be intimately tied to the synthesis and assembly
of the viral genome and/or Gag (NC) and the budding of these
new virions. Indeed, A3G assembles with viral RNA in
producer cells (Figure 5 and ) and maintains this
association within virion cores (Figures 3 and 5). Since newly
synthesized A3G assembles into HMM complexes in cells
within 30 min in the presence or absence of HIV RNA (Figure
2A–2D), we cannot determine in these experiments whether
the newly synthesized A3G (less than 1.5 h old) recruited into
virions represents A3G newly assembled into any one specific
cellular complex, a viral-specific HMM complex, or a
combination of cellular and viral complexes. However,
several other observations in conjunction with these pulse-
chase data strongly support a model in which newly
synthesized cellular A3G not yet fully assembled into cellular
HMM complexes forms the major pool for recruitment into
HIVDVif virions. First, A3G incorporation into virions is
mediated by assembly with viral determinants for encapsida-
tion, including the viral RNA genome and/or Gag [15–22], and
virion-incorporated A3G ultimately forms a large RNP
complex (IVAC) with viral RNA, NC, and IN (Figures 3 and
5). Thus, virion-bound A3G forms a complex distinct from
the cellular HMM complexes, at the very least distinguished
by the presence of the viral encapsidation determinants (viral
RNA genome and/or Gag). Second, since none of the cellular
cofactors identified in the cellular HMM A3G complexes are
corecruited with A3G into virions in an A3G-specific manner
( and unpublished data), viral determinants for A3G
virion incorporation would have to extract A3G out of
mature multisubunit complexes if they do serve as a reservoir
for virion incorporation. If such a mechanism is employed, it
is difficult to explain why A3G incorporation into virions is
not also readily detected at much later time points in the
pulse-chase radiolabeling studies. Finally, overexpression of
A3G in cells leads to packaging of additional amounts of A3G
into virions that localize outside of the virion core (Figure
3C), and this form of A3G is enzymatically active in vitro in
the absence of addition of RNase A (Figure 4C). Thus, this
additional extra-core A3G appears to be the LMM monomer/
dimer that forms upon RNase A treatment  and could
possibly arise from (1) newly synthesized LMM A3G not yet
assembled into HMM complexes or (2) LMM A3G not
assembled into HMM complexes due to saturation of cellular
cofactors upon A3G overexpression.
We thus favor a model in which, upon translation, newly
synthesized LMM A3G assembles with viral RNA and protein
factors to gain access to newly assembling virions (and, in so
doing, forms an IVAC-like complex). Since viral genomic
RNA is subject to cellular processing that may be common to
RNA that nucleates the cellular HMM complexes, a subset of
common cellular RNA-binding factors can be predicted to be
found in both the cellular HMM complexes [27,28,48] and
viral RNP complexes. For example, RNA helicase A, a
component of the Staufen-containing HMM A3G complex,
has been reported to be packaged into virions , but its
incorporation occurs independently of A3G and is unaffected
by A3G virion incorporation (unpublished data). However, we
cannot completely exclude the possibility that A3G is
recruited into virions by viral cofactors from very recently
assembled HMM complex(es), as recently suggested .
However, because we do not observe virion incorporation
from older HMM A3G complexes, we do not favor such a
model. One limitation of the pulse-chase studies is that we
cannot calculate the percentage of radiolabeled (newly
synthesized) to unlabeled (mature) A3G that is in virions at
any of the given collection times.
The recruitment of A3G into HIV virions is ultimately
detrimental to the virus, underscoring the essential function
of the HIV Vif protein in blocking encapsidation of the
deaminase. The principal mechanism by which Vif abrogates
antiviral A3G activity is believed to involve proteasome-
mediated degradation of A3G, most of which is resident in
HMM A3G complexes. The observation that newly translated
A3G (less than 1.5 h old) is preferentially recruited into
virions (Figure 2B) implies that Vif must also effectively target
this newly synthesized pool of cellular A3G. Recently, it has
been reported that more Vif binds to A3G in the presence of
RNase that in its absence , suggesting that LMM A3G
unbound to RNA may be a good target for Vif. Our prior
studies have shown that Vif expression promotes polyubiqui-
tinylation of A3G that resolves as HMM . Whether Vif
activity leads to A3G ubiquitylation before, during, or after
the assembly of this enzyme into cellular HMM complexes
remains to be determined. Similarly, it remains to be
determined whether ubiquitylated A3G resolving into HMM
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150162
Biochemical Analysis of Intravirion A3G
fractions upon FPLC represents the modification of A3G in
the recently identified cellular complexes [27,28,48] or a
separate complex of A3G, Vif, Cul5, SCF, and the proteasome.
Perhaps Vif targets ribosome-associated A3G, thus destroying
newly synthesized A3G and removing the key pool of enzyme
that is selectively incorporated into virions. Such a scenario is
consistent with the observation that Vif partially inhibits the
synthesis of A3G [7,50]. Alternatively, Vif could target A3G
bound for virion incorporation by targeting viral RNA–
associated A3G. Indeed, Vif has been reported to interact
with viral genomic RNA, suggesting a mechanism by which it
might preferentially target virion-bound A3G [51–53]. Sim-
ilarly, the reported association of Vif with the plasma
membrane [54,55], the site of virion assembly, could localize
this viral protein in proximity to A3G undergoing active
encapsidation. While Vif expression ultimately depletes cells
of all A3G, others have suggested that such global degradation
of the enzyme may not be strictly required for Vif to exert its
countereffects on A3G [56,57]. Regardless, our findings
suggest that an important target for Vif is the newly
synthesized pool of A3G, rather than A3G already assembled
into cellular HMM complexes.
Because A3G can hypermutate the nascent minus-strand
DNA of HIV, we found it surprising that intravirion A3G is
inactive in in vitro deoxycytidine deaminase assays. Indeed,
we observed that the binding of HIV RNA to A3G within
virions prevented ssDNA binding and/or occluded the A3G
catalytic site(s). As shown in Figure 4A, the addition of free
ssDNA substrate to virion A3G proved insufficient to
compete RNA binding and/or access the catalytic pocket(s).
Rather, the inhibitory RNA had to be removed before A3G
enzymatic activation was observed. In view of the emerging
findings that A3G can also exert antiviral activity independ-
ent of deoxycytidine deamination [14,46,58–62], we propose
that virion A3G may employ two mechanisms acting
sequentially to produce its full antiviral effect. First, the
enzymatically latent form of A3G bound to HIV RNA may
impair the generation of minus-strand DNA by physically
blocking the movement of RT on its viral RNA template.
Indeed, short interfering RNA–mediated knockdown of
endogenous A3G in resting CD4 T cells enhances the
synthesis of late reverse transcription products  and the
generation of both early and late reverse transcription
products is reduced by the presence of A3G in virions .
However, because this inhibition is incomplete, minus-strand
viral DNA is occasionally generated, setting the stage for the
second, enzyme-dependent antiviral action of A3G. During
reverse transcription, we now show that RNase H degrades
the viral RNA that impairs A3G activity, allowing the enzyme
to extensively deaminate the minus-strand DNA. Perhaps
incomplete inhibition of reverse transcription by A3G is
caused by the occasional ability of RT displace A3G off the
RNA template. Although this could result in the generation of
enzymatically active A3G, A3G may also be able to rebind the
RNA–DNA duplex, reestablishing the inactivated state and
dependence upon RNase H for enzymatic activation. Indeed,
the lack of A3G activity induced by reverse transcription but
under conditions where RNase H activity is inhibited (Figure
6E) suggests that if A3G is displaced by RT, it rapidly rebinds
inhibitory nucleic acid. These events of initial inhibition and
subsequent activation of A3G enzymatic activity by various
components of the virus highlight an unexpectedly complex
but interesting interplay between HIV and its cellular host.
Such a dual strategy for A3G inhibition of retroviral
replication could account for its potent antiviral activity and
explain reports of both enzyme-dependent and -independent
antiviral activities. This model could also explain the
conflicting results concerning the ability of A3G to inhibit
the replication of hepatitis B virus. In some cells, A3G acts
independently of deoxycytidine deamination [46,58], while in
others, prominent DNA mutation is evident [64–66]. Cell-type
differences in the relative effectiveness of these two sequen-
tial antiviral actions of A3G could underlie these findings.
Materials and Methods
Cells and viruses. The 293T cells were maintained in DMEM
supplemented with 10% FBS (Gemini Bio-Products, http://www.
gembio.com). H9 cells were maintained in RPMI supplemented with
10% FBS. Primary CD4 T cells were isolated from fresh human
peripheral blood mononuclear cells on CD4 magnetic microbeads
(Miltenyi Biotec, http://www.miltenyibiotec.com). The isolated CD4 T
cells were then activated by 36-h treatment with PHA (5 lg/ml)
followed by 36-h IL-2 treatment (20 U/ml; Roche, http://www.roche.
com) in RPMI supplemented with 10% FBS, 100 lg/ml streptomycin,
and 100 U/ml penicillin. Virions were generated by calcium
phosphate–mediated cotransfection of subconfluent 293T in T175
flasks with a proviral plasmid (60 lg), pCMV4-HA-A3G vector (0 to 20
lg), and/or pCMV4-HA (0 to 20 lg). The medium was changed after
16 h, and the supernatant and cells collected were collected after 48
h. The virus-containing supernatant was clarified by low-speed
centrifugation, filtered through a 0.22-lm membrane, and sedi-
mented by ultracentrifugation over a 2-ml cushion of 8.4% iodixanol
at 20,000 rpm using an SW28 rotor (Beckman Coulter, http://www.
beckmancoulter.com) for 2 h at 4 8C. The virus-containing pellet was
resuspended in 1 ml of PBS, DNase-treated (RNase-free; Roche),
underlaid with a 100-ll cushion of 8.4% iodixanol and ultra-
centrifuged at 20,000 rpm in an HFA 22.1 rotor (Heraeus, http://
www.thermo.com) for 1 h at 4 8C. Unless otherwise indicated, 0.1 U of
RNase A inhibitor (RNaseOUT; Invitrogen, http://www.invitrogen.
com) was added to virion pellets, which were then immediately lysed
or flash-frozen on liquid nitrogen and stored at?80 8C until lysis. The
addition of RNaseOUT had no effect on the intrinsic activity of HA-
A3G (Figure S4). Cells were washed with PBS, and the pellet was
either immediately lysed or flash-frozen on liquid nitrogen and
stored at ?80 8C until use. To generate VSV-G–pseudotyped DVif
virions, 293T cells were cotransfected with expression vectors for the
DVif provirus and the envelope of VSV-G. At 48 h after transfection,
supernatants were cleared by low-speed centrifugation and filtration
as described above and then used directly on fresh H9 or primary
CD4 T cells. The T cells were spinoculated with the pseudotyped
virion-containing supernatant as previously described . Briefly,
0.43106cells/well of a 48-well plate were centrifuged at low speed for
2 h at room temperature with VSV-G–pseudotyped viruses. Cells were
then washed five times with cold medium and returned to complete
media for an additional 40 h. Supernatants and cells were then
collected and processed as described above for the transfected 293T
Plasmids. The proviral clone of pNL4–3DVif used to generate the
HIV-1DVif virions has been previously described [68,69]. pNL4–
3DVifH?(E478Q) contains a point mutation in the catalytic site of the
RNase H domain of RT that compromises RNase H activity. This
plasmid was generated by first subcloning the SpeI-EcoR1 Pol-
containing restriction fragment of pNL4–3DVif into pEF1A. The
mutagenesis primer 59–ACAACAAATCAGAAGACTCAGTTACAAG-
CAATTCATCTAGC–39 and its complement (Operon, http://www.
operon.eu.com) were used to generate the E478Q mutation in the
subclone using the QuikChange site-directed mutagenesis kit
(Stratagene, http://www.stratagene.com). The mutation was confirmed
by DNA sequencing. The pol region in the subclone was then
recloned back into pNL4–3DVif. The introduction of the E478Q
mutation into NL4–3DVifH–(E478Q) was confirmed by sequencing.
pCMV4-HA and pCMV4-HA-A3G  expression vectors were
cotransfected with pNL4–3DVif to generate HIV-1DVif virions
lacking or containing HA-tagged A3G.
Virion fractionations. Virion cores were obtained using a
previously published method . Briefly, virion pellets were
PLoS Pathogens | www.plospathogens.orgFebruary 2007 | Volume 3 | Issue 2 | e15 0163
Biochemical Analysis of Intravirion A3G
resuspended in MOPS Buffer I (200 mM NaCl, 100 mM MOPS [pH
7.0]) and Triton X-100 added to a final concentration of 0.5% for 2
min at room temperature. The cores were then pelleted from the
solubilized enveloped by spinning the samples at 14,000g for 8 min at
4 8C. The core pellets were then washed twice with MOPS buffer II
(100 mM NaCl, 50 mM MOPS [pH 7.0]). The cores were then either
analyzed by immunoblotting or further fractionated to remove the
p24-CA shell, as previously described . Briefly, cores were
resuspended in STE buffer (10 mM Tris [pH 6.7], 1 M NaCl, 0.5 mM
EDTA), incubated at 37 8C for 4 h and subsequently centrifuged at
14,000g to pellet the RNP complex.
FPLC analyses. Virions present in the supernatants of 293T cells
were transiently transfected with HIV proviral plasmids, and the cells
themselves were lysed in ice-cold lysis buffer (50 mM HEPES [pH 7.4],
125 mM NaCl, 0.2% NP-40, and 13 EDTA-free protease inhibitor
cocktail [Calbiochem/EMD Biosciences, http://www.emdbiosciences.
com]). Lysates were clarified by sedimentation, quantified with a
protein assay (Bio-Rad, http://www.bio-rad.com), and applied to a
calibrated Superose 6 HR 10/30 gel filtration column run by an FPLC
One column-volume (24 ml) using FPLC running buffer (50 mM
HEPES [pH 7.4], 125 mM NaCl, 0.1% NP-40, 1 mM dithiothreitol, and
10% glycerol) was collected in 1-ml aliquots. Equal volumes of
collected fractions were either directly run on SDS-PAGE gels or
concentrated with YM-3 Microcon filters with a cutoff of 3,000 Da
(Millipore, http://www.millipore.com) before running on SDS-PAGE
after normalization for resultant concentrate volume. The size-
and immunoblotted. To test nuclease sensitivity, the lysates were
U/ml DNase (RNase-free; Roche) for 1 h at 37 8C before gel filtration.
Antibodies. Polyclonal antibodies againstA3G and Vpr have
been previously described. Through the National Institutes of Health
AIDS Research and Reference Reagent Program, HIV-1 RT mono-
antiserum to HIV-1 IN (757) was obtained from Dr. Duane Grand-
by Dr. Robert J. Gorelick (National Cancer Institute, Frederick,
Maryland, United States). Mouse monoclonal anti-p24 Gag ascites
was generously provided by Beckman Coulter. Other antibodies used
include polyclonal anti-HA antibody Y11, monoclonal anti–14-3-3c
antibody C-16, monoclonal anti-CD45 antibody 2D-1, and polyclonal
anti-GFP antibody (FL) (all Santa Cruz Biotechnology, http://www.scbt.
com) and monoclonal anti-HA antibody HA.11 unlinked or linked to
beads (Covance, http://www.covance.com).
Immunoblot analysis of proteins was performed using horseradish-
linked secondary antibodies followed by ECL detection (Pierce
Biotechnology, http://www.piercenet.com). In Figure 1, A3G and
p24-CA were detected and quantified by using fluorescently linked
secondary antibodies (LI-COR Biosciences, http://www.licor.com).
Blotted proteins were then detected and quantified using the Odyssey
infrared imaging system and software (LI-COR).
Pulse-chase radiolabeling experiments. Four plates of 293T cells
were transfected with pCMV4-A3G-HA alone or with pNL4–3DVif.
After 36 h, the cells were rinsed once and incubated for 1 h with
pulse-radiolabeling medium (DMEM without methionine and cys-
teine; GIBCO, http://www.invitrogen.com) plus 10% dialysed FBS).
The cells were pulse labeled for 10 min with 500 lCi/ml EasyTag
perkinelmer.com) containing radiolabeled methionine and cysteine
in fresh pulse-radiolabeling medium. At the end of the pulse-
radiolabeling period, the radiolabel was removed and one plate of
cells harvested. The remaining radiolabeled samples were incubated
with chase medium (DMEM supplemented with 10% FBS, 4.02 mM
methionine , and 3 mM cysteine ). Cells were harvested
following incubation for 0.5, 1, or 2 h. Cells pellets were lysed in ELB
lysis buffer. Each lysate was size-fractionated on gel filtration columns
packed with Sepharose CL-6B beads, which crudely separate HMM
from LMM proteins (Figure S1). For each sample, ten fractions of 300
ll each were collected, and equal volumes of each fraction were
immunoprecipitated with anti-HA antibody. The immunoprecipi-
tates were run on SDS-PAGE, and the signal was detected by
autoradiography. The signal from each radiolabeled A3G-HA band
was quantitated using Scion Image for Windows software (Version
1.62; Scion Corporation, http://www.scioncorp.com) and divided by
the sum of the total signal, in order to assign a relative percent
density versus the fraction number for every chase time point sample.
For the pulse-chase analysis of virus-producing cells, 293T cells
were cotransfected with pNL4–3DVif, pCMV4-HA-A3G, and pEGFP-
35S Protein Labeling Mix (Perkin Elmer, http://www.
C1 to generate HA-A3G–containing HIV-1DVif virions. After 48 h,
the medium was changed, and the cells were rinsed and incubated for
1 h with pulse-radiolabeling medium as described above. The cells
were then pulse-radiolabeled for 30 min with 125 lCi/ml EasyTag
radiolabeling medium. At the end of the pulse-radiolabeling period,
the radiolabel was removed. Supernatant from the initial pulse-
labeled samples (t ¼ 0, pulse) was harvested, and radiolabeled cells
were incubated with chase medium (DMEM supplemented with 10%
FBS, 4.02 mM methionine , and 3 mM cysteine ) for 0.5 h.
Again, supernatant was collected (t ¼ 0.5 h), and the cells were
incubated with chase medium for a further 30 min to generate the t¼
1 h sample. The process was repeated twice more with an incubation
of 1 h and 2 h to generate the t ¼2 h and t¼4 h samples. At all time
points, a fraction of radiolabeled cells were also collected, washed
with PBS, pelleted by centrifugation, flash-frozen on liquid nitrogen,
and stored at?80 8C. The virus-containing supernatants were filtered
through a 0.22-lm membrane, and virions were sedimented by
ultracentrifugation over a 2-ml cushion of 8.4% iodixanol at 20,000
rpm in an SW28 rotor (Beckman) at 4 8C. The pellets were
resuspended in 1 ml of PBS, underlaid with a 100-ll cushion of
8.4% iodixanol, and ultracentrifuged at 20,000 rpm in an HFA 22.1
rotor (Hereaus) for 1 h at 4 8C. The resultant virion pellets were flash-
frozen on liquid nitrogen and stored at ?80 8C.
After virion and cell pellets had been obtained for all time points,
the samples were lysed in the lysis buffer described above. Lysates
were clarified by sedimentation and quantified with a protein assay
(Bio-Rad), and immunoprecipitations were set up at equal protein
concentration/volume in the presence of monoclonal anti-p24 ascites
or monoclonal anti-HA antibody and incubated for 2 h at 4 8C. The
immunoprecipitates were washed once with lysis buffer and subjected
to SDS-PAGE. The proteins were transferred to nitrocellulose and
immunoblotted for GFP or HA with polyclonal antibodies or for p24
with monoclonal antibody.
GFP, HA-A3G, and p24-CA identified by immunoblotting were
excised from the membranes and subjected to scintillation analysis.
Bands were first identified by immunoblotting since Gag and HA-
A3G coimmunoprecipitate with each other [15,17,20,21] and are close
in size. Scintillation counts were normalized to the amount of
immunoprecipitated material assessed, determined with ImageJ
(http://rsb.info.nih.gov/ij). The normalized counts were divided by
the sum of the total counts to assign a relative percent density for
every sample. No GFP was detected in virions (unpublished data).
In an alternate approach (Figure S2E), 293T cells were first
transfected with HA-A3G expression vector DNA using Fugene
(Roche) followed by infection of these cells with VSV-G-pseudotyped
NL4–3DVif for 12 h. The cells were then pulse-radiolabeled with 125
lCi/ml EasyTag, as described above. Also as described above, after the
pulse, cells were chased with cold medium and cells and virions were
harvested at 1, 3, 5, and 9 h after the pulse-radiolabeling period. In
these experiments, samples were subjected to denaturing lysis (50 mM
Tris [pH 7.5], 1% SDS, 5 mM dithiothreitol) followed by anti-HA or
anti-p24 immunoprecipitations (50 mM Tris [pH 7.5], 250 mM NaCl,
5 mM EDTA, 0.5% NP-40) and immunoblotting or PhosphorImaging
(Bio-Rad), as indicated.
In vitro deoxycytidine deaminase assays. Samples for analysis were
either (1) whole virion lysates or (2) FPLC fractions from cell or virion
lysates. FPLC fractions were immunoprecipitated with monoclonal
anti-HA antibody to concentrate HA-A3G. In all cases, the amount of
HA-A3G in the input samples was confirmed by immunoblotting
before analysis. DNA oligonucleotides (59-ATTATTATTATTCCCA
target sites for A3G deamination [italicized] were labeled at the 59
end with [32P]ATP using T4 polynucleotide kinase (New England
Biolabs, http://www.neb.com) or with an FITC fluorophore (Operon).
Labeled oligonucleotides and input samples were incubated in 20 ll of
h unless otherwise indicated. For incubations under conditions
stimulating endogenous reverse transcription, KCl (final concentra-
tion, 60 mM), MgCl2(final concentration, 4 mM), and dNTPs (final
concentration, 1 mM) were added. The RNase H inhibitor Compound
I, generously provided by Dr. Daria Hazuda (Merck), was used at a final
concentration of 0.1, 1, 10, or 100 lM. To terminate the reactions and
purify the labeled oligonucleotides, the reactions were subjected to G-
25 Mini Quick Spin Columns (Roche). Any uracil bases generated by
A3G were converted to abasic sites by treatment of the purified
oligonucleotides with 1 U of uracil DNA glycosylase (New England
Biolabs) for 30 min at 37 8C. After 10 min of heat inactivation at 95 8C,
the reactions were subjected to alkaline hydrolysis by the addition of
NaOH (final concentration, 0.2 M) for 10 min at 95 8C. Cleavage
35S Protein Labeling Mix (Perkin Elmer) in fresh pulse-
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150164
Biochemical Analysis of Intravirion A3G
products were resolved on 15% PAGE TBE-urea gels (Bio-Rad) and
visualized with a Personal FX Imager (Bio-Rad), for radiography or
RNase H assays. Samples for analysis were either virion lysates or
recombinant protein. Recombinant HIV-1 RT, WT or E478Q, was
generously provided by Dr. Matthias Gotte (McGill University) and
used at a final concentration of 1 nM. Test substrates included ssDNA,
ssRNA, RNA–RNA hybrid, and DNA–RNA hybrid. Unmodified 18-
mer PAGE-purified complementary DNA and RNA oligonucleotides
were from Operon and are based on the oligonucleotides 18-DAB-
DNA and 18-FAM-RNA described by Shaw-Reid et al. . In
addition, an unmodified RNA oligonucleotide complementary to
18-FAM-RNA was used to generate the RNA–RNA hybrid (Operon).
All oligonucleotides were end-labeled with [32P]ATP using polynu-
cleotide kinase (New England Biolabs). Hybrids were formed by
annealing hot oligonucleotide to cold complementary oligonucleo-
tide. Samples were incubated in 20 ll of RNase H buffer (50 mM Tris-
Cl [pH 8.0], 60 mM KCl) either with or without 5 mM MgCl2, as
indicated, for 10 min at 37 8C. Compound I was added to a final
concentration of 0.1, 1, 10, or 100 lM. Radiolabeled substrate (single-
stranded or hybrid) was added to a final concentration of 100 nM, and
the reactions were allowed to continue at 37 8C for 30 min. After the
addition of loading dye to stop the reactions, the cleavage products
were resolved on 20% PAGE-TBE-urea gels and were visualized with a
Personal FX PhosphorImager (Bio-Rad).
RT-PCR. FPLC samples and immunoprecipitates for RNA analysis
were first treated with 20 U of DNase (RNase-free; Roche) at 37 8C.
The RNA was then extracted with the QiaAmp RNA purification kit
(Qiagen) according to the manufacturer’s instructions. Viral genomic
RNA was detected by reverse transcription with a primer comple-
mentary to the gag region of HIV-1 (59-TGCTATGTCACTTCC-
CCTTGG-39, generously provided by Jerry Kropp [Gladstone
Institute of Virology and Immunology]) followed by PCR using
primers complementary to the R (F496; nucleotides 496–517) and U5
(R573; nucleotides 552–573) or U5 (F592; nucleotides 592–613) and
PBS (R666; nucleotides 645–666) regions of HIV-1. All these primers
have been described . In addition, reverse transcription was also
performed using an antisense primer complementary to the Vpu
region (59-TCATTGCCACTGTCTTCTGCTCT-39) followed by PCR
using the Vpr primer and a primer complementary to Pol (59-GT-
Figure S1. Analysis of the Purity of the Virions Generated
(A) The 293T cells expressing DVif provirus with (þ) or without (?)
HA-A3G, and virions derived from these cells, were assessed by
immunoblotting (IB) for an abundant cellular factor 14-3-3c that is
not incorporated into virions . This factor was readily detected in
the producer cell lysates but not in the virion lysates.
(B) Activated primary CD4 T cells derived from peripheral blood of
two donors were spinoculated with VSV-G–pseudotyped NL4–3DVif.
Both the infected cells and virions derived thereof were assessed by
immunoblotting (IB) for the presence of CD45 (molecular weight
180–220 kDa), which is highly expressed on miscrovesicles but
excluded from virions .
Found at doi:10.1371/journal.ppat.0030015.sg001 (452 KB AI).
Figure S2. Additional Pulse-Chase Data
(A) Gel filtration of cell lysates by mini Sepharose CL-6B columns
distinguishes HMM A3G from LMM A3G. The 293T cells transiently
expressing A3G-HA or HA-A3G were lysed and subjected to CL-6B
size-fractionation. The ten fractions collected were assessed by
immunoblotting (IB) to determine the resolution of A3G and
monomeric 55-kDa a-tubulin. Shown are eight representative gel
filtrations. HMM A3G, which peaks in fractions 4 and 5, is readily
distinguishable from LMM a-tubulin, which peaks in fractions 6 and 7.
(B) Pulse labeling for 30 min resolves newly synthesized A3G as HMM
within the 30-min pulse period and persists as HMM for at least 3 h.
Chase lysates were similarly subjected to gel filtration as in (A). Shown
are the autoradiograms of the anti-HA immunoprecipitates from
each fraction after gel filtration.
(C) Normalization of radiolabeled HA-A3G incorporated into virions
(as originally plotted in Figure 2E, middle panel), by the amount of
radioactive HA-A3G available in the producer cell lysates (Figure 2E,
top panel). Plotted is the percent radioactive density of thus-
normalized A3G for any given time point relative to the total
radioactive density of all the time points. P, pulse.
(D) The decline in radiolabel content over the chase is similar for HA-
A3G in the absence or presence of DVif proviral gene expression. The
293T cells in a six-well plate were transfected with HA-A3G and GFP
plasmids but in the absence of provirus, in parallel with the cells in
Experiment 1 of Figure 2E. The pulse chase was performed as in
Figure 2E, in parallel with the provirus-expressing samples. Plotted is
the percent density of radiolabeled HA-A3G relative to the whole
(E) Extension of the chase time reveals that even 9-h-old HMM A3G is
not recruited into virions. The 293T cells transfected with HA-A3G
were infected with VSV-G–pseudotyped HIVDVif, pulse labeled, and
then chased with cold medium for the following 9 h. HA-A3G and
p24-CA were immunoprecipitated and subjected to either immuno-
blotting (IB) with antibodies to either A3G or p24-CA or Phosphor-
Imager analysis, as indicated. P, pulse.
Found at doi:10.1371/journal.ppat.0030015.sg002 (4.9 MB AI).
Figure S3. Treatment of Virion Lysates with EDTA Activates A3G
Virions bearing a WT or mutant RNase H domain were lysed in lysis
buffer supplemented with 10 mM EDTA and assessed in the
deoxycytidine deaminase assay. Virions contained or lacked HA-
A3G as indicated.
Found at doi:10.1371/journal.ppat.0030015.sg003 (964 KB AI).
Figure S4. A3G Enzymatic Activity Is Unaffected by RNaseOUT
HA-A3G from RNase A–treated virion lysates was assessed in the in
vitro deoxycytidine deaminase assay with increasing doses of
RNaseOUT (0.2, 4, and 40 U, as indicated by the slope of the triangle)
to assess whether the inhibitor affects intrinsic A3G enzymatic
Found at doi:10.1371/journal.ppat.0030015.sg004 (1.4 MB AI).
The authors would like to particularly thank Dr. Daria Hazuda
(Merck) for generously providing Compound I, Dr. Matthias Go ¨tte
(McGill University) for generously providing the two recombinant
reverse transcriptase enzymes, and Jerry Kropp (Gladstone Institute
of Virology and Immunology) for generously providing the reverse
transcription primer. Special thanks are given to Y. L. Chiu, J.
Kreisberg, K. Stopak, M. Cavrois, and A. O’Mahony for helpful
discussions and/or assistance, S. Ordway for editorial assistance, J.
Carroll for graphics support, and R. Givens and S. Cammack for
Author contributions. VBS, WY, and WCG conceived and designed
the experiments. VBS and WY performed the experiments, analyzed
the data, and contributed reagents/materials/analysis tools. VBS and
WCG wrote the paper.
Funding. We are grateful for funding support including National
Institutes of Health (NIH) grants R01AI065329 and P01 HD40543 to
WCG and funding from the University of California San Francisco–
Gladstone Institute of Virology and Immunology Center for AIDS
Research (UCSF-GIVI CFAR) (P30 A127763). The project was also
supported in part by an NIH core equipment grant awarded to the J.
David Gladstone Institutes (RR1 892801). VBS is supported by a
fellowship from the Universitywide AIDS Research Program.
Competing interests. The authors have declared that no competing
1.Sheehy AM, Gaddis NC, Choi JD, Malim MH (2002) Isolation of a human
gene that inhibits HIV-1 infection and is suppressed by the viral Vif
protein. Nature 418: 646–650.
2. Harris RS, Bishop KN, Sheehy AM, Craig HM, Petersen-Mahrt SK, et al.
(2003) DNA deamination mediates innate immunity to retroviral infection.
Cell 113: 803–809.
3. Mangeat B, Turelli P, Caron G, Friedli M, Perrin L, et al. (2003) Broad
antiretroviral defence by human APOBEC3G through lethal editing of
nascent reverse transcripts. Nature 424: 99–103.
Lecossier D, Bouchonnet F, Clavel F, Hance AJ (2003) Hypermutation of
HIV-1 DNA in the absence of the Vif protein. Science 300: 1112.
Zhang H, Yang B, Pomerantz RJ, Zhang C, Arunachalam SC, et al. (2003)
The cytidine deaminase CEM15 induces hypermutation in newly synthe-
sized HIV-1 DNA. Nature 424: 94–98.
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150165
Biochemical Analysis of Intravirion A3G
6. Yu Q, Konig R, Pillai S, Chiles K, Kearney M, et al. (2004) Single-strand
specificity of APOBEC3G accounts for minus-strand deamination of the
HIV genome. Nat Struct Mol Biol 11: 435–442.
Stopak K, de Noronha C, Yonemoto W, Greene WC (2003) HIV-1 Vif blocks
the antiviral activity of APOBEC3G by impairing both its translation and
intracellular stability. Mol Cell 12: 591–601.
Marin M, Rose KM, Kozak SL, Kabat D (2003) HIV-1 Vif protein binds the
editing enzyme APOBEC3G and induces its degradation. Nat Med 9: 1398–
Sheehy AM, Gaddis NC, Malim MH (2003) The antiretroviral enzyme
APOBEC3G is degraded by the proteasome in response to HIV-1 Vif. Nat
Med 9: 1404–1407.
10. Conticello SG, Harris RS, Neuberger MS (2003) The Vif protein of HIV
triggers degradation of the human antiretroviral DNA deaminase
APOBEC3G. Curr Biol 13: 2009–2013.
11. Mehle A, Strack B, Ancuta P, Zhang C, McPike M, et al. (2004) Vif
overcomes the innate antiviral activity of APOBEC3G by promoting its
degradation in the ubiquitin-proteasome pathway. J Biol Chem 279: 7792–
12. Yu X, Yu Y, Liu B, Luo K, Kong W, et al. (2003) Induction of APOBEC3G
ubiquitination and degradation by an HIV-1 Vif-Cul5-SCF complex.
Science 302: 1056–1060.
13. Mariani R, Chen D, Schrofelbauer B, Navarro F, Konig R, et al. (2003)
Species-specific exclusion of APOBEC3G from HIV-1 virions by Vif. Cell
14. Chiu YL, Soros VB, Kreisberg JF, Stopak K, Yonemoto W, et al. (2005)
Cellular APOBEC3G restricts HIV-1 infection in resting CD4þT cells.
Nature 435: 108–114.
15. Cen S, Guo F, Niu M, Saadatmand J, Deflassieux J, et al. (2004) The
interaction between HIV-1 Gag and APOBEC3G. J Biol Chem 279: 33177–
16. Svarovskaia ES, Xu H, Mbisa JL, Barr R, Gorelick RJ, et al. (2004) Human
apolipoprotein B mRNA-editing enzyme-catalytic polypeptide-like 3G
(APOBEC3G) is incorporated into HIV-1 virions through interactions with
viral and nonviral RNAs. J Biol Chem 279: 35822–35828.
17. Luo K, Liu B, Xiao Z, Yu Y, Yu X, et al. (2004) Amino-terminal region of the
human immunodeficiency virus type 1 nucleocapsid is required for human
APOBEC3G packaging. J Virol 78: 11841–11852.
18. Khan MA, Kao S, Miyagi E, Takeuchi H, Goila-Gaur R, et al. (2005) Viral
RNA is required for the association of APOBEC3G with human
immunodeficiency virus type 1 nucleoprotein complexes. J Virol 79:
19. Schafer A, Bogerd HP, Cullen BR (2004) Specific packaging of APOBEC3G
into HIV-1 virions is mediated by the nucleocapsid domain of the gag
polyprotein precursor. Virology 328: 163–168.
20. Alce TM, Popik W (2004) APOBEC3G is incorporated into virus-like
particles by a direct interaction with HIV-1 Gag nucleocapsid protein. J
Biol Chem 279: 34083–34086.
21. Douaisi M, Dussart S, Courcoul M, Bessou G, Vigne R, et al. (2004) HIV-1
and MLV Gag proteins are sufficient to recruit APOBEC3G into virus-like
particles. Biochem Biophys Res Commun 321: 566–573.
22. Zennou V, Perez-Caballero D, Gottlinger H, Bieniasz PD (2004) APOBEC3G
incorporation into human immunodeficiency virus type 1 particles. J Virol
23. Dutko JA, Schafer A, Kenny AE, Cullen BR, Curcio MJ (2005) Inhibition of a
yeast LTR retrotransposon by human APOBEC3 cytidine deaminases. Curr
Biol 15: 661–666.
24. Suspene R, Sommer P, Henry M, Ferris S, Guetard D, et al. (2004)
APOBEC3G is a single-stranded DNA cytidine deaminase and functions
independently of HIV reverse transcriptase. Nucleic Acids Res 32: 2421–
25. Schrofelbauer B, Yu Q, Zeitlin SG, Landau NR (2005) Human immunode-
ficiency virus type 1 Vpr induces the degradation of the UNG and SMUG
uracil-DNA glycosylases. J Virol 79: 10978–10987.
26. Kaiser SM, Emerman M (2006) Uracil DNA glycosylase is dispensable for
human immunodeficiency virus type 1 replication and does not contribute
to the antiviral effects of the cytidine deaminase APOBEC3G. J Virol 80:
27. Chiu YL, Witkowska HE, Hall SC, Santiago M, Soros VB, et al. (2006) High-
molecular-mass APOBEC3G complexes restrict Alu retrotransposition.
Proc Natl Acad Sci U S A 103: 15588–15593.
28. Kozak SL, Marin M, Rose KM, Bystrom C, Kabat D (2006) The anti-HIV-1
editing enzyme APOBEC3G binds HIV-1 RNA and messenger RNAs that
shuttle between polysomes and stress granules. J Biol Chem 281: 29105–
29. Briggs JA, Wilk T, Welker R, Krausslich HG, Fuller SD (2003) Structural
organization of authentic, mature HIV-1 virions and cores. EMBO J 22:
30. Briggs JA, Simon MN, Gross I, Krausslich HG, Fuller SD, et al. (2004) The
stoichiometry of Gag protein in HIV-1. Nat Struct Mol Biol 11: 672–675.
31. Park J, Morrow CD (1993) Mutations in the protease gene of human
immunodeficiency virus type 1 affect release and stability of virus particles.
Virology 194: 843–850.
32. Chassagne J, Verrelle P, Dionet C, Clavel F, Barre-Sinoussi F, et al. (1986) A
monoclonal antibody against LAV gag precursor: Use for viral protein
analysis and antigenic expression in infected cells. J Immunol 136: 1442–
33. Forshey BM, Aiken C (2003) Disassembly of human immunodeficiency virus
type 1 cores in vitro reveals association of Nef with the subviral
ribonucleoprotein complex. J Virol 77: 4409–4414.
34. Forshey BM, von Schwedler U, Sundquist WI, Aiken C (2002) Formation of
a human immunodeficiency virus type 1 core of optimal stability is crucial
for viral replication. J Virol 76: 5667–5677.
35. Fassati A, Goff SP (2001) Characterization of intracellular reverse
transcription complexes of human immunodeficiency virus type 1. J Virol
36. Blobel G (1971) Isolation of a 5S RNA-protein complex from mammalian
ribosomes. Proc Natl Acad Sci U S A 68: 1881–1885.
37. Stefani G, Fraser CE, Darnell JC, Darnell RB (2004) Fragile X mental
retardation protein is associated with translating polyribosomes in neuro-
nal cells. J Neurosci 24: 7272–7276.
38. Chelico L, Pham P, Calabrese P, Goodman MF (2006) APOBEC3G DNA
deaminase acts processively 3’ ! 5’ on single-stranded DNA. Nat Struct
Mol Biol 13: 392–399.
39. DiMarzo SJ, Rakoff JS (1986) Intrauterine insemination with husband’s
washed sperm. Fertil Steril 46: 470–475.
40. Lightfoote MM, Coligan JE, Folks TM, Fauci AS, Martin MA, et al. (1986)
Structural characterization of reverse transcriptase and endonuclease
polypeptides of the acquired immunodeficiency syndrome retrovirus. J
Virol 60: 771–775.
41. Cirino NM, Cameron CE, Smith JS, Rausch JW, Roth MJ, et al. (1995)
Divalent cation modulation of the ribonuclease functions of human
immunodeficiency virus reverse transcriptase. Biochemistry 34: 9936–9943.
42. Shaw-Reid CA, Feuston B, Munshi V, Getty K, Krueger J, et al. (2005)
Dissecting the effects of DNA polymerase and ribonuclease H inhibitor
combinations on HIV-1 reverse-transcriptase activities. Biochemistry 44:
43. Shaw-Reid CA, Munshi V, Graham P, Wolfe A, Witmer M, et al. (2003)
Inhibition of HIV-1 ribonuclease H by a novel diketo acid, 4-[5-
(benzoylamino)thien-2-yl]-2,4-dioxobutanoic acid. J Biol Chem 278: 2777–
44. Schatz O, Cromme FV, Gruninger-Leitch F, Le Grice SF (1989) Point
mutations in conserved amino acid residues within the C-terminal domain
of HIV-1 reverse transcriptase specifically repress RNase H function. FEBS
Lett 257: 311–314.
45. Russell RA, Wiegand HL, Moore MD, Schafer A, McClure MO, et al. (2005)
Foamy virus Bet proteins function as novel inhibitors of the APOBEC3
family of innate antiretroviral defense factors. J Virol 79: 8724–8731.
46. Turelli P, Mangeat B, Jost S, Vianin S, Trono D (2004) Inhibition of
hepatitis B virus replication by APOBEC3G. Science 303: 1829.
47. Doehle BP, Schafer A, Wiegand HL, Bogerd HP, Cullen BR (2005)
Differential sensitivity of murine leukemia virus to APOBEC3-mediated
inhibition is governed by virion exclusion. J Virol 79: 8201–8207.
48. Wichroski MJ, Robb GB, Rana TM (2006) Human retroviral host restriction
factors APOBEC3G and APOBEC3F localize to mRNA processing bodies.
PLoS Pathog 2: e41. doi:10.1371/journal.ppat.0020041
49. Roy BB, Hu J, Guo X, Russell RS, Guo F, et al. (2006) Association of RNA
helicase a with human immunodeficiency virus type 1 particles. J Biol Chem
50. Kao S, Khan MA, Miyagi E, Plishka R, Buckler-White A, et al. (2003) The
human immunodeficiency virus type 1 Vif protein reduces intracellular
expression and inhibits packaging of APOBEC3G (CEM15), a cellular
inhibitor of virus infectivity. J Virol 77: 11398–11407.
51. Henriet S, Richer D, Bernacchi S, Decroly E, Vigne R, et al. (2005)
Cooperative and specific binding of Vif to the 5’ region of HIV-1 genomic
RNA. J Mol Biol 354: 55–72.
52. Zhang H, Pomerantz RJ, Dornadula G, Sun Y (2000) Human immunode-
ficiency virus type 1 Vif protein is an integral component of an mRNP
complex of viral RNA and could be involved in the viral RNA folding and
packaging process. J Virol 74: 8252–8261.
53. Khan MA, Aberham C, Kao S, Akari H, Gorelick R, et al. (2001) Human
immunodeficiency virus type 1 Vif protein is packaged into the
nucleoprotein complex through an interaction with viral genomic RNA. J
Virol 75: 7252–7265.
54. Simon JH, Fouchier RA, Southerling TE, Guerra CB, Grant CK, et al. (1997)
The Vif and Gag proteins of human immunodeficiency virus type 1
colocalize in infected human T cells. J Virol 71: 5259–5267.
55. Goncalves J, Shi B, Yang X, Gabuzda D (1995) Biological activity of human
immunodeficiency virus type 1 Vif requires membrane targeting by C-
terminal basic domains. J Virol 69: 7196–7204.
56. Kao S, Miyagi E, Khan MA, Takeuchi H, Opi S, et al. (2004) Production of
infectious human immunodeficiency virus type 1 does not require
depletion of APOBEC3G from virus-producing cells. Retrovirology 1: 27.
57. Mehle A, Goncalves J, Santa-Marta M, McPike M, Gabuzda D (2004)
Phosphorylation of a novel SOCS-box regulates assembly of the HIV-1 Vif-
Cul5 complex that promotes APOBEC3G degradation. Genes Dev 18: 2861–
58. Rosler C, Kock J, Kann M, Malim MH, Blum HE, et al. (2005) APOBEC-
mediated interference with hepadnavirus production. Hepatology 42: 301–
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150166
Biochemical Analysis of Intravirion A3G
59. Sasada A, Takaori-Kondo A, Shirakawa K, Kobayashi M, Abudu A, et al. Download full-text
(2005) APOBEC3G targets human T-cell leukemia virus type 1. Retrovir-
ology 2: 32.
60. Newman EN, Holmes RK, Craig HM, Klein KC, Lingappa JR, et al. (2005)
Antiviral function of APOBEC3G can be dissociated from cytidine
deaminase activity. Curr Biol 15: 166–170.
61. Navarro F, Bollman B, Chen H, Konig R, Yu Q, et al. (2005) Complementary
function of the two catalytic domains of APOBEC3G. Virology 333: 374–
62. Stenglein MD, Harris RS (2006) APOBEC3B and APOBEC3F inhibit L1
retrotransposition by a DNA deamination-independent mechanism. J Biol
Chem 281: 16839–16841.
63. Guo F, Cen S, Niu M, Saadatmand J, Kleiman L (2006) Inhibition of
formula-primed reverse transcription by human APOBEC3G during
human immunodeficiency virus type 1 replication. J Virol 80: 11710–11722.
64. Noguchi C, Ishino H, Tsuge M, Fujimoto Y, Imamura M, et al. (2005) G to A
hypermutation of hepatitis B virus. Hepatology 41: 626–633.
65. Rosler C, Kock J, Malim MH, Blum HE, von Weizsacker F (2004) Comment
on ‘‘Inhibition of hepatitis B virus replication by APOBEC3G.’’ Science
305: 1403; author reply 1403.
66. Suspene R, Guetard D, Henry M, Sommer P, Wain-Hobson S, et al. (2005)
Extensive editing of both hepatitis B virus DNA strands by APOBEC3
cytidine deaminases in vitro and in vivo. Proc Natl Acad Sci U S A 102:
67. O’Doherty U, Swiggard WJ, Malim MH (2000) Human immunodeficiency
virus type 1 spinoculation enhances infection through virus binding. J Virol
68. Adachi A, Ono N, Sakai H, Ogawa K, Shibata R, et al. (1991) Generation and
characterization of the human immunodeficiency virus type 1 mutants.
Arch Virol 117: 45–58.
69. Sakai H, Shibata R, Sakuragi J, Sakuragi S, Kawamura M, et al. (1993) Cell-
dependent requirement of human immunodeficiency virus type 1 Vif
protein for maturation of virus particles. J Virol 67: 1663–1666.
70. Jenkins Y, McEntee M, Weis K, Greene WC (1998) Characterization of HIV-
1 vpr nuclear import: Analysis of signals and pathways. J Cell Biol 143: 875–
71. von Schwedler UK, Stuchell M, Muller B, Ward DM, Chung HY, et al. (2003)
The protein network of HIV budding. Cell 114: 701–713.
72. Esser MT, Graham DR, Coren LV, Trubey CM, Bess JW Jr, et al. (2001)
Differential incorporation of CD45, CD80 (B7–1), CD86 (B7–2), and major
histocompatibility complex class I and II molecules into human immuno-
deficiency virus type 1 virions and microvesicles: Implications for viral
pathogenesis and immune regulation. J Virol 75: 6173–6182.
PLoS Pathogens | www.plospathogens.org February 2007 | Volume 3 | Issue 2 | e150167
Biochemical Analysis of Intravirion A3G