JOURNAL OF VIROLOGY, Nov. 2005, p. 13538–13547
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
Vol. 79, No. 21
?-Adrenoreceptors Reactivate Kaposi’s Sarcoma-Associated
Herpesvirus Lytic Replication via PKA-Dependent
Control of Viral RTA
Margaret Chang,1Helen J. Brown,2Alicia Collado-Hidalgo,3,6Jesusa M. Arevalo,3
Zoran Galic,3Tonia L. Symensma,2,4Lena Tanaka,4Hongyu Deng,2
Jerome A. Zack,1,3,5Ren Sun,2,5and Steve W. Cole3,5,6*
Departments of Microbiology, Immunology and Molecular Genetics,1Molecular and Medical Pharmacology,2and
Medicine,3David E. Geffen School of Medicine, University of California, Los Angeles, California; Department
of Biological Sciences, Mount Saint Mary’s College, Los Angeles, California4; and Jonsson Comprehensive Cancer Center,5
UCLA AIDS Institute, and UCLA Molecular Biology Institute5and Norman Cousins Center,6
University of California, Los Angeles, California
Received 25 April 2005/Accepted 5 August 2005
Reactivation of Kaposi’s sarcoma-associated herpesvirus (KSHV) lytic replication is mediated by the viral
RTA transcription factor, but little is known about the physiological processes controlling its expression or
activity. Links between autonomic nervous system activity and AIDS-associated Kaposi’s sarcoma led us to
examine the potential influence of catecholamine neurotransmitters. Physiological concentrations of epineph-
rine and norepinephrine efficiently reactivated lytic replication of KSHV in latently infected primary effusion
lymphoma cells via ?-adrenergic activation of the cellular cyclic AMP/protein kinase A (PKA) signaling
pathway. Effects were blocked by PKA antagonists and mimicked by pharmacological and physiological PKA
activators (prostaglandin E2and histamine) or overexpression of the PKA catalytic subunit. PKA up-regulated
RTA gene expression, enhanced activity of the RTA promoter, and posttranslationally enhanced RTA’s trans-
activating capacity for its own promoter and heterologous lytic promoters (e.g., the viral PAN gene). Mutation
of predicted phosphorylation targets at RTA serines 525 and 526 inhibited PKA-mediated enhancement of RTA
trans-activating capacity. Given the high catecholamine levels at sites of KSHV latency such as the vasculature
and lymphoid organs, these data suggest that ?-adrenergic control of RTA might constitute a significant
physiological regulator of KSHV lytic replication. These findings also suggest novel therapeutic strategies for
controlling the activity of this oncogenic gammaherpesvirus in vivo.
known as human herpesvirus 8 [HHV-8]) is a lymphotropic
gammaherpesvirus originally identified in the context of Ka-
posi’s sarcoma (8) and subsequently implicated in primary
effusion lymphoma and multicentric Castleman’s disease (6,
66). The KSHV genome most closely resembles herpesvirus
saimiri (60), but the virus also bears more distant structural
and functional similarities to Epstein-Barr virus (48, 49). Like
all herpesviruses, KSHV displays a bifurcating gene expression
program that allows it to defer lytic replication and enter a
protracted state of latency during which only a small minority
of viral genes are expressed (19, 21, 71, 78). KSHV establishes
latency in B lymphocytes and vascular endothelial cells, but it
must resume lytic replication to disseminate or colonize a new
host. Discovery of the physiological signals that control KSHV
reactivation is thus key to controlling its pathogenic potential.
Suppression of such signals could block the spread of infection,
and pharmacological induction of such signals might flush la-
tently infected cells into lytic replication for elimination by
nucleoside analogue drugs (31).
Impaired cellular immunity plays a critical role in allowing
KSHV replication, but the physiological stimuli that positively
induce lytic gene expression are poorly understood. Chemical
agents such as phorbol esters or N-butyrate can reactivate
KSHV in vitro (45, 50, 57), and proinflammatory cytokines
have similar, though weaker effects (7, 44, 46). At the level of
the viral genome, lytic reactivation is mediated by the KSHV-
encoded transcription factor RTA. Overexpression of the RTA
gene is sufficient to trigger lytic replication in latently infected
B-cell lines (37, 70), and RTA mutations can block viral reac-
tivation in vitro (36). RTA protein can also autoactivate the
RTA promoter (16, 25, 62), but the cellular signals that initiate
RTA expression are not well understood.
Several recent studies have shown that high levels of auto-
nomic nervous system activity can accelerate the onset of
AIDS-defining conditions during human immunodeficiency vi-
rus type 1 infection (11, 12, 14). These effects have been at-
tributed to autonomic nervous system regulation of human
immunodeficiency virus type 1 replication (10, 13, 14), but it is
also conceivable that autonomic nervous system activity might
directly activate opportunistic pathogens such as KSHV. The
present studies examined that hypothesis, with an emphasis on
the cellular signal transduction pathways that might allow cat-
echolamine neurotransmitters from the autonomic nervous
system to regulate key molecular events in KSHV reactivation.
Results show that physiological concentrations of epinephrine
and norepinephrine can induce lytic replication of KSHV in
* Corresponding author. Mailing address: Department of Medicine,
Division of Hematology-Oncology, 11-934, Factor Building, David
Geffen School of Medicine at UCLA, Los Angeles, CA 90095-1678.
Phone: 310-267-4243. Fax: 310-206-1318. E-mail: email@example.com.
latently infected lymphoid cells via ?-adrenergic activation of
the cellular protein kinase A (PKA) signaling pathway. These
effects are mediated by increased expression of RTA and post-
translational enhancement of its trans-activating capacity. Re-
sults provide an endocrinological perspective on the control of
KSHV replication and suggest novel strategies for therapeutic
MATERIALS AND METHODS
Cell culture. KSHV reactivation was analyzed in the primary effusion lym-
phoma (PEL) cell lines KS-1, BC3, and BCBL-1. DG75 and Ramos served as B
lymphoid cells uninfected with KSHV or Epstein-Barr virus. Cells were cultured
under standard conditions in RPMI plus 10% fetal bovine serum or in serum-
free X-VIVO 15 (Cambrex, East Rutherford, NJ) supplemented by a defined mix
of growth factors (MITO?, BD Biosciences, Bedford, MA). Reactivation exper-
iments were performed as described (70), using the PKC inducer phorbol-12-
myristate-13-acetate (PMA, also known as TPA; Calbiochem, San Diego CA),
the PKA inducer dibutyrl cyclic AMP (db-cAMP), norepinephrine ([?]-artere-
nol; Sigma-Aldrich, St. Louis, MO), or other indicated agents acting on the
?-adrenoreceptor (?AR)/cAMP/PKA signaling pathway (all from Sigma-Aldrich
or Calbiochem). Norepinephrine and PKA modulating agents were diluted in
phosphate-buffered saline and added to cultures in volumes ?1% of total culture
volume. Unstimulated controls received an equivalent volume of vehicle.
Lytic protein expression. Intracellular expression of KSHV ORF59 was as-
sayed by flow cytometric quantification of indirect immunofluorescence in per-
meabilized KS1, BC3, or BCBL1 cells; 2 ? 106cells were stained for 30 min at
4°C with 1 ?g of mouse-anti KSHV ORF59 monoclonal antibody (Advanced
Biotechnology Incorporated, Colombia, MD) in 50 ?l of BD Cytofix/Cytoperm
(BD Immunocytometry Systems, San Jose, CA), washed in 450 ?l of permeabi-
lization buffer, stained for 30 min with 0.5 ?g of fluorescein isothiocyanate-
conjugated rat anti-mouse immunoglobulin G2b Ab (BD Immunocytometry
Systems), and washed again in 450 ?l of permeabilization buffer. Fluorescence
intensity data were acquired on a FACScan flow cytometer (BD Immunocytom-
etry) with dead cells and debris excluded on the basis of forward- versus side-
scatter gating using CellQuest software. KSHV lytic proteins and cellular ?-actin
were also assayed by Western blot with enhanced chemiluminescence using
KSHV patient serum, as previously described (70).
Lytic gene expression. mRNA for KSHV gene products and human ?-adre-
noreceptor subtypes were quantified relative to cellular housekeeping genes
using real-time reverse transcription (RT)-PCR. Reactions utilized 1/10 of the
total DNase-treated (QIAGEN, Valencia, CA) RNA extracted from 106cells in
a one-step thermal cycling protocol (QIAGEN One-Step RT-PCR) with 30 min
of reverse transcription at 50°C, 15 min. of RT denaturation at 95°C, and 40
cycles of DNA amplification (15 s at 95°C, 60 s at 60°C). Reactions utilized
established primers and fluorescent detection probes for human glyceraldehyde-
3-phosphate dehydrogenase (GAPDH) and KSHV K8.1, ORF29, ORF50,
ORF72, and ORF57 (21). Primers for human ?-adrenoreceptor subtypes were:
?1(forward: TCG GAA TCC AAG GTG TAG GG, reverse:TGG CTT TTC
TCT TTG CCT CG), ?2(forward: CAT GTC TCT CAT CGT CCT GGC CA,
reverse:CAC GAT GGA AGA GGC AAT GGC A), and ?3(forward: GGC
TTC TTG GGG AGT TTC TTA GG, reverse: TTC TGG AGG GTA GAG
TGT CAC AGC), derived from GenBank sequences ADRB1: J03019, ADRB2:
M15169, and ADRB3: X70811, respectively. mRNA expression was normalized
to GAPDH by subtraction of threshold cycles (Ct; normalized target Ct? target
Ct? GAPDH Ct) and quantified as a fold change relative to an unstimulated
baseline (fold change ? 2[stimulated normalized target Ct– baseline normalized
DNA replication. Replicating KSHV DNA was assayed by Gardella gel anal-
ysis as previously described (23). Briefly, 2 ? 107KS1 cells were lysed during
electrophoresis through a 0.75% agarose gel for 3 h at 0.8 V/cm followed by 17 h
at 4.5 V/cm. Resolved DNA was transferred to a Hybond N? membrane (Am-
ersham-Pharmacia, Buckinghamshire England), UV cross-linked, and probed
with a 3-kb32P-labeled PCR product spanning the majority of the KSHV ORF50
locus. Supernatant particle-associated KSHV DNA was assayed by treating 1 ml
of 0.45-?m-filtered PEL cell supernatants with 2 ?g DNase (Worthington Bio-
chemical Corporation, Lakewood, NJ) and 10 mM MgCl for 15 min, followed by
extraction of particle-protected DNA (QIAGEN MiniElute virus spin kit) and
real-time PCR amplification of KSHV K8.1 DNA (45 cycles of 95°C for 15
seconds and 60°C for 60 seconds) with resolution of products on an 3% agarose
Infectious virus. PEL cell production of infectious KSHV was assayed by
suspending peripheral blood mononuclear cells (PBMC) from healthy donors in
PEL cell supernatants that had been filtered at 0.45 ?m and treated with DNase
as described above; 2 ? 106PBMC were stimulated with PHA for 72 h, washed,
and incubated for 1 h. in 2 ml of cell culture supernatant from PEL cells treated
with phorbol myristate acetate (PMA), norepinephrine, db-cAMP, or vehicle for
48 h. After extensive washing, PBMC were cultured for another 24 h and
establishment of KSHV genomic DNA in PBMC was assayed by PCR detection
of ORF50 (forward: AAC CAG AAG CCT CGG GCG AAG, reverse: GTG
CAC GCC ACG GAT GTC) or K8.1 (forward: AAA GCG TCC AGG CCA
CCA CAG A, reverse: GGC AGA AAA TGG CAC ACG GTT AC) in cellular
DNA (45 cycles of 95°C for 15 seconds and 60°C for 60 seconds, with SYBR
green real-time detection). Cellular RNA was also extracted from 8 ? 106
PBMC, treated with RNase-free DNase (QIAGEN RNEasy), and assayed for
expression of the latent gene product ORF72 by real-time RT-PCR as described
(21). Results were normalized to GAPDH DNA amplified in parallel and visu-
alized on a 2% agarose gel.
Overexpression of PKA catalytic subunit. PEL cells were transduced with a
self-inactivating lentiviral vector expressing a constitutively active form of PKA
(43) and an enhanced green fluorescent protein (EGFP) reporter gene under
control of a recombinant Rh-MLV promoter (34). Both sequences were trans-
lated from a single transcript bearing the PS3 internal ribosome entry site (IRES)
(73). cDNA for the human immunodeficiency virus type 1 central polypyrimidine
tract (77) was introduced into pSIN-18-Rh (34) at the XhoI site upstream of the
Rh murine leukemia virus promoter to produce pSIN18RhMLV-E-CPPT.
cDNA of the PKA catalytic subunit ? (PRKACA: X07767) was amplified with
primers bearing AgeI and SalI restriction sites and subcloned into pCR-Blunt
TOPO (Invitrogen, Carlsbad, CA), sequenced, released by digestion, purified,
and subcloned into AgeI and SalI sites of pSIN18RhMLV-E-CPPT to produce
pSIN18RhMLV-E-CPPT-PKA. The PS3 IRES-EGFP sequence was excised
from pDF-PS3 (73) and subcloned into the NotI site of a circularized pCRII-
TOPO vector (Invitrogen). The EcoRV site upstream of the subcloned fragment
was changed into a SalI site, and a SalI/XhoI fragment was subcloned into
pSIN18RhMLV-E-CPPT-PKA to produce pSIN18RhMLV-E-CPPT-PKA-PS3-
EGFP. The negative control vector pSIN18RhMLV-E-CPPT-EGFP included
EGFP sequences in the absence of upstream PRKACA-PS3 sequences. Vectors
were constructed by transfection into 293T cells (34), and target cells were
transduced by 1 h of incubation in vector-containing supernatants supplemented
with 10 ?g/ml Polybrene.
RTA promoter activity. Luciferase reporter assays utilized a 3-kb sequence
upstream of the RTA translation start site (pRpluc) (16), and three truncation
variants generated by restriction enzyme digestion of pRpluc (pRp1 cut at PstI,
pRp2 at NdeI, and pRp8 at KpnI) to delete six potential cAMP response
elements (CREs) detected by sequence-based bioinformatics (76); 4 ?g of each
reporter construct was electroporated along with 50 ng of the control pRLCMV
(Renilla luciferase driven by the cytomegalovirus [CMV] promoter) (Promega,
Madison, WI) and 6 ?g of empty pcDNA3 vector (10 ?g total) into 107KS-1,
BC3, BCBL-1, or DG75 cells (240 V, 125 ?, 950 ?F) or 5 ? 107Ramos cells
supplemented with 26 ?g/ml DEAE dextran (2 cycles of 320 V, 0 ?, 950 ?F),
using a BTX ECM 630 pulse generator. Dual luciferase assays (Promega) were
performed in triplicate, with firefly luciferase light units normalized to Renilla
luciferase light units prior to analysis of log fold change by Student’s t test.
RTA trans-activating capacity. RTA protein was expressed in the context of
RTA promoter assays by replacing 4 ?g of pcDNA3 with pcDNA3/RTA (a
pcDNA3-based vector expressing genomic KSHV RTA under control of the
CMV promoter) (16) or pFLAG/RTA (C-terminally FLAG-tagged KSHV RTA)
(3, 65). To control for any effect of PKA on the CMV promoter (56), data were
normalized to Renilla luciferase levels driven by the CMV promoter (pRLCMV).
To assess RTA-mediated trans-activation of a heterologous KSHV promoter,
luciferase reporter assays were conducted as above after replacing pRpluc with
pLUC/-69, a pGL3-Basic vector expressing firefly luciferase from a 69-nucleotide
fragment of the KSHV PAN RNA promoter (64).
Expressed RTA protein levels were assessed in parallel using flow cytometry to
detect C-terminal EGFP-tagged ORF50 expressed under control of the same
promoter (a kind gift from Joonho Choe, Department of Biological Sciences,
Korean Advanced Institute of Science and Technology) (26). Fluorescence in-
tensity data were acquired on a FACScan flow cytometer (BD Immunocytom-
etry) with dead cells and debris excluded on the basis of forward versus side
scatter gating using CellQuest software. In each of three replicate experiments,
GFP expression was quantified as a percent change above background fluores-
cence levels, with statistical significance of differences assessed by paired t test.
RTA phosphorylation targets. The predicted amino acid sequence of RTA
(AF091348) was scanned by Phosphobase 2.0 (32) to identify consensus PKA
VOL. 79, 2005
?-ADRENERGIC REACTIVATION OF KSHV13539
phosphorylation sites (55). Functional significance of predicted PKA and PKC
phosphorylation candidates at serines 526 (S525) and 526 (S526) was assessed by
converting both codons to alanines through site-directed mutagenesis of
pcDNA3/RTA-FLAG, with PCR primers changing RTA nucleotides 1573 to
1574 from GC to AG to produce S525A; and nucleotide 1576 from G to T to
produce S526A. Mutations were verified by sequencing and characterized in
PAN promoter luciferase assays as above.
Overexpression of ?-adrenoreceptors. To assess the effects of ?-adrenorecep-
tor overexpression in the absence of exogenous ligands, PEL cells were electro-
porated as described above with 4 ?g of mFLAG-?-pcDNA3 (a gift from B.
Kobilka, Stanford University) or pcDNA3 (empty vector). At 24 and 48 h later,
cells were assayed for lytic protein expression by ORF59 flow cytometry as
Effect of catecholamines on latent KSHV infection. To de-
termine whether autonomic nervous system activity might re-
activate lytic replication of KSHV, we treated three latently
infected B-cell lines with physiological concentrations of the
catecholamine neurotransmitters epinephrine and norepi-
nephrine (18, 39, 58, 61) and assayed lytic protein expression
48 h later. Catecholamine concentrations ranging from 1 nM to
100 ?M activated lytic gene expression in KS-1, BC3, and
BCBL-1 cells when assessed at the level of protein (Fig. 1A)
and mRNA (Fig. 1B). All lytic gene classes were induced,
ranging from the immediate-early gene RTA/ORF50 to the
late lytic gene ORF29 (Fig. 1C). Similar effects were observed
for representative early genes (e.g., ORF59; data not shown).
Norepinephrine induced KSHV DNA replication (Fig. 1D),
and PCR analysis of cell culture supernatants confirmed an
increased concentration of particle-associated KSHV DNA
(Fig. 1E). Supernatants from norepinephrine-treated PEL
cells contained sufficient concentrations of viral particles to
permit de novo infection of primary PBMC (Fig. 1F). Thus,
physiological concentrations of norepinephrine can induce the
entire lytic replication cycle in lymphoid cells latently infected
Mediating receptors. Most catecholamine effects on leuko-
cytes are mediated by ?-adrenergic receptors (63) (?AR), but
?-adrenoreceptors can also be expressed under certain condi-
tions (30). To identify the specific receptors mediating cate-
cholamine reactivation of KSHV, we pretreated PEL cells with
either the ? antagonist propranalol or the ? antagonist phen-
tolamine for 1 h prior to norepinephrine exposure. ?-Blockade
efficiently inhibited norepinephrine-mediated reactivation of
KSHV lytic protein (Fig. 2A) and mRNA (Fig. 2B). In con-
trast, ?-adrenergic blockade did not affect norepinephrine-
mediated reactivation of KSHV lytic gene expression (Fig. 2B).
Neither inhibitor had any effect on HHV-8 lytic replication in
the absence of exogenous norepinephrine (data not shown).
To define the subtype of ? receptor responsible for these
effects, we assayed mRNA for each of the three known ?AR
types by RT-PCR and found only ?1adrenoreceptor mRNA to
be detectable in appreciable quantities in PEL cells (Fig. 2C).
Catecholamine sensitivity profiles matched the known pharma-
cology of ?1ARs, with epinephrine triggering a low-range in-
crease in KSHV lytic gene expression (?1 nM range) in addi-
tion to paralleling norepinephrine-mediated induction over the
range of 100 nM to 10 ?M (Fig. 1B). ?-Adrenoreceptors re-
quired exogenous stimulation to activate KSHV lytic genes, as
shown by studies in which overexpression of ?-adrenoreceptors
in the absence of adrenergic ligands failed to induce ORF59
protein expression (data not shown; average 0.4% cells positive
by flow cytometry at 24 h versus 0.2% positive for the empty
vector control, difference P ? 0.625).
Role of the cAMP/PKA signaling pathway. In lymphoid cells,
?ARs signal primarily through G?s-mediated activation of the
adenylyl cyclase/cAMP/PKA signaling pathway (29, 63). Other
signaling pathways can potentially be activated (15, 38, 79), so
we sought to verify PKA’s role by treating PEL cells with the
PKA antagonist KT5720 prior to catecholamine exposure.
KT5720 inhibited norepinephrine induction of ORF29 mRNA
by more than 90% (Fig. 2D), and had similar effects on RTA/
ORF50 (data not shown). In contrast, the protein kinase C
(PKC) antagonists chelerythrine chloride and bisindolylmale-
imide hydrochloride failed to block norepinephrine-mediated
induction of KSHV lytic genes (results for chelerythrine chlo-
ride shown in Fig. 2D, and similar data for bisindolylmaleimide
hydrochloride not shown). Both PKC antagonists blocked
KSHV reactivation by PMA, but KT5720 had no effect on
PMA-mediated lytic gene expression. Thus, PKA and PKC
represent functionally independent signaling pathways for
KSHV reactivation in PEL cells.
Direct activation of cAMP signaling with the cell-permeable
analogue db-cAMP also activated KSHV lytic gene expression,
and other physiological cAMP inducers such as prostaglandin
E2, and histamine (H2) had similar effects (Fig. 2E). Hormone-
induced reactivation was specific to ligands that stimulate G?s/
adenylyl cyclase/cAMP/PKA signaling. Hormones that signal
through other pathways failed to induce KSHV lytic gene ex-
pression (e.g., acetylcholine or the glucocorticoid receptor ag-
onist dexamethasone) (Fig. 2E).
Activation of PKA is the most common mechanism by which
cAMP regulates cell function, but other cAMP targets have
been identified (17). To determine whether PKA activity can
account for cAMP-induced KSHV reactivation, we expressed a
constitutively active form of PKA in KS-1 and BCBL-1 cells
using a self-inactivating lentiviral vector. The ? catalytic sub-
unit of PKA (PKAc) was expressed from a bicistronic mRNA
that included a C-terminal EGFP reporter sequence indepen-
dently translated via a synthetic internal ribosome entry site.
This vector transduced more then 50% of KS-1 and BCBL-1
cells (Fig. 2F). Expression of the reporter gene alone had
minimal impact on KSHV lytic gene mRNA levels, but expres-
sion of the reporter gene in conjunction with PKAcup-regu-
lated expression of both immediate-early (RTA) and late lytic
transcripts (ORF29) (Fig. 2G). Thus, PKA activity alone is
sufficient to induce lytic gene expression in lymphoid cells
latently infected with KSHV.
Effect of PKA signaling on RTA expression and activity.
KSHV reactivation from latency is controlled by RTA-medi-
ated trans-activation, and we sought to determine how ?AR/
PKA signaling might modulate that process. To evaluate direct
effects on the RTA promoter in the absence of any contribution
from viral gene products, uninfected DG75 or Ramos cells
were electroporated with a reporter construct expressing firefly
luciferase under the control of ?3 kb of KSHV genomic DNA
upstream of ORF50 (previously described in reference 16).
Cells were then stimulated with the PKA activator db-cAMP
or the PKC activator PMA, and luciferase reporter activity was
assessed 18 h later. Reporter gene activity increased by fivefold
13540CHANG ET AL.J. VIROL.
following PKA activation in DG75 cells (Fig. 3A) and 14-fold
in Ramos cells (not shown), indicating that cellular transcrip-
tion factors alone are sufficient to mediate PKA induction of
the RTA promoter. Similar effects emerged in the context of
BC3 and KS-1 PEL cells (Fig. 3A and data not shown, respec-
Several cellular transcription factors respond to PKA, in-
cluding the CREB/ATF family of basic leucine zipper proteins
(5, 40, 47, 53). Bioinformatic analysis of the 3-kb sequence
upstream of ORF50 revealed six putative cAMP response el-
ements (CREs) that could potentially support CREB/ATF-
mediated activation of the RTA promoter (Fig. 3B). To define
the functional significance of these sites, we compared PKA
responsiveness of the full-length promoter (?3.09 kb) with
FIG. 1. Effect of norepinephrine (NE) on KSHV reactivation from latency. (A) Expression of the KSHV lytic cycle protein ORF59 was assayed
by flow cytometry using indirect immunofluorescence in BC3 cells fixed and permeabilized 24 h after exposure to 20 ng/ml PMA, 10 ?M
norepinephrine, or an equivalent volume of vehicle. (B) Dose dependence of catecholamine effects on KSHV lytic gene expression were also
assessed by quantitative real-time RT-PCR detection of KSHV ORF29 mRNA expression (normalized to GAPDH mRNA) 24 h following
exposure of BCBL-1 cells to indicated concentrations of epinephrine or norepinephrine. Similar results were observed for ORF50/RTA mRNA
(data not shown). (C) Kinetics of norepinephrine-induced lytic gene expression were assessed for immediate-early ORF50/RTA and late lytic
ORF29 at the indicated time points by real-time RT-PCR (data represent mean ? standard error fold change in target mRNA concentration in
three replicates after normalization to cellular GAPDH). Norepinephrine induction of ORF50/RTA mRNA was statistically significant at all time
points (12 h, P ? 0.0001; 24 h, P ? 0.0011; 36 h, P ? 0.0004; 48 h, P ? 0.0027; all by t test on log-transformed induction values). Norepinephrine
induction of ORF29 was not significant by 12 h (P ? 0.2086), but was highly significant at all subsequent time points (24 h, P ? 0.0003; 36 h, P
? 0.0006; 48 h, P ? 0.0003). Similar kinetics were observed in BCBL-1 cells (data not shown). (D) Gardella gel analysis of replicating KSHV DNA
was carried out by electrophoresis of KS-1 cell-associated DNA and hybridization with a32P-labeled PCR amplicon from the KSHV ORF50 locus
to distinguish episomal genomic DNA from linear replicating DNA. Data show cells treated for 48 h with vehicle (Control), PMA, norepinephrine,
or the PKA activator db-cAMP. (E) KSHV particle-associated DNA was assayed by PCR amplification of KSHV K8.1 sequences in 20 ?l of
DNase-treated 0.45-?m-filtered supernatant from KS-1 cell cultures treated with vehicle, PMA, or norepinephrine for 48 h (positive control is KS-1
cell-associated DNA). (F) Presence of infectious KSHV in cell culture supernatants from E was assessed by incubating PHA-stimulated PBMC
in filtered/DNased supernatants for 1 h followed by washing. 24 h later, PBMC were assayed for KSHV K8.1 DNA and cellular GAPDH by
real-time PCR. PBMC not exposed to PEL cell supernatants served as negative controls, and KS-1 cells served as positive controls.
VOL. 79, 2005
?-ADRENERGIC REACTIVATION OF KSHV13541
FIG. 2. Role of the ?-adrenergic receptor and cAMP/PKA signaling pathway in norepinephrine-induced reactivation of KSHV. (A) To define
the role of ?-adrenergic receptors in norepinephrine-mediated reactivation of lytic gene expression, BCBL-1 cells were treated with indicated
concentrations of the ?-antagonist propranalol for 1 h before exposure to 10 ?M norepinephrine, and then assayed for KSHV lytic protein
expression by Western blot 48 h later (primary antibody: Kaposi’s sarcoma patient serum). Results show induction of the major KSHV lytic protein
species at MW 49,800 which was the only band diagnostic of reactivation by the PMA positive control. (B) Specificity of adrenergic receptor
involvement was tested by treating KS-1 cells with the ?-antagonist phentolamine or the ?-antagonist propranalol for 1 h prior to norepinephrine
(NE) exposure; 24 h later, concentrations of mRNA for the immediate-early ORF50/RTA and late ORF29 were assessed by real-time RT-PCR
(values normalized to GAPDH and expressed as a ratio relative to vehicle-treated controls). (C) Expression of mRNA for ?1, ?2, and ?3adrenergic
receptors was assessed in untreated KS-1 and BCBL-1 cells by real-time RT-PCR. Products were resolved on a 3.5% agarose gel and compared
to positive control PCR products (parallel amplification of genomic DNA for ?1and ?3adrenergic receptors or a PBMC cDNA library for ?2
adrenergic receptors and GAPDH). To verify that RT-PCR results were free of contaminating DNA, BCBL-1 and KS-1 RNA samples were
amplified in parallel in the absence of reverse transcriptase (No RT). Data are representative of four independent experiments in which ?1
adrenergic receptors were consistently detected at high levels, and ?2and ?3receptors were not significantly expressed. (D) To evaluate the role
of PKA and PKC in norepinephrine activation of KSHV lytic gene expression, KS-1 cells were pretreated with the PKA antagonist KT5720 (1 ?M)
or the PKC antagonists chelerythrine chloride (Chel, 1 ?M) or bisindolylmaleimide HCl (300 nM) for 1 h prior to norepinephrine exposure.
Expression of mRNA for the late lytic gene ORF29 was assayed by real-time RT-PCR 24 h later. PKA blockade inhibited norepinephrine-induced
lytic gene expression by ?90%, but PKC inhibitors failed to block norepinephrine effects. Chelerythrine and bisindolylmaleimide (data not shown)
efficiently blocked PMA induction of ORF29, verifying that inhibitors were capable of blocking known PKC activators. KT5720 failed to block
PMA-mediated ORF29 expression, indicating that norepinephrine/PKA and PMA/PKC signaling pathways are functionally independent in PEL
cells. (E) To determine whether cAMP activity was sufficient to induce lytic gene expression, KS-1 cells were treated with graded doses of
pharmacological db-cAMP or with indicated concentrations of physiological cAMP inducers prostaglandin E2(PGE2) and histamine (H2).
cAMP/PKA specificity was evaluated using hormones that signal through alternative pathways, such as acetylcholine (ACh) or the glucocorticoid
dexamethasone (Dex). (F) To determine if PKA activity alone is sufficient to induce lytic gene expression, KS-1 cells were transduced with a
lentiviral vector (34) expressing the catalytic subunit of PKA (PKAc) and a downstream EGFP reporter sequence translated from a single mRNA
bearing the PS3 synthetic internal ribosome entry site (73). At 48 h following transduction with an empty vector, or vector bearing bicistronic EGFP
and PKAc, EGFP-positive cells were quantified by flow cytometry to assess transduction efficiency. Cells transduced with EGFP alone showed
transduction efficiencies comparable to comparable to EGFP plus PKAc(data not shown). (G) Also at 48 h posttransduction, concentrations of
mRNA for immediate-early ORF50/RTA and late lytic ORF29 were quantified by real-time RT-PCR (data normalized to GAPDH and expressed
as a ratio relative to empty vector controls). Data represent the mean (? standard error) of three independent experiments, with statistical
significance evaluated by paired t test. Similar effects were observed with BCBL-1 cells (data not shown).
13542 CHANG ET AL.J. VIROL.
that of truncated variants lacking the distal four CREs (be-
tween ?1.32 and ?2.04 kb) or all six CREs (?0.29 kb) (Fig.
3B). Reporter constructs were electroporated into BC3 PEL
cells (to assess activity in a the context of KSHV latency) or
uninfected Ramos cells (to assess activity in the absence of
KSHV gene products) and cells were subsequently treated for
18 h with db-cAMP or PMA. Deletion of the distal four CREs
reduced PKA inducibility by approximately 50% in BC3 PEL
cells (Fig. 3B), and by a similar amount in uninfected Ramos
cells (data not shown). Despite this quantitative reduction in
PKA responsiveness, the RTA promoter continued to show
significant induction by db-cAMP even when all six putative
CRE sites were eliminated (an average 17-fold enhancement
over basal activity in the ?0.29-kb construct, P ? 0.001; Fig.
3B). Thus, CREB/ATF factors may quantitatively enhance
PKA effects on the RTA promoter, but other transcription
factors also play a significant role.
Reporter studies conducted in KSHV-infected PEL cells
suggested that viral gene products might participate in PKA-
mediated activation of the RTA promoter. As shown in Fig.
3A, PKA activation up-regulated RTA promoter activity by
?16-fold in KSHV-infected BC3 cells, compared to ?5-fold in
uninfected DG75 tested in parallel (difference, P ? 0.0063 as
assessed by the cell type ? db-cAMP interaction term from a
factorial analysis of variance, F[1, 8] ? 13.49). Norepinephrine
had similar effects (data not shown), with RTA promoter ac-
tivity increasing by an average 9.37-fold in BCBL-1 PEL cells
(standard error ? 1.6-fold, P ? 0.0092 by t test), and signifi-
cantly less in DG75 cells (difference from BCBL-1, P ? 0.0012,
F[1, 4] ? 56.47).
One of the most powerful inducers of the RTA promoter is
the RTA protein itself (16, 25, 62). To determine whether PKA
might enhance RTA’s trans-activating capacity independently
of its effects on RTA protein levels, we expressed RTA in trans
from a heterologous promoter and assessed db-cAMP effects
on RTA promoter activity. As in previous studies (16, 25, 62),
RTA protein significantly enhanced activity of the RTA pro-
moter (Fig. 4A). Addition of db-cAMP increased that effect by
fourfold, from an average of sevenfold above basal activity to
more than 30-fold (Fig. 4A). This increase in trans-activating
capacity did not stem from changes in RTA protein levels,
which were measured in parallel using flow cytometry to quan-
tify expression of GFP-tagged ORF50 (right panel Fig. 4A).
Similar effects were observed in latently infected KS-1 and
BCBL-1 cells, with promoter activity increasing from ?16-fold
above basal activity levels in the presence of RTA alone to
?125-fold above basal activity following the addition of db-
cAMP (difference P ? 0.007 by t test; data not shown). Nor-
FIG. 3. Regulation of RTA promoter activity by PKA. (A) Activity of the RTA promoter was assessed by luciferase reporter assays in which
pRpluc (firefly luciferase coding sequence controlled by ?3 kb of KSHV genomic DNA upstream of ORF50) was electroporated into the Ramos
or DG75 B cell lines, which are known to be free of KSHV and Epstein-Barr virus, or into BC3 PEL cells containing latent KSHV. Following
electroporation, cells were incubated for 18 h in medium supplemented as indicated with the PKA activator db-cAMP (300 ?M) or the PKC
activator PMA (20 ng/ml). Firefly luciferase activity was normalized to Renilla luciferase activity generated by pRLCMV (Renilla luciferase under
control of the CMV promoter), and the statistical significance of triplicate determinations was evaluated by t test. Results showed significant
PKA-mediated induction of RTA promoter activity in both cell types, but db-cAMP up-regulated RTA promoter activity more strongly
KSHV-containing BC3 cells (as indicated by a cell type ? db-cAMP interaction term from a factorial analysis of variance, P ? 0.0001). Similar
effects were observed when PEL cell lines were treated with 10 ?M norepinephrine (data not shown). (B) Bioinformatic analysis of the RTA
promoter revealed six potential cAMP response elements (CREs) within 2 kb of the ORF50 transcription start site. To assess their role, we
compared db-cAMP effects on the full-length promoter (pRpluc) with effects on a series of truncation mutants omitting the distal 1.05 kb but
retaining all six putative CREs (pRp1), omitting the distal four CREs (pRp2), and omitting all predicted CREs (pRp8). Experiments were carried
out KS-1 cells as described above (A), with data represented as the mean (? standard error) fold enhancement in normalized luciferase activity
for db-cAMP-treated cells relative to controls. Deletion of the distal four CREs led to a quantitative reduction in PKA-inducibility, but db-cAMP
continued to activate the RTA promoter even in constructs lacking all predicted CRE sites (e.g., pRp8). Similar results emerged in BC3 cells (data
VOL. 79, 2005
?-ADRENERGIC REACTIVATION OF KSHV 13543
epinephrine also exerted similar effects, with RTA promoter
activity increasing from 15.5- ? 2.9-fold above basal levels with
RTA protein alone to 40.8- ? 1.4-fold above basal levels when
RTA was supplemented by 10 ?M norepinephrine (data not
shown). Thus, ?AR/PKA signaling appears to posttranslation-
ally enhance the ability of RTA protein to trans-activate the
To determine whether PKA enhancement of RTA trans-
activating capacity might impact other KSHV lytic genes, we
also analyzed activity of the promoter driving expression of the
KSHV polyadenylated nuclear (PAN) RNA gene (3, 64). In
uninfected DG75 or Ramos cells, db-cAMP had a negligible
effect on the PAN promoter (Fig. 4B and Fig. 5). Expression of
RTA alone significantly increased PAN promoter activity, as
previously reported (36, 64, 65, 70). Activation of PKA in
conjunction with RTA expression led to a further enhancement
of reporter gene activity, rising from ?60-fold above basal
activity to ?200-fold (Fig. 4B). Thus, PKA signaling can en-
hance RTA-mediated induction of KSHV lytic promoters even
when the PKA pathway has no direct effect on the promoter
via cellular transcription factors. Norepinephrine induced sim-
ilar dynamics, with RTA-mediated activation of the PAN pro-
moter rising from 150.4- ? 6.0-fold above basal activity to
1,518.0- ? 333.0-fold in the presence of norepinephrine (dif-
ference P ? 0.0097 by t test; data not shown).
Posttranslational effects on RTA. RTA protein is extensively
phosphorylated (36), but it is unclear what role this plays in
regulating lytic gene expression. Bioinformatic analysis of the
RTA coding sequence identified 10 potential PKA phosphor-
ylation sites (32, 55). One site at serine 526 (S526) fell adjacent
to a predicted PKC phosphorylation target at serine 525
(S525), suggesting that this region might represent a general-
ized target for serine/threonine kinases that modulate KSHV
activity. To evaluate that hypothesis, we converted both serine
codons to alanines via site-directed mutagenesis and tested the
resulting mutant RTA (S525A-S526A) for PKA-mediated en-
hancement of PAN promoter activity.
The S525A-S526A mutations substantially impaired PKA’s
ability to enhance the basal trans-activating capacity of RTA
(Fig. 5). PAN promoter activity increased ?50-fold above basal
levels when wild-type RTA was supplemented with PKA sig-
naling, but less than 2-fold when the S525A-S526A mutant was
accompanied by PKA activation (difference P ? 0.0001, as-
sessed by the interaction term from a factorial analysis of
variance, F[1, 8] ? 55.77). Thus, PKA-induced up-regulation
of RTA activity requires the S525/S526 tandem serines. PKC
also enhanced the trans-activating capacity of wild-type RTA
(PMA effects in Fig. 5), although this effect was less pro-
nounced than that of PKA. PKC’s effects were also abrogated
by the S525A-S526A mutations, suggesting that this region of
RTA might represent a generalized target for modulation by
serine/threonine kinases. Although the S525A-S526A mutant
was resistant to trans-activational enhancement by both PKA
and PKC, it remained a functional transcription factor as
shown by its capacity to induce the PAN promoter by an av-
FIG. 4. Posttranslational enhancement of RTA trans-activating capacity by PKA. (A) To determine whether RTA protein might contribute to
PKA-mediated activation of RTA promoter activity, FLAG-tagged RTA was expressed in trans (vector: pFLAG/RTA) in uninfected DG-75 cells
and supplemented by 1 mM db-cAMP or 20 ng/ml PMA as indicated. Luciferase activity was expressed as the mean fold change above basal
promoter activity for triplicate determinations, with statistical significance assessed by t test. Firefly luciferase activity was normalized to Renilla
luciferase from pRLCMV, driven by the same CMV immediate-early enhancer-promoter used to express RTA. To ensure that PKA effects were
not mediated by altered RTA protein levels, parallel electroporation studies quantified the density of GFP-tagged RTA by flow cytometry. Results
of three independent studies are reported as the mean ? standard error of green fluorescence intensity after subtraction of background
fluorescence intensity in cells electroporated with vector DNA alone. Neither db-cAMP nor PMA significantly enhanced RTA protein levels.
(B) To determine whether PKA-induced enhancement of RTA trans-activating capacity affects heterologous viral promoters, reporter assays were
carried out as in A substituting pLUC/-69 (firefly luciferase driven by the KSHV PAN promoter) (64) for the RTA reporter construct. Effects of
PKA signaling were tested in uninfected Ramos cells with no ectopic RTA (equivalent quantity of vector DNA) or ectopic RTA as in A. Parallel
flow cytometry analyses of cells electroporated with an ORF50-GFP expression vector verified substantial expression of ORF50 protein (P ? 0.001
by t test), but db-cAMP and PMA did not significantly alter protein quantity.
13544 CHANG ET AL.J. VIROL.
erage of 20-fold above its basal activity level (P ? 0.0001 by t
The present data show that physiological concentrations of
catecholamine neurotransmitters can efficiently reactivate
KSHV lytic replication in latently infected lymphoid cells.
These effects are mediated by ?1adrenoreceptors and subse-
quent activation of the cAMP/PKA signaling pathway. Other
activators of the PKA signaling pathway have similar effects
(e.g., db-cAMP, prostaglandin E2, histamine), and overexpres-
sion of the PKA catalytic subunit alone is sufficient to induce
KSHV lytic gene expression. ?-Adrenergic signaling induces
expression of KSHV genes from all stages of the lytic replica-
tion cycle and is accompanied by genome replication and pro-
duction of infectious viral particles. These effects appear to be
mediated by PKA regulation of the viral RTA gene–the pri-
mary “molecular switch” controlling KSHV lytic replication
(36, 70). Both norepinephrine and PKA induce RTA gene
expression via cellular transcription control pathways, and both
signals can also enhance the capacity of existing RTA protein
to trans-activate viral lytic cycle promoters (including the RTA
promoter). These results suggest that autonomic nervous sys-
tem activity could constitute an important determinant of
KSHV reactivation dynamics in vivo. ?drenergic reactivity also
suggests several potential therapeutic strategies, including
?-blockade to limit reactivation or use of ?-agonists to flush
latent virus into lytic replication for eradication by nucleoside
PKA signaling alone appears to be sufficient to reactivate
KSHV, as shown by the parallel effects of physiological PKA
inducers (e.g., catecholamines, prostaglandin E2, and hista-
mine), pharmacological PKA activators (e.g., db-cAMP), and
genetic overexpression of the PKA catalytic subunit. Comple-
mentary findings have recently emerged from a genome-wide
overexpression scan that identified the PKA catalytic subunit
as one of the most powerful inducers of RTA-mediated trans-
activation (J. Harada, Genomics Institute of the Novartis
Foundation). These data suggest that KSHV might reactivate
in response to a broad spectrum of extracellular signals that
converge on the cAMP/PKA second messenger system. Most
previous studies of KSHV have relied on PKC activators to
induce lytic replication (e.g., PMA/tetradecanoyl phorbol ace-
tate) (36). Like PKA, PKC is a ubiquitous serine/threonine
kinase mediating transcriptional response to a diverse array of
extracellular stimuli (51, 52). Although the biological effects of
these two kinases are often antagonistic, PKA and PKC target
similar amino acid motifs (52, 55). The PKA-mediated path-
way for KSHV reactivation is independent of the PKC-medi-
ated pathway, as shown by the fact that PKC inhibitors failed
to block norepinephrine induction of lytic gene expression.
However, the parallel effects of PKA and PKC on KSHV lytic
gene expression suggest a generalized role for cellular serine/
threonine kinases in regulating the balance between viral la-
tency and lytic replication.
The present studies identify two distinct mechanisms by
which the ?AR/cAMP/PKA signaling pathway can modulate
the activity of RTA. Cellular transcription control pathways
can directly activate the RTA promoter in the absence of
KSHV gene products (e.g., in uninfected B lymphoid cells),
and PKA can posttranslationally enhance the trans-activating
capacity of RTA protein. PKA control of RTA gene expression
appears to be mediated in part by CREB/ATF transcription
factors, as shown by a 50% decline in PKA-mediated induction
of RTA promoter constructs that lack any identifiable CRE.
However, PKA can still enhance RTA promoter activity by
?15-fold in the absence of CRE sites, suggesting that other
transcription factors also play a significant role. Identifying
those factors and mapping their mechanisms of action will
provide further information about the host cell processes reg-
ulating KSHV replication. As in the present analysis of the
?AR/cAMP/PKA pathway, such studies could also identify
new systemic regulators of KSHV activity and suggest further
targets for therapeutic intervention.
Alternative strategies for controlling KSHV reactivation
may also come from the discovery that PKA and PKC share a
common capacity to regulate the RTA protein’s activity as a
FIG. 5. Role of RTA serines 525 and 526 in PKA-mediated en-
hancement of trans-activation. To evaluate the functional role of pre-
dicted phosphorylation targets at S525 and S526 of RTA, both residues
were converted to alanines through PCR-based site-directed mutagen-
esis. RTA’s trans-activating capacity was assayed using a luciferase
reporter driven by the KSHV PAN promoter as in Fig. 4B. Reporter
constructs were electroporated into uninfected Ramos cells accompa-
nied by either 1 ?g of pFLAG/RTA (WT RTA), 1 ?g of pFLAG/
RTA-S525A-S526A (mutant), or 1 ?g of empty pFLAG (Vector).
Cells were subsequently cultured in the presence of 300 ?M db-cAMP
or 20 ng/ml PMA as indicated, and Firefly luciferase activity was
measured 18 h later as a fold change above basal activity after nor-
malization to control Renilla luciferase levels. Values represent the
mean (? standard error) fold increase in PAN promoter activity rel-
ative to the vehicle-treated control for each construct (Vector, WT
RTA, S525A-S526A mutant RTA). Similar results emerged when data
were normalized to a common basal condition (e.g., vehicle-treated
cells electroporated with the empty vector). In similar studies using
KS-1 cells and norepinephrine, endogenous regulation of RTA expres-
sion complicated analyses but data continued to show greater PKA-
induced up-regulation of RTA activity on the PAN promoter for wild-
type RTA versus the S525A-S526A mutant (results reported in main
VOL. 79, 2005
?-ADRENERGIC REACTIVATION OF KSHV13545
transcription factor. Kinase-mediated posttranslational modi-
fication of RTA could explain why PKA activates KSHV lytic
promoters more strongly in PEL cells than in similar B lym-
phoid cells lacking endogenous viral gene products. Given the
parallel effects of PKA and PKC on RTA activity, and the
functional blockade of those effects by phoshpho-resistant mu-
tations at serines 525 and 526, it is tempting to speculate that
kinase-induced phosphorylation mediates this posttransla-
tional effect. The presence of these serines within a nuclear
localization signal (33) also suggests a potential mechanism for
functional modulation. However, we have not been able to
directly verify changes in the phosphorylation of either wild-
type serine due to the high background levels of phosphoryla-
tion previously noted (36). In addition, mutation of either
serine alone failed to block PKA’s effect. This could indicate a
requirement for coordinated activity of PKA and PKC, or the
need for more complex secondary structural alterations to
completely block the effects of either kinase alone.
Posttranslational enhancement of RTA trans-activating ca-
pacity requires more detailed analysis, but the present data
clearly show that cellular kinases can regulate the activity of
this key viral switch protein. Similar results have emerged in
analyses of PKC-mediated control of the Epstein-Barr virus
BZLF transcription factor (1, 20, 22), suggesting that gamma-
herpesvirus lytic regulators may have evolved a functional sen-
sitivity to cellular kinases as a general mechanism for synchro-
nizing reactivation with favorable cellular conditions. If so,
pharmacological manipulation of cellular kinases could pro-
vide potential strategies for controlling gammaherpesvirus rep-
?drenergic reactivity is a common characteristic of ?- and
?-herpesviruses (2, 56, 59, 74, 75) and may explain the pro-
pensity of those viruses to reactivate in response to environ-
mental stress (27, 42, 54, 59, 74, 75). The gammaherpesvirus
Epstein-Barr virus is sensitive to stress-induced glucocorticoids
(4, 24, 35, 41, 67, 68), and the present studies suggest that
KSHV might also reactivate during stress-induced autonomic
nervous system activity. The emergence of stress-reactive biol-
ogy in all major classes of Herpesviridae suggests a potential
evolutionary advantage for latent viruses to monitor organis-
mal stress levels, which could be diagnostic of reduced cellular
immune threat (28, 54, 72) or lowered host-transmission hur-
dles (9, 69). Reactivation of KSHV in response to cat-
echolamines may thus represent one example in which herpes-
viruses monitor their host to detect ecological conditions
favoring their survival in much the same way they seek to
create those conditions by manipulating the cellular microen-
This work was supported by the National Institutes of Health (S.C.:
AI49135, AI52737; R.S.: CA91791 and DE14153; J.Z.: AI36059 and
AI 36554), the Norman Cousins Center at UCLA, and the James L.
Pendelton Charitable Trust.
1. Baumann, M., H. Mischak, S. Dammeier, W. Kolch, O. Gires, D. Pich, R.
Zeidler, H. J. Delecluse, and W. Hammerschmidt. 1998. Activation of the
Epstein-Barr virus transcription factor BZLF1 by 12-O-tetradecanoylphor-
bol-13-acetate-induced phosphorylation. J. Virol. 72:8105–8114.
2. Bloom, D. C., J. G. Stevens, J. M. Hill, and R. K. Tran. 1997. Mutagenesis
of a cAMP response element within the latency-associated transcript pro-
moter of HSV-1 reduces adrenergic reactivation. Virology 236:202–207.
3. Brown, H. J., M. J. Song, H. Deng, T. T. Wu, G. Cheng, and R. Sun. 2003.
NF-?B inhibits gammaherpesvirus lytic replication. J. Virol. 77:8532–8540.
4. Cacioppo, J. T., J. K. Kiecolt-Glaser, W. B. Malarkey, B. F. Laskowski, L. A.
Rozlog, K. M. Poehlmann, M. H. Burleson, and R. Glaser. 2002. Autonomic
and glucocorticoid associations with the steady-state expression of latent
Epstein-Barr virus. Horm. Behav. 42:32–41.
5. Carey, M., and S. T. Smale. 2000. Transcriptional regulation in eukaryotes:
concepts, strategies, and techniques. Cold Spring Harbor Laboratory Press,
Cold Spring Harbor, N.Y.
6. Cesarman, E., Y. Chang, P. S. Moore, J. W. Said, and D. M. Knowles. 1995.
Kaposi’s sarcoma-associated herpesvirus-like DNA sequences in AIDS-re-
lated body-cavity-based lymphomas. N. Engl. J. Med. 332:1186–1191.
7. Chang, J., R. Renne, D. Dittmer, and D. Ganem. 2000. Inflammatory cyto-
kines and the reactivation of Kaposi’s sarcoma-associated herpesvirus lytic
replication. Virology 266:17–25.
8. Chang, Y., E. Cesarman, M. S. Pessin, F. Lee, J. Culpepper, D. M. Knowles,
and P. S. Moore. 1994. Identification of herpesvirus-like DNA sequences in
AIDS-associated Kaposi’s sarcoma. Science 266:1865–1869.
9. Cole, S. W. 2005. The complexity of dynamic host networks, p. 605–628. In
T. S. Deisboeck and S. Kauffman (ed.), Complex systems science in biomed-
icine. Kluwer Academic-Plenum Publishers, New York, N.Y.
10. Cole, S. W., B. D. Jamieson, and J. A. Zack. 1999. cAMP externalizes
lymphocyte CXCR4: implications for chemotaxis and HIV infection. J. Im-
11. Cole, S. W., M. E. Kemeny, J. L. Fahey, J. A. Zack, and B. D. Naliboff. 2003.
Psychological risk factors for HIV pathogenesis: mediation by the autonomic
nervous system. Biol. Psychiatry 54:1444–1456.
12. Cole, S. W., M. E. Kemeny, S. E. Taylor, B. R. Visscher, and J. L. Fahey.
1996. Accelerated course of human immunodeficiency virus infection in gay
men who conceal their homosexual identity. Psychosomatic Med. 58:219–
13. Cole, S. W., Y. D. Korin, J. L. Fahey, and J. A. Zack. 1998. Norepinephrine
accelerates HIV replication via protein kinase A-dependent effects on cyto-
kine production. J. Immunol. 161:610–616.
14. Cole, S. W., B. D. Naliboff, M. E. Kemeny, M. P. Griswold, J. L. Fahey, and
J. A. Zack. 2001. Impaired response to HAART in HIV-infected individuals
with high autonomic nervous system activity. Proceedings of the National
Academy of Sciences of the USA. 98:12695–12700.
15. Davare, M. A., V. Avdonin, D. D. Hall, E. M. Peden, A. Burette, R. J.
Weinberg, M. C. Horne, T. Hoshi, and J. W. Hell. 2001. A beta2 adrenergic
receptor signaling complex assembled with the Ca2?channel Cav1.2. Science
16. Deng, H., A. Young, and R. Sun. 2000. Auto-activation of the rta gene of
human herpesvirus-8/Kaposi’s sarcoma-associated herpesvirus. J Gen. Virol.
17. de Rooij, J., F. J. Zwartkruis, M. H. Verheijen, R. H. Cool, S. M. Nijman, A.
Wittinghofer, and J. L. Bos. 1998. Epac is a Rap1 guanine-nucleotide-
exchange factor directly activated by cyclic AMP. Nature 396:474–477.
18. Dimsdale, J. E., and J. Moss. 1980. Plasma catecholamines in stress and
exercise. JAMA 243:340–342.
19. Dittmer, D., M. Lagunoff, R. Renne, K. Staskus, A. Haase, and D. Ganem.
1998. A cluster of latently expressed genes in Kaposi’s sarcoma-associated
herpesvirus. J. Virol. 72:8309–8315.
20. El-Guindy, A. S., L. Heston, Y. Endo, M. S. Cho, and G. Miller. 2002.
Disruption of Epstein-Barr virus latency in the absence of phosphorylation
of ZEBRA by protein kinase C. J. Virol. 76:11199–11208.
21. Fakhari, F. D., and D. P. Dittmer. 2002. Charting latency transcripts in
Kaposi’s sarcoma-associated herpesvirus by whole-genome real-time quan-
titative PCR. J. Virol. 76:6213–6223.
22. Francis, A., T. Ragoczy, L. Gradoville, L. Heston, A. El-Guindy, Y. Endo, and
G. Miller. 1999. Amino acid substitutions reveal distinct functions of serine
186 of the ZEBRA protein in activation of early lytic cycle genes and synergy
with the Epstein-Barr virus R transactivator. J. Virol. 73:4543–4551.
23. Gardella, T., P. Medveczky, T. Sairenji, and C. Mulder. 1984. Detection of
circular and linear herpesvirus DNA molecules in mammalian cells by gel
electrophoresis. J. Virol. 50:248–254.
24. Glaser, R., L. A. Kutz, R. C. MacCallum, and W. B. Malarkey. 1995. Hor-
monal modulation of Epstein-Barr virus replication. Neuroendocrinology
25. Gradoville, L., J. Gerlach, E. Grogan, D. Shedd, S. Nikiforow, C. Metroka,
and G. Miller. 2000. Kaposi’s sarcoma-associated herpesvirus open reading
frame 50/Rta protein activates the entire viral lytic cycle in the HH-B2
primary effusion lymphoma cell line. J. Virol. 74:6207–6212.
26. Gwack, Y., H. Byun, S. Hwang, C. Lim, and J. Choe. 2001. CREB-binding
protein and histone deacetylase regulate the transcriptional activity of Ka-
posi’s sarcoma-associated herpesvirus open reading frame 50. J. Virol. 75:
27. Jenkins, F. J., and A. Baum. 1995. Stress and reactivation of latent herpes
simplex virus: a fusion of behavioral medicine and molecular biology. Ann.
Behav. Med. 17:116–123.
28. Kalinichenko, V. V., M. B. Mokyr, L. H. Graf, Jr., R. L. Cohen, and D. A.
Chambers. 1999. Norepinephrine-mediated inhibition of antitumor cytotoxic
13546CHANG ET AL.J. VIROL.
T lymphocyte generation involves a beta-adrenergic receptor mechanism and
decreased TNF-alpha gene expression. J. Immunol. 163:2492–2499.
29. Kammer, G. M. 1988. The adenylate cyclase-cAMP-protein kinase A path-
way and the regulation of the immune response. Immunol. Today 9:222.
30. Kavelaars, A. 2002. Regulated expression of alpha-1 adrenergic receptors in
the immune system. Brain Behav. Immunol. 16:799–807.
31. Kedes, D. H., and D. Ganem. 1997. Sensitivity of Kaposi’s sarcoma-associ-
ated herpesvirus replication to antiviral drugs. Implications for potential
therapy. J. Clin. Investig. 99:2082–2086.
32. Kreegipuu, A., N. Blom, and S. Brunak. 1999. PhosphoBase, a database of
phosphorylation sites: release 2.0. Nucleic Acids Res. 27:237–239.
33. Krishnan, R., A. Kavirayani, K. Driscoll, W. Bu, D. Palmeri, and D. M.
Lukac. 2004. Nuclear/cytoplasmic localization of the KSHV ORF50/RTA
protein can be manipulated to regulate its activity. Abstract from the Seventh
international workshop on KSHV and related agents, University of Califor-
nia, Santa Cruz, 21 August 2004.
34. Kung, S. K., S. S. An, and I. S. Chen. 2000. A murine leukemia virus (MuLV)
long terminal repeat derived from rhesus macaques in the context of a
lentivirus vector and MuLV gag sequence results in high-level gene expres-
sion in human T lymphocytes. J. Virol. 74:3668–3681.
35. Kupfer, S. R., and W. C. Summers. 1990. Identification of a glucocorticoid-
responsive element in Epstein-Barr virus. J. Virol. 64:1984–1990.
36. Lukac, D. M., J. R. Kirshner, and D. Ganem. 1999. Transcriptional activa-
tion by the product of open reading frame 50 of Kaposi’s sarcoma-associated
herpesvirus is required for lytic viral reactivation in B cells. J. Virol. 73:9348–
37. Lukac, D. M., R. Renne, J. R. Kirshner, and D. Ganem. 1998. Reactivation
of Kaposi’s sarcoma-associated herpesvirus infection from latency by expres-
sion of the ORF 50 transactivator, a homolog of the EBV R protein. Virol-
38. Luttrell, L. M., S. S. Ferguson, Y. Daaka, W. E. Miller, S. Maudsley, G. J.
Della Rocca, F. Lin, H. Kawakatsu, K. Owada, D. K. Luttrell, M. G. Caron,
and R. J. Lefkowitz. 1999. Beta-arrestin-dependent formation of beta2 ad-
renergic receptor-Src protein kinase complexes. Science 283:655–661.
39. Madden, K. S., V. M. Sanders, and D. L. Felten. 1995. Catecholamine
influences and sympathetic neural modulation of immune responsiveness.
Annu. Rev. Pharmacol. Toxicol. 35:417–448.
40. Mayr, B., and M. Montminy. 2001. Transcriptional regulation by the phos-
phorylation-dependent factor CREB. Nat. Rev. Mol. Cell. Biol. 2:599–609.
41. Mehta, S. K., D. L. Pierson, H. Cooley, R. Dubow, and D. Lugg. 2000.
Epstein-Barr virus reactivation associated with diminished cell-mediated im-
munity in Antarctic expeditioners. J. Med. Virol. 61:235–240.
42. Mehta, S. K., R. P. Stowe, A. H. Feiveson, S. K. Tyring, and D. L. Pierson.
2000. Reactivation and shedding of cytomegalovirus in astronauts during
spaceflight. J. Infect. Dis. 182:1761–1764.
43. Mellon, P. L., C. H. Clegg, L. A. Correll, and G. S. McKnight. 1989. Regu-
lation of transcription by cyclic AMP-dependent protein kinase. Proc. Natl.
Acad. Sci. USA 86:4887–4891.
44. Mercader, M., B. Taddeo, J. R. Panella, B. Chandran, B. J. Nickoloff, and
K. E. Foreman. 2000. Induction of HHV-8 lytic cycle replication by inflam-
matory cytokines produced by HIV-1-infected T cells. Am. J. Pathol. 156:
45. Miller, G., M. O. Rigsby, L. Heston, E. Grogan, R. Sun, C. Metroka, J. A.
Levy, S. J. Gao, Y. Chang, and P. Moore. 1996. Antibodies to butyrate-
inducible antigens of Kaposi’s sarcoma-associated herpesvirus in patients
with HIV-1 infection. N. Engl. J. Med. 334:1292–1297.
46. Monini, P., S. Colombini, M. Sturzl, D. Goletti, A. Cafaro, C. Sgadari, S.
Butto, M. Franco, P. Leone, S. Fais, G. Melucci-Vigo, C. Chiozzini, F.
Carlini, G. Ascherl, E. Cornali, C. Zietz, E. Ramazzotti, F. Ensoli, M.
Andreoni, P. Pezzotti, G. Rezza, R. Yarchoan, R. C. Gallo, and B. Ensoli.
1999. Reactivation and persistence of human herpesvirus-8 infection in B
cells and monocytes by Th-1 cytokines increased in Kaposi’s sarcoma. Blood
47. Montminy, M. 1997. Transcriptional regulation by cyclic AMP. Annu. Rev.
48. Moore, P. S., and Y. Chang. 2001. Kaposi’s sarcoma-associated herpesvirus,
p. 2803–2833. In D. M. Knipe and P. M. Howley (ed.), Fields’ virology, vol.
2. Lippincott Williams & Wilkins, Philadelphia, Pa.
49. Moore, P. S., and Y. Chang. 2001. Molecular virology of Kaposi’s sarcoma-
associated herpesvirus. Phil. Trans. R. Soc. Lond. B Biol. Sci. 356:499–516.
50. Moore, P. S., S. J. Gao, G. Dominguez, E. Cesarman, O. Lungu, D. M.
Knowles, R. Garber, P. E. Pellett, D. J. McGeoch, and Y. Chang. 1996.
Primary characterization of a herpesvirus agent associated with Kaposi’s
sarcomae. J. Virol. 70:549–558.
51. Nishizuka, Y. 1992. Intracellular signaling by hydrolysis of phospholipids and
activation of protein kinase C. Science 258:607–614.
52. Nishizuka, Y. 1986. Studies and perspectives of protein kinase C. Science
53. Pabo, C. O., and R. T. Sauer. 1992. Transcription factors: structural families
and principles of DNA recognition. Annu. Rev. Biochem. 61:1053–1095.
54. Padgett, D. A., J. F. Sheridan, J. Dorne, G. G. Berntson, J. Candelora, and
R. Glaser. 1998. Social stress and the reactivation of latent herpes simplex
virus type 1. Proc. Natl. Acad. Sci. USA 95:7231–7235.
55. Pearson, R. B., and B. E. Kemp. 1991. Protein kinase phosphorylation site
sequences and consensus specificity motifs: tabulations. Methods Enzymol.
56. Prosch, S., C. E. C. Wendt, P. Reinke, C. Priemer, M. Pooert, D. H. Kruger,
H.-D. Volk, and W.-D. Docke. 2000. A novel link between stress and human
cytomegalovirus (HCMV) infection: sympathetic hyperactivity stimulates
HCMV activation. Virology 272:357–365.
57. Renne, R., W. Zhong, B. Herndier, M. McGrath, N. Abbey, D. Kedes, and D.
Ganem. 1996. Lytic growth of Kaposi’s sarcoma-associated herpesvirus (hu-
man herpesvirus 8) in culture. Nat. Med. 2:342–346.
58. Richter, S. D., T. H. Schurmeyer, M. Schedlowski, A. Hadicke, U. Tewes,
R. E. Schmidt, and T. O. Wagner. 1996. Time kinetics of the endocrine
response to acute psychological stress. J. Clin. Endocrinol. Metab. 81:1956–
59. Roizman, B., and D. M. Knipe. 2001. Herpes simplex viruses and their
replication, p. 2399–2459. In D. M. Knipe and P. M. Howley (ed.), Fields
virology, 4th ed., vol. 2. Lippincott Williams and Wilkins, Philadelphia, Pa.
60. Russo, J. J., R. A. Bohenzky, M. C. Chien, J. Chen, M. Yan, D. Maddalena,
J. P. Parry, D. Peruzzi, I. S. Edelman, Y. Chang, and P. S. Moore. 1996.
Nucleotide sequence of the Kaposi sarcoma-associated herpesvirus (HHV8).
Proc. Natl. Acad. Sci. USA 93:14862–14867.
61. Saitoh, M., T. Yanagawa, T. Kondoh, H. Miyakoda, H. Kotake, and H.
Mashiba. 1995. Neurohumoral factor responses to mental (arithmetic) stress
and dynamic exercise in normal subjects. Intern. Med. 34:618–622.
62. Sakakibara, S., K. Ueda, J. Chen, T. Okuno, and K. Yamanishi. 2001.
Octamer-binding sequence is a key element for the autoregulation of Kapo-
si’s sarcoma-associated herpesvirus ORF50/lytA gene expression. J. Virol.
63. Sanders, V. M., and R. H. Straub. 2002. Norepinephrine, the beta-adrenergic
receptor, and immunity. Brain Behav. Immun. 16:290–332.
64. Song, M. J., H. J. Brown, T. T. Wu, and R. Sun. 2001. Transcription activa-
tion of polyadenylated nuclear RNA by RTA in human herpesvirus 8/Ka-
posi’s sarcoma-associated herpesvirus. J. Virol. 75:3129–3140.
65. Song, M. J., X. Li, H. J. Brown, and R. Sun. 2002. Characterization of
interactions between RTA and the promoter of polyadenylated nuclear
RNA in Kaposi’s sarcoma-associated herpesvirus/human herpesvirus 8.
J. Virol. 76:5000–5013.
66. Soulier, J., L. Grollet, E. Oksenhendler, P. Cacoub, D. Cazals-Hatem, P.
Babinet, M. F. d’Agay, J. P. Clauvel, M. Raphael, L. Degos, et al. 1995.
Kaposi’s sarcoma-associated herpesvirus-like DNA sequences in multicen-
tric Castleman’s disease. Blood 86:1276–1280.
67. Stowe, R. P., S. K. Mehta, A. A. Ferrando, D. L. Feeback, and D. L. Pierson.
2001. Immune responses and latent herpesvirus reactivation in spaceflight.
Aviat. Space Environ. Med. 72:884–891.
68. Stowe, R. P., D. L. Pierson, D. L. Feeback, and A. D. Barrett. 2000. Stress-
induced reactivation of Epstein-Barr virus in astronauts. Neuroimmuno-
69. Stumpf, M. P. H., Z. Laidlaw, and V. A. A. Jansen. 2002. Herpesviruses
hedge their bets. Proc. Natl. Acad. Sci. USA 99:15234–15237.
70. Sun, R., S. F. Lin, L. Gradoville, Y. Yuan, F. Zhu, and G. Miller. 1998. A
viral gene that activates lytic cycle expression of Kaposi’s sarcoma-associated
herpesvirus. Proc. Natl. Acad. Sci. USA 95:10866–10871.
71. Sun, R., S. F. Lin, K. Staskus, L. Gradoville, E. Grogan, A. Haase, and G.
Miller. 1999. Kinetics of Kaposi’s sarcoma-associated herpesvirus gene ex-
pression. J. Virol. 73:2232–2242.
72. Valitutti, S., M. Dessing, and A. Lanzavecchia. 1993. Role of cAMP in
regulating cytotoxic T lymphocyte adhesion and motility. Eur. J. Immunol.
73. Venkatesan, A., and A. Dasgupta. 2001. Novel fluorescence-based screen to
identify small synthetic internal ribosome entry site elements. Mol. Cell.
74. Wagner, E. K., and D. C. Bloom. 1997. Experimental investigation of herpes
simplex virus latency. Clin. Microbiol. Rev. 10:419–443.
75. Whitley, R. J. 2001. Herpes simplex viruses, p. 2461–2509. In D. M. Knipe
and P. M. Howley (ed.), Fields virology, 4th ed., vol. 2. Lippincott Williams
and Wilkins, Philadelphia, Pa.
76. Wingender, E., P. Dietze, H. Karas, and R. Knuppel. 1996. TRANSFAC: a
database on transcription factors and their DNA binding sites. Nucleic Acids
77. Zennou, V., C. Petit, D. Guetard, U. Nerhbass, L. Montagnier, and P.
Charneau. 2000. HIV-1 genome nuclear import is mediated by a central
DNA flap. Cell 101:173–185.
78. Zhong, W., H. Wang, B. Herndier, and D. Ganem. 1996. Restricted expres-
sion of Kaposi sarcoma-associated herpesvirus (human herpesvirus 8) genes
in Kaposi sarcoma. Proc. Natl. Acad. Sci. USA 93:6641–6646.
79. Zou, Y., I. Komuro, T. Yamazaki, S. Kudoh, H. Uozumi, T. Kadowaki, and
Y. Yazaki. 1999. Both Gs and Gi proteins are critically involved in isopro-
terenol-induced cardiomyocyte hypertrophy. J. Biol. Chem. 274:9760–9770.
VOL. 79, 2005
?-ADRENERGIC REACTIVATION OF KSHV 13547