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NEOPLASIA
Hypoxia induces lytic replication of Kaposi sarcoma–associated herpesvirus
David A. Davis, Andrea S. Rinderknecht, J. Paul Zoeteweij, Yoshiyasu Aoki, Elizabeth L. Read-Connole, Giovanna Tosato,
Andrew Blauvelt, and Robert Yarchoan
There is substantial evidence that Kaposi
sarcoma–associated herpesvirus (KSHV)
plays an important role in the pathogenesis of all forms of Kaposi sarcoma (KS). It
has been noted that KS commonly occurs
in locations, such as the feet, where tissue may be poorly oxygenated. On the
basis of this observation, the potential
role of hypoxia in the reactivation of KSHV
replication was explored by studying 2
KSHV-infected primary effusion lymphoma B-cell lines (BC-3 and BCBL-1)
latently infected with KSHV. Acute and
chronic exposure of these cells to hypoxia (1% O2) induced KSHV lytic replication, as indicated by an increase in intracellular lytic protein expression and
detection of virus in cell supernatants by
Western immunoblotting. In addition, hypoxia increased the levels of secreted viral
interleukin-6. Moreover, hypoxia enhanced the lytic replication initiated by
the viral inducer 12-O-tetradecanoylphorbol-13-acetate. Desferoxamine and cobalt
chloride, 2 compounds that increase the
intracellular levels of hypoxia-inducible
factor 1, were also able to induce KSHV
lytic replication. These studies suggest
that hypoxia is an inducer of KSHV replication. This process may play an important role in the pathogenesis of
KS. (Blood. 2001;97:3244-3250)
© 2001 by The American Society of Hematology
Introduction
replication may contribute to the development of KS. Cells latently
infected with KSHV can be activated to undergo lytic replication
by phorbol esters. However, relatively little is presently known
about the physiologic signals that serve to activate KSHV in
infected patients.
In his original description of KS in 1872, Moritz Kaposi noted
that the painful edema and characteristic lesions of the condition
occurred primarily on the feet of his patients, and we have been
struck by a predilection of KS to involve the feet in both classic and
AIDS-associated forms.17 The lower extremities often have relatively low tissue oxygen (O2) concentrations,18 and we hypothesized that such hypoxic conditions might reactivate KSHV and
lead to the subsequent production of proangiogenic factors, thus
favoring the development of KS. In this report, we investigated a
possible role for hypoxia in the re-activation of KSHV in 2
KSHV-infected PEL cell lines.
Kaposi sarcoma (KS) is a multifocal proliferative disease of
vascular origin. KS lesions are characterized by proliferation and
growth of spindle-shaped neoplastic cells that appear to be of
lymphatic endothelial cell origin.1-3 KS is the most common
neoplasm in human immunodeficiency virus type 1–infected
individuals.1 Even before the advent of acquired immunodeficiency
syndrome (AIDS), KS was known to be more prevalent in certain
geographic areas and groups of patients (classic and endemic KS).
A classic form of KS is described as occurring in elderly men in
southern Italy and other Mediterranean countries,4 and an endemic
form of KS has been observed to occur in sub-Saharan Africa.5,6 In
addition, transplant recipients and other immunosuppressed patients are at risk for developing KS.4
Since the discovery of Kaposi sarcoma–associated herpesvirus
(KSHV), also called human herpesvirus-8 (HHV-8), in 1994,7
substantial evidence has been accumulated that implicates it as an
essential factor in the pathogenesis of all forms of KS.8,9 It is also
involved in the pathogenesis of primary effusion lymphoma (PEL)
and certain cases of multicentric Castleman disease.10 KSHV DNA
is detected in nearly all KS lesions.7,11,12 As with other human
herpesviruses, infection with KSHV can be latent or lytic. Activation of viral replication in latently infected B cells, endothelial
cells, or other target cells may be responsible for viral spread and
contribute to the development of KS.4 KSHV encodes for several
cellular gene mimics that are produced during lytic replication and
that have proangiogenic activity.13-16 The virus also encodes several
cellular protein mimics with oncogenic potential, such as v-cyc, a
D-type cyclin, and a member of the interferon regulatory factor
family.16 The production of these viral factors during KSHV lytic
The KSHV-positive and Epstein-Barr virus–negative PEL cell lines BCBL119 (National Institutes of Health AIDS Research and Reagent Program,
Rockville, MD) and BC-320 (American Type Culture Collection, Rockville
MD) were grown in complete medium (RPMI 1640 medium containing
15% fetal calf serum, 2 mm/L glutamine, 50 U/mL penicillin, and 50
␮g/mL streptomycin; all from Gibco BRL [Gaithersburg, MD]). Cells were
cultured in either a standard incubator in atmospheric O2 or an incubator at
1% O2 by means of the Proox model 110 O2 controller (which controls O2
From the HIV and AIDS Malignancy Branch, the Dermatology Branch, and the
Medicine Branch, National Cancer Institute, National Institutes of Health,
Bethesda, MD.
Reprints: David A. Davis, HIV and AIDS Malignancy Branch, National Cancer
Institute, National Institutes of Health, Bldg 10, Rm 10S255, MSC 1868, 9000
Rockville Pike, Bethesda, MD 20892; e-mail: [email protected].
Submitted October 6, 2000; accepted January 23, 2001.
D.A.D and A.S.R. contributed equally to this manuscript.
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.
This is a US goverment work. There are no restrictions on its use.
© 2001 by The American Society of Hematology
3244
Materials and methods
Cell culture at 21% and 1% O2
BLOOD, 15 MAY 2001 䡠 VOLUME 97, NUMBER 10
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BLOOD, 15 MAY 2001 䡠 VOLUME 97, NUMBER 10
levels from 1% to 99%) (Reming Bioinstruments, Redfield, NY). The CO2
was maintained at 5% in both chambers, and the air in the low-O2 incubator
was displaced with nitrogen and continually monitored to maintain 1% O2.
Cells were induced with 12-O-tetradecanoylphorbol-13-acetate (TPA)
(Sigma, St Louis, MO). Two other compounds tested as inducers of KSHV
lytic replication, desferoxamine mesylate (Dfx) (10 mM and 5 mM)
(Sigma) and cobalt chloride (CoCl2) (100 mM and 5 mM) (Sigma), were
dissolved in phosphate-buffered salined (PBS), filter-sterilized, and stored
as stock solutions at ⫺20°C until use.
Analysis of KSHV lytic protein expression by flow cytometry
Cells were seeded at 300 000 viable cells per milliliter and incubated for 2
to 3 days as indicated. When inducers were used, cells were incubated
overnight, followed by the addition of the inducer, and incubated for
another 2 to 3 days under the same conditions. The intracellular expression
of processivity factor 8 (PF-8, an early-lytic protein) and glycoprotein 8.1
(gpK8.1, a late-lytic protein) in the viable cells was then determined by flow
cytometric analysis as described.21
HYPOXIC INDUCTION OF KSHV
3245
antibody (0.5 ␮g/mL) was added to the wells in PBS-T containing 0.5%
bovine serum albumin (BSA) (PBS-T/BSA). Plates were incubated for 2
hours at room temperature and washed. Affinity-purified human serum
protein–absorbed goat antirabbit immunoglobulin G antibody conjugated to
alkaline phosphatase (Sigma) (1:400 dilution in PBS-T/BSA) was added.
Plates were incubated at room temperature for 1.5 hours and washed with
PBS-T. A p-nitrophenolphosphate substrate (Sigma) was added to the wells,
and the plates were read at 405 nm with ␭ correction at 595 nm. A purified
fusion protein of MBP (42.7 kd) and amino acids 22 through 204 of vIL-6
(21.6 kd) were used as standards. The concentration of vIL-6 was calculated
from absorbance values in relation to the standard curve, corrected for the
presence of the MBP fusion protein in the vIL-6 standard (vIL-6 corresponds to 33.6% of MBP vIL-6). The assay sensitivity is approximately 30
pg/mL for vIL-6. Vascular endothelial growth factor (VEGF) was determined in the supernatant of cells by means of the Human VEGF kit from
Accucyte (College Park, MD) with a dynamic range of 0.195 to 50
ng/mL VEGF.
Statistical analysis
Western blot analysis for the viral minor capsid protein
The level of virus in the cellular supernatants of KSHV-infected cells was
determined by means of a peptide affinity–purified rabbit polyclonal
antibody directed toward a 17–amino acid peptide of the open reading
frame 26 (ORF 26) minor capsid protein (mCP) of KSHV as described
previously.22 To determine if we could use this assay to quantitatively
measure the level of virus in the media, we measured the mCP band
intensity for increasing concentrations of sucrose-purified KSHV (ABI,
Columbia, MD) by Western blot. We found that the mCP band intensity, as
determined by means of Un-Scan-It (Silk Scientific, Orem, UT), was
directly proportional to the amount of virus loaded from 1 ␮g (equivalent to
3.3 ⫻ 107 viral particles) to 16 ␮g of viral lysate protein loaded. Cell
culture samples were first centrifuged (10 minutes at 1000g) to remove the
cells. The supernatant was ultracentrifuged in 1 to 1.5 mL aliquots at
100 000g for 35 minutes at 4°C. The pelleted material was washed with 1
mL cold PBS, pelleted as above, and then resuspended in sodium dodecyl
sulfate sample buffer (Novex, San Diego, CA) and boiled at 70°C for 10
minutes. Samples that represented equal volumes of pelleted supernatant
were electrophoresed on 10% or on 4% to 12% bis-Tris polyacrylamide gels
with 2-(N-morphilino) ethanesulfonic acid running buffer, by means of the
NuPage system (Novex). Proteins were electroblotted onto nitrocellulose,
and the mCP was detected with a peptide affinity–purified rabbit antibody
as previously described.22 Briefly, blots were incubated with anti-mCP
rabbit antibody for 2 hours at room temperature and then for 30 minutes
with an antirabbit secondary antibody conjugated to alkaline phosphatase.
Bands were detected by means of the Western Blue–stabilized substrate for
alkaline phosphatase (Promega, Madison, WI). The band was identified by
comparison with a viral standard (ABI) containing the mCP band that
migrated to an approximate molecular mass of 34 kd as determined by
comparison with the migration positions of SeeBlue-prestained markers
(Novex). Immunoblots were scanned, and densitometry analyses of the
scanned images were performed.
Enzyme-linked immunosorbent assays for viral interleukin 6
and vascular endothelial growth factor
Levels of viral interleukin 6 (vIL-6) in culture supernatants were determined by means of a specific enzyme-linked immunosorbent assay as
previously described.23 Briefly, polystyrene plates (Immunol 1B; Dynex
Technologies, Chantilly, VA) were coated with mouse monoclonal antivIL-6 antibody (v6m12.1.1, 4 ␮g/mL in carbonate buffer, pH 9) overnight
at 4°C. After washing the plates with PBS containing 0.05% Tween 20
(PBS-T) and blocking with SuperBlock (Pierce, Rockford, IL), test samples
were added in duplicate to the wells. These samples were diluted in advance
with the same volume of PBS-T containing 10% fetal bovine serum (FBS);
final concentration of FBS was 10% for both samples and standards. Plates
were incubated overnight at room temperature and washed with PBS-T.
Maltose-binding protein (MBP)–absorbed rabbit polyclonal anti–vIL-6
The exact Wilcoxon rank sum test was used to compare expression of
KSHV-encoded antigens in cells cultured under normoxia and cells cultured
under chronic hypoxia. The Wilcoxon signed rank test was used to compare
parallel cultures established under normoxia and acute hypoxia at the
same time.
Results
Hypoxia increases KSHV lytic protein expression
in KSHV-infected PEL B-cell lines
BC-3 and BCBL-1 PEL B-cell lines were used to study the role of
hypoxia in KSHV reactivation. Viral reactivation was determined,
in part, by studying the expression of 2 lytic proteins, PF-8 and
gpK8.1, by flow cytometry. In normoxia (21% O2), the percentage
of viable cells expressing the early-lytic protein PF-8 was very low:
0.12% and 0.23% for BC-3 and BCBL-1 cells, respectively. Also,
the median percentage of cells expressing the late-lytic protein
gpK8.1 was very low, 0.04% and 0.09% for BC-3 and BCBL-1
cells, respectively (Figure 1). However, when BC-3 cells were
maintained under chronic hypoxic conditions (1% O2 for more than
3 cell passages), the median percentage of viable BC-3 cells
expressing PF-8 increased to 2.7%, a 22-fold increase (P ⫽ .012)
(Figure 1A). A similar increase in PF-8 expression from 0.23% to
2.86% was seen in BCBL-1 cells, representing a 12-fold increase
(P ⫽ .004) (Figure 1B). Late-lytic gpK8.1 protein expression was
also greater under hypoxic conditions, with BC-3 cells having a
median increase from 0.04% to 0.55% (13-fold increase) (P ⫽ .016)
and BCBL-1 cells having a median increase from 0.09% to 0.94%
(10-fold increase) (P ⫽ .028) (Figure 1A,B). It was observed
throughout these experiments that the cells grown in chronic
hypoxia had a somewhat lower cell growth rate and viability. This
decrease in viability seen in chronic hypoxia ranged from 10% to
25% compared with cells grown in normoxia.
The effect of acute hypoxia on KSHV lytic replication was also
investigated. An increase in the expression of both lytic proteins,
similar to that observed in chronic hypoxia, occurred in the B-cell
lines when acutely exposed to hypoxia (Figure 1C,D). However,
the extent of the increase in PF-8 expression as a result of acute
hypoxic exposure varied somewhat from experiment to experiment, ranging from a 7.7- to a 34-fold induction in BC-3 (median
increase, 9.7-fold) (Figure 1C,D). Similarly, there was a 3- to
12-fold induction of gpK8.1 (median increase, 6-fold) in BC-3
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3246
DAVIS et al
BLOOD, 15 MAY 2001 䡠 VOLUME 97, NUMBER 10
We then assessed the effect of both acute and chronic
hypoxia on the lytic protein expression induced by various doses
of TPA. As expected, TPA increased PF-8 and gpK8.1 expression in a dose-dependent manner 2 days after treatment in
normoxic conditions (Figure 2). However, the level of lytic
protein expression in response to TPA was significantly increased in cells exposed to both acute and chronic hypoxia
(Figure 2). Acute hypoxia was more effective than chronic
hypoxia at enhancing the induction of PF-8 and gpK8.1
expression in the presence of TPA for both PEL cell lines.
Detection of KSHV mCP under normoxic and
hypoxic conditions
Figure 1. Effect of chronic hypoxia (1% O2), acute hypoxia (1% O2), and acute
hypoxia plus TPA on KSHV intracellular lytic protein expression in BC-3 and
BCBL-1 cells 48 to 72 hours after treatment. Cells were seeded at 300 000 viable
cells per milliliter. (A, B) Cells were maintained in either 21% O2 or 1% O2 for at least 3
passages before being used in experiments (chronic exposure). The plotted values
for PF-8 (u) and gpK8.1 (o) expression represent the median of at least 3
independent experiments ⫾ the 25% and 75% quartiles. Exact Wilcoxon rank sum
test (unpaired) for panel A: (BC-3, PF-8 21% O2 vs 1% O2) p2 ⫽0 .0012, and (BC-3,
gpK8.1 21% O2 vs 1% O2) p2 ⫽ 0.016. Exact Wilcoxon rank sum test (unpaired) for
panel B: (BCBl-1, PF-8 21% O2 vs 1% O2) p2 ⫽ 0.004, and (BCBL-1, gpK8.1 21% O2
vs 1% O2) p2 ⫽ 0.028. (C, D) Cells grown in normoxia were incubated in 21% O2 or
1% O2 overnight. They were then exposed to TPA (6.4 nM) or to media control and
returned to their respective conditions. The plotted values for PF-8 expression is the
median percentage of positive cells for 6 independent experiments ⫾ the 25% and
75% quartiles. Wilcoxon signed rank test (paired) for panel C: (BC-3, PF-8 21% O2 vs
1% O2) p2 ⫽ 0.031, and (BC-3/TPA, PF-8 21% O2 vs 1% O2) p2 ⫽ 0.031. Wilcoxon
signed rank test (paired) for panel D: (BCBL-1, PF-8 21% O2 vs 1% O2) p2 ⫽ 0.031,
and (BCBL-1/TPA, PF-8 21% O2 vs 1% O2) p2 ⫽ 0.031.
To determine whether exposure to 1% O2 (hypoxia) also affected virus
production from KSHV-infected cells, we assayed for mCP in culture
supernatants under these conditions. Under normoxic conditions, we
could not detect mCP in pellets from 1 to 1.5 mL supernatants from
BCBL-1 or BC-3 cells. However, mCP was detected in supernatants
from BCBL-1 and BC-3 cells exposed to chronic hypoxia (Figure 3A).
BCBL-1 cells cultured in 5% O2 also produced increased levels of mCP,
although to a lesser degree than that seen at 1% O2 (data not shown). In
addition, BCBL-1 and BC-3 cells exposed to a suboptimal concentration of TPA produced higher levels of mCP when exposed to acute
hypoxia (Figure 3B). A low level of the mCP was detected 2 days after
6.4-nM TPA treatment in the supernatant of cells grown in 21% O2, and
the level further increased over the next 2 days, reaching a maximum on
day 4 (Figure 3B). When cells were exposed to acute hypoxia and 6.4
nM TPA, there was a 5-fold increase in the level of mCP observed 2
days after transfer to hypoxic conditions, as compared with TPA-treated
cells in normoxia. In addition, when cells were exposed to hypoxia, the
level of mCP reached a maximum a day earlier than for cells in
normoxia (Figure 3B). Qualitatively similar results were obtained with
BC-3 cells (data not shown). We also performed a TPA dose-response
cells (not shown). Acute hypoxia also increased PF-8 expression in
BCBL-1 cells, but to a lesser degree than that seen for BC-3 cells
(Figure 1C,D).
Hypoxia enhances TPA induction of KSHV lytic replication
TPA is an inducer of KSHV lytic replication in PEL cell lines
infected with KSHV.19,24 It was previously shown that the maximum percentages of cells that could be induced to undergo lytic
replication (based on PF-8 expression quantitated by flow cytometry) by 30 nM TPA 3 days after treatment in BCBL-1 and BC-3
cells were approximately 12% and 20%, respectively.21 We were
interested in determining whether hypoxia would affect lytic
induction by TPA in the 2 cell lines used. When cells exposed to
chronic hypoxia were treated with 30 nM TPA, substantial toxicity
(less than 40% viability) occurred in both of the cell lines. A
viability of less than 40% was our cutoff for assaying the level of
lytic protein expression, and protein expression was therefore not
determined for this concentration of TPA. Therefore, the effect of
chronic hypoxia on viral reactivation by TPA was assessed at lower
TPA concentrations. When cells were grown in normoxia and
treated with a suboptimal concentration of TPA (6.4 nM), the
median percentage of cells expressing PF-8 was 8.6% for BC-3 and
3.9% for BCBL-1 cells as determined by flow cytometry (Figure
1C,D). However, the median level of PF-8 expression more than
doubled when these TPA-treated cells were exposed to acute
hypoxia, resulting in 10.3% expression for BCBL-1 cells and
20.0% expression for BC-3 cells (Figure 1C,D).
Figure 2. Dose-dependent expression of PF-8 and gpK8.1 in TPA-stimulated
BCBL-1 cells in normoxia and hypoxia (acute and chronic exposure). BCBL-1
cells that had been cultured in normoxia (21% O2) or hypoxia (1% O2) were seeded at
300 000 viable cells per milliliter and incubated overnight at the indicated conditions.
Following the overnight incubation, the cells were treated with increasing concentrations of TPA and maintained at the indicated conditions following TPA treatment for 48
hours. In each panel, the values represent 1 of 2 similar experiments. (A) PF-8
expression in BCBL-1 cells in normoxia (f) or acute hypoxia (F). (B) gpK8.1
expression in BCBL-1 cells in normoxia (f) or acute hypoxia (F). (C) PF-8 expression
in BCBL-1 cells in normoxia (f) or chronic hypoxia (F). (D) gpK8.1 expression in
BCBL-1 cells in normoxia (f) or chronic hypoxia (F).
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BLOOD, 15 MAY 2001 䡠 VOLUME 97, NUMBER 10
experiment to determine the effect of chronic hypoxia on TPA induction
of virus. Again, the cells grown in chronic hypoxia produced detectable
mCP even in the absence of TPA induction; those grown in normoxia
did not. TPA treatment at all concentrations further increased mCP
production in cells exposed to hypoxia; cells exposed to normoxia
produced measurable virus only at the 2 highest TPA concentrations (6.4
and 30 nM) (results not shown).
Hypoxia is known to induce the production of VEGF in normal
cells, and we considered whether VEGF production induced by
hypoxia in the PEL cells might have acted in an autocrine manner
to reactive KSHV replication. Consistent with a previous report,25
BC-3 cells produced substantial VEGF (1.43 ng/106 cells; 2.8
ng/mL media) after 3 days of culture under normoxic conditions.
However, although there was a small increase in the VEGF
produced per cell when these cells were exposed to 1% O2 (to 2.5
ng/106 cells), there was no net change in the VEGF released into the
media (2.75 ng/mL media). Also, addition of VEGF at 10 or 100
ng/mL for 3 days had no substantial effect on the level of
expression of PF-8 or gpK8.1 by BC-3 or BCBL-1 cells over that
seen in control cells. Taken together, these results suggest that
hypoxia-induced KSHV reactivation was not the result of an
up-regulation of VEGF in these cells by hypoxia.
Effect of hypoxia on the induction of KSHV vIL-6 production
Viral IL-6 is a cytokine thought to play a role in the pathogenesis of
KS by stimulating spindle cell growth and activating the produc-
HYPOXIC INDUCTION OF KSHV
3247
Figure 4. Effect of acute hypoxia and TPA on secreted vIL-6 production and
intracellular PF-8 expression from KSHV-infected BCBL-1 cells. BCBL-1 cells
were seeded at 300 000 cells per milliliter and then incubated in normoxia or hypoxia
(1% O2). Following incubation overnight, cells were either left untreated or treated
with TPA (6.4 nM) and returned to their respective conditions. (A) After incubation for
72 hours, the cells were harvested and analyzed for vIL-6 in the supernatant. (B) PF-8
(f) and gpK8.1 (o) lytic protein expression in the viable cells.
tion of angiogenic factors from these cells.15,26 Acute exposure of
BCBL-1 cells to hypoxia alone was found to induce vIL-6
production approaching the levels induced with 6.4 nM TPA
(Figure 4A). When cells were exposed to hypoxia and treated with
6.4 nM TPA, there was a further increase in vIL-6 production over
that caused by each treatment alone (Figure 4A). The level of vIL-6
increased in parallel with the extent of lytic intracellular protein
expression as determined by flow cytometry (Figure 4B). In 3
different experiments with BC-3 cells, the level of vIL-6 increased
in response to acute hypoxia from an average of 245 pg/mL in
normoxia to 611 pg/mL in hypoxia.
Effect of hypoxia-inducible-factor–inducing agents Dfx and
CoCl2 on KSHV protein expression
One of the primary mediators of the response of cells to low O2 is
hypoxia-inducible factor 1 alpha (HIF-1␣). Under normal conditions, HIF-1␣ is rapidly degraded, but this process is inhibited
under hypoxic conditions.27 To investigate the ability of increased
HIF-1␣ to reactivate KSHV replication, we tested the effects of 2
commonly used inducers of HIF-1␣, Dfx and CoCl2, on KSHVinfected cells.28 In a dose-dependent manner, CoCl2 treatment in
normoxic conditions increased the percentage of BCBL-1 cells
Figure 3. Western blot detection of the mCP of KSHV from supernatants of
KSHV-infected cells in normoxia (21% O2) and hypoxia (1% O2) in the absence
or presence of TPA induction. (A) 96-hour supernatants were collected from BC-3
or BCBL-1 cells that had been cultured for at least 3 passages at 21% O2 or 1% O2,
washed, seeded at 300 000 cells per milliliter, and returned to their respective
conditions. For BC-3, the band represents approximately 5.5 ⫻ 107 viral particles per
milliliter of supernatant. For BCBL-1, the band represents approximately 10 ⫻ 107
viral particles per milliliter. These values are based on comparison with a viral
standard electrophoresed on the same blot. (B) BCBL-1 cells cultured at 300 000
cells per milliliter in normoxia (21% O2) or acute hypoxia (1% O2) were then treated
with 6.4 nM TPA. The time indicated represents the time following TPA addition after
incubation of cells overnight. Each band was scanned, and the relative band intensity
expressed as total pixels was determined and plotted for samples in normoxia (o and
top band) and hypoxia (f and bottom band).
Figure 5. Dose-dependent expression of PF-8 and gpK8.1 in CoCl2-treated
BCBL-1 cells in normoxia. BCBL-1 cells, treated with increasing concentrations of
CoCl2 and harvested 72 hours after seeding, were analyzed for PF-8 (u) and gpK8.1
(o) protein expression. The plotted values represent the mean percentage of positive
cells ⫾ SD of 3 experiments.
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3248
DAVIS et al
expressing PF-8 and gpK8.1 (Figure 5); at 200 ␮M CoCL2, the
level of antigen expression was close to that obtained with cells
exposed to 30 nM TPA21 (compare Figure 2). CoCl2 also enhanced
the lytic protein expression induced by 6.4 nM TPA up to 2-fold
and increased vIL-6 levels in culture supernatants (data not shown).
Although less effective than CoCl2, treatment of BCBL-1 cells with
Dfx increased PF-8 protein expression from an average of 0.43%
for untreated cells to 0.88% and 1.2% for concentrations of 5 ␮M
and 10 ␮M, respectively. Concentrations above 200 ␮M CoCl2 and
20 ␮M Dfx led to substantial toxicity (less than 40% viability), and
therefore samples were not measured for lytic protein expression.
Discussion
Although KSHV can be activated by phorbol esters in the
laboratory, the physiologic stimuli involved in the reactivation of
KSHV from its latent state in B cells, endothelial cells, and other
target cells in the body are poorly understood. Our studies indicate
that hypoxia (1% O2) can activate lytic replication of KSHV in 2
different PEL B-cell lines and increase the expression of the
proangiogenic factor vIL-6. Some activation was observed with a
higher O2 saturation (5% O2). An O2 saturation as low as 2.5% can
be found in certain tissues under physiological conditions,29 and a
saturation of 1% or lower can be found under certain pathologic
conditions,30,31 including vascular ischemia of the legs.32 This
suggests that hypoxia-induced KSHV activation may be important
in the life cycle of this virus and contribute to KS pathogenesis. In
addition, we found that hypoxia potentiates TPA-induced KSHV
activation, suggesting that this stimulus may work in concert with
other inducers of KSHV replication. We have not determined if
endothelial cells infected with HHV-8 would respond similarly to
hypoxia, and therefore future studies on these cells will be required
to determine the extent of involvement of hypoxia in HHV-8
pathogenesis.
Although the percentage of cells induced to undergo lytic replication
by hypoxia is less than the maximal level induced by phorbol esters, it
represents a median increase of up to 22-fold over the background
replication obtained under normoxic conditions. It is worth noting that
our data, as well as previous studies,21 suggest that there may be a limit
to the percentage of cells that can be activated into the lytic phase. It was
previously reported that a maximum of approximately 15% live
BCBL-1 cells can be induced into the lytic phase and that for BC-3 cells
the maximum is closer to 20% to 25%. The reason a substantially higher
percentage of live cells do not undergo lytic replication with hypoxia
alone is unclear, but could be related to a cell-cycle–related restriction of
KSHV reactivation. Also, although hypoxia enhanced TPA-induced
replication, we found that combining a high level of TPA (eg, 30 nM)
and hypoxia led to significant cellular toxicity. To what extent this
represents direct cellular toxicity as a result of the combined effects of
hypoxia and TPA, as opposed to cell death from increased KSHV lytic
replication under these conditions, is not clear at this time.
Hypoxia induces a number of effects through an increase in the
intracellular levels of HIF-1␣. Under normoxic conditions, this protein
is degraded by the ubiquitin proteosome system in conjunction with the
von Hippel–Lindau gene product, but under hypoxic conditions, this
process is inhibited.33,34 We hypothesized that increased levels of
HIF-1␣ might mediate the reactivation of KSHV. This hypothesis was
supported by our results showing that 2 different chemicals, Dfx and
CoCl2, that are known to mimic hypoxia by raising levels of HIF-1␣
were both found to induce KSHV replication when added to cells grown
BLOOD, 15 MAY 2001 䡠 VOLUME 97, NUMBER 10
under standard laboratory conditions. Moreover, the level of induction
by CoCl2 exceeded the level observed in cells exposed to 1% O2. It is
known that HIF-1␣ reaches one half of maximal levels in cells exposed
to 1% O2, with higher levels being reached as the O2 level is lowered
below 1%.29 Thus, O2 levels below 1% might further increase the level
of KSHV lytic induction. HIF-1␣ mediates cellular effects by binding to
hypoxia-responsive elements (HREs) in the cellular genome. Analysis
of the KSHV genome reveals the existence of several regions that
contain the HRE consensus sequence (G/C/T)ACGTGC(G/T)35 upstream of KSHV ORFs. In some regions of the genome, there is also a
sequence with similarity to the identified HIF-enhancer sequence
TCCTCTT36 between the HREs and the coding region. These regions
include the immediate early protein encoded by ORF 57 and the lytic
genes encoded by ORF 26, 29b, K10, and 64. In addition, the HRE
consensus sequence is found upstream of ORF 50, a gene whose
expression has been reported to be required for reactivation of HHV-8
lytic replication in B-cells.37 It therefore remains possible that the direct
binding of HIF-1 activates these genes and/or others leading to HHV-8
lytic replication. This matter is currently under investigation. Alternatively, it is possible that KSHV is indirectly activated by a cellular
response to hypoxia, such as VEGF production. However, we did not
detect KSHV activation when VEGF was exogenously added to the
PEL cells. Also, hypoxia did not result in a substantial increase in the
already high constitutive level of VEGF production by the BC-3 cells.
Taken together, these results suggest that the increase in KSHV
reactivation induced by hypoxia was not mediated through stimulation
of VEGF under these conditions. However, it remains possible that other
indirect mechanisms may be mediating this effect.
KSHV encodes a number of proangiogenic human factors,
including vIL-6, viral macrophage inflammatory protein 1 (vMIPI), and vMIP-II, that are thought to be involved in the pathogenesis
of KS. In addition, ORF 74 of KSHV encodes a constitutively
active G-protein–coupled receptor that stimulates the production of
cellular proangiogenic factors, and by these means, the virus could
induce the proliferation of vascular endothelial cells.38,39 These
viral-encoded proteins are all produced or increased during lytic
KSHV replication.24 Thus, hypoxic conditions such as may occur at
the edges of a wound may induce both lytic KSHV replication and
the growth of target endothelial cells. This may provide a mechanism to aid the establishment of infection if this virus enters the
body through a cut in the skin or mucosal tissue. Hypoxic
conditions may activate the virus and contribute to the development of KS after localized infection of cells by KSHV.
There is some evidence to indicate that hypoxia may affect the
replication of certain other viruses as well.40 In particular, it has
been reported that replication of herpes simplex virus type 2 can be
increased 3- to 5-fold by exposure of infected cells to low O2
concentrations.40 Also, there is some evidence that production of
Coxsackie virus can be either decreased or increased by exposure
to hypoxia and that expression of HBx protein by hepatitis
B–infected hepatoma cells is enhanced by hypoxia.41 What is
noteworthy about KSHV, however, is the combination of viral
induction by hypoxia and hypoxia-induced production of proangiogenic factors that could promote the growth of endothelial
target cells.
The findings in this study may also explain certain clinical and
epidemiological features of KS. Although there are reports that
advanced KS may involve a monoclonal proliferation,42 the best
available evidence suggests that spindle cell proliferation induced
directly and/or indirectly by KSHV-encoded genes plays an
From www.bloodjournal.org by guest on June 17, 2017. For personal use only.
BLOOD, 15 MAY 2001 䡠 VOLUME 97, NUMBER 10
HYPOXIC INDUCTION OF KSHV
important role in the development and progression of this condition.43-45 Hypoxia-induced KSHV reactivation may contribute to
the increased prevalence of KS in patients with diabetes mellitus,46,47 as this condition is often associated with vascular disease
and poor circulation. It may also contribute to the tendency of KS
to occur on the lower legs, especially in the elderly men who
develop classic KS. Hypoxic tissues may also produce VEGF as a
direct result of hypoxia, and this may work in tandem with
hypoxia-induced KSHV replication to promote spindle cell growth.
Additional studies will be needed to further understand how these
factors may interact in the pathogenesis of KS.
Activation of lytic KSHV infection by hypoxia may also help
explain the pattern of distribution of KSHV infection and KS in the
world. Unlike other human herpesviruses, which infect almost all
individuals worldwide, the prevalence of KSHV infection varies
widely from region to region.9,48,49 Interestingly, the prevalence of
KSHV infection and KS is especially high in areas of the world
where malaria is still endemic (eg, sub-Saharan Africa) or where a
subset of the population has hereditary anemias, such as thalasse-
3249
mia, that confer partial protection against malaria (eg, southern
Italy and other Mediterranean areas).4,50,51 It is possible that tissue
hypoxia in patients with malaria or hereditary animas leads to
increased shedding of KSHV and spread of the virus to other
individuals.
In summary, our study provides evidence that KSHV can be
activated by exposure of latently infected B-cell lines to hypoxia
and that hypoxia can potentiate activation of KSHV by TPA. This
finding may be important in the epidemiology and life cycle of
KSHV and in the pathogenesis of KS. Further study of this
phenomenon may lead to strategies to prevent or treat KSHVrelated diseases.
Acknowledgments
We thank Isaac Rodriguez-Chavez for critical reading of the
manuscript and Seth M. Steinberg and Jodi Black for assistance
and advice.
References
1. Longo DL, Steis RG, Lane HC, et al. Malignancies in the AIDS patient: natural history, treatment
strategies, and preliminary results. Ann N Y Acad
Sci. 1984;437:421-430.
2. Dupin N, Fisher C, Kellam P, et al. Distribution of
human herpesvirus-8 latently infected cells in Kaposi’s sarcoma, multicentric Castleman’s disease, and primary effusion lymphoma. Proc Natl
Acad Sci U S A. 1999;96:4546-4551.
3. Antman K, Chang Y. Kaposi’s sarcoma. N Engl
J Med. 2000;342:1027-1038.
4. Iscovich J, Boffetta P, Franceschi S, Azizi E, Sarid
R. Classic Kaposi sarcoma: epidemiology and
risk factors. Cancer. 2000;88:500-517.
5. Stein ME, Spencer D, Ruff P, Lakier R, MacPhail
P, Bezwoda WR. Endemic African Kaposi’s sarcoma: clinical and therapeutic implications: 10year experience in the Johannesburg Hospital
(1980-1990). Oncology. 1994;51:63-69.
6. Friedman-Kien AE, Saltzman BR. Clinical manifestations of classical, endemic African, and epidemic AIDS-associated Kaposi’s sarcoma. J Am
Acad Dermatol. 1990;22:1237-1250.
7. Chang Y, Cesarman E, Pessin MS, et al. Identification of herpesvirus-like DNA sequences in
AIDS-associated Kaposi’s sarcoma. Science.
1994;266:1865-1869.
8. Whitby D, Howard MR, Tenant-Flowers M, et al.
Detection of Kaposi sarcoma associated herpesvirus in peripheral blood of HIV-infected individuals and progression to Kaposi’s sarcoma. Lancet.
1995;346:799-802.
9. Gao SJ, Kingsley L, Li M, et al. KSHV antibodies
among Americans, Italians and Ugandans with
and without Kaposi’s sarcoma. Nat Med. 1996;2:
925-928.
10. Cesarman E, Chang Y, Moore PS, Said JW,
Knowles DM. Kaposi’s sarcoma-associated herpesvirus-like DNA sequences in AIDS-related
body-cavity-based lymphomas. N Engl J Med.
1995;332:1186-1191.
11. Moore PS, Chang Y. Detection of herpesvirus-like
DNA sequences in Kaposi’s sarcoma in patients
with and without HIV infection. N Engl J Med.
1995;332:1181-1185.
12. Chuck S, Grant RM, Katongole-Mbidde E, Conant M, Ganem D. Frequent presence of a novel
herpesvirus genome in lesions of human immunodeficiency virus-negative Kaposi’s sarcoma.
J Infect Dis. 1996;173:248-251.
13. Boshoff C, Endo Y, Collins PD, et al. Angiogenic
and HIV-inhibitory functions of KSHV-encoded
chemokines. Science. 1997;278:290-294.
14. Nicholas J, Ruvolo VR, Burns WH, et al. Kaposi’s
sarcoma-associated human herpesvirus-8 encodes homologues of macrophage inflammatory
protein-1 and interleukin-6. Nat Med. 1997;3:287292.
15. Neipel F, Albrecht JC, Ensser A, et al. Human
herpesvirus 8 encodes a homolog of interleukin-6. J Virol. 1997;71:839-842.
16. Moore PS, Boshoff C, Weiss RA, Chang Y. Molecular mimicry of human cytokine and cytokine
response pathway genes by KSHV. Science.
1996;274:1739-1744.
17. Kaposi M. Idiopathic multiple pigmented sarcoma
of the skin. CA Cancer J Clin. 1982;32:342-347.
18. Ubbink DT, Tulevski II, den Hartog D, Koelemay
MJ, Legemate DA, Jacobs MJ. The value of noninvasive techniques for the assessment of critical
limb ischaemia. Eur J Vasc Endovasc Surg. 1997;
13:296-300.
19. Renne R, Zhong W, Herndier B, et al. Lytic
growth of Kaposi’s sarcoma-associated herpesvirus (human herpesvirus 8) in culture. Nat Med.
1996;2:342-346.
20. Arvanitakis L, Mesri EA, Nador RG, et al. Establishment and characterization of a primary
effusion (body cavity-based) lymphoma cell
line (BC-3) harboring Kaposi’s sarcoma-associated herpesvirus (KSHV/HHV-8) in the absence
of Epstein-Barr virus. Blood. 1996;88:26482654.
21. Zoeteweij JP, Eyes ST, Orenstein JM, et al. Identification and rapid quantification of early- and
late-lytic human herpesvirus 8 infection in single
cells by flow cytometric analysis: characterization
of antiherpesvirus agents. J Virol. 1999;73:58945902.
22. Davis DA, Humphrey RW, Newcomb FM, et al.
Detection of serum antibodies to a Kaposi’s sarcoma-associated herpesvirus-specific peptide.
J Infect Dis. 1997;175:1071-1079.
23. Aoki Y, Yarchoan R, Braun J, Iwamoto A, Tosato
G. Viral and cellular cytokines in AIDS-related
malignant lymphomatous effusions. Blood. 2000;
96:1599-1601.
24. Sarid R, Flore O, Bohenzky RA, Chang Y, Moore
PS. Transcription mapping of the Kaposi’s sarcoma-associated herpesvirus (human herpesvirus 8) genome in a body cavity-based lymphoma
cell line (BC-1). J Virol. 1998;72:1005-1012.
25. Aoki Y, Tosato G. Role of vascular endothelial
growth factor/vascular permeability factor in the
pathogenesis of Kaposi’s sarcoma-associated
herpesvirus-infected primary effusion lymphomas. Blood. 1999;94:4247-4254.
26. Molden J, Chang Y, You Y, Moore PS, Goldsmith MA. A Kaposi’s sarcoma-associated herpesvirus-encoded cytokine homolog (vIL-6) activates signaling through the shared gp130
receptor subunit. J Biol Chem. 1997;272:
19625-19631.
27. Semenza GL. Regulation of mammalian O2 homeostasis by hypoxia-inducible factor 1. Annu
Rev Cell Dev Biol. 1999;15:551-578.
28. Wang GL, Semenza GL. Desferrioxamine induces erythropoietin gene expression and hypoxia-inducible factor 1 DNA-binding activity: implications for models of hypoxia signal transduction.
Blood. 1993;82:3610-3615.
29. Jiang BH, Semenza GL, Bauer C, Marti HH.
Hypoxia-inducible factor 1 levels vary exponentially over a physiologically relevant range of
O2 tension. Am J Physiol. 1996;271:C1172–
1180.
30. Chang N, Goodson WH, Gottrup F, Hunt TK. Direct measurement of wound and tissue oxygen
tension in postoperative patients. Ann Surg.
1983;197:470-478.
31. Semenza GL. HIF-1: mediator of physiological
and pathophysiological responses to hypoxia.
J Appl Physiol. 2000;88:1474-1480.
32. Lantsberg L, Goldman M. Laser Doppler flowmetry, transcutaneous oxygen tension measurements and Doppler pressure compared in patients undergoing amputation. Eur J Vasc Surg.
1991;5:195-197.
33. Richard DE, Berra E, Pouyssegur J. Angiogenesis: how a tumor adapts to hypoxia. Biochem
Biophys Res Commun. 1999;266:718-722.
34. Semenza GL. Perspectives on oxygen sensing.
Cell. 1999;98:281-284.
35. Semenza GL, Roth PH, Fang HM, Wang GL.
Transcriptional regulation of genes encoding glycolytic enzymes by hypoxia-inducible factor 1.
J Biol Chem. 1994;269:23757-23763.
36. Liu Y, Cox SR, Morita T, Kourembanas S. Hypoxia regulates vascular endothelial growth factor gene expression in endothelial cells: identification of a 5⬘ enhancer. Circ Res. 1995;77:638643.
37. Lukac DM, Kirshner JR, Ganem D. Transcriptional activation by the product of open reading
From www.bloodjournal.org by guest on June 17, 2017. For personal use only.
3250
BLOOD, 15 MAY 2001 䡠 VOLUME 97, NUMBER 10
DAVIS et al
frame 50 of Kaposi’s sarcoma-associated herpesvirus is required for lytic viral reactivation in B
cells. J Virol. 1999;73:9348-93461.
38. Masood R, Cai J, Zheng T, Smith DL, Naidu Y,
Gill PS. Vascular endothelial growth factor/vascular permeability factor is an autocrine growth factor for AIDS-Kaposi sarcoma. Proc Natl Acad Sci
U S A. 1997;94:979-984.
39. Bais C, Santomasso B, Coso O, et al. G-proteincoupled receptor of Kaposi’s sarcoma-associated
herpesvirus is a viral oncogene and angiogenesis
activator. Nature. 1998;391:86-89.
40. Ebbesen P, Toth FD, Villadsen JA, Norskov-Lauritsen N. In vitro interferon and virus production at
in vivo physiologic oxygen tensions. In Vivo.
1991;5:355-358.
41. Lee SW, Lee YM, Bae SK, Murakami S, Yun Y,
Kim KW. Human hepatitis B virus X protein is a
possible mediator of hypoxia-induced angiogen-
esis in hepatocarcinogenesis. Biochem Biophys
Res Commun. 2000;268:456-461.
42. Rabkin CS, Janz S, Lash A, et al. Monoclonal origin of multicentric Kaposi’s sarcoma lesions.
N Engl J Med. 1997;336:988-993.
43. Ensoli B, Barillari G, Gallo RC. Cytokines and
growth factors in the pathogenesis of AIDS-associated Kaposi’s sarcoma. Immunol Rev. 1992;
127:147-155.
44. Yarchoan R. Therapy for Kaposi’s sarcoma: recent advances and experimental approaches. J
Acquir Immune Defic Syndr. 1999;21(suppl 1):
S66–S73.
45. Miles SA, Rezai AR, Salazar-Gonzalez JF, et al.:
AIDS Kaposi sarcoma-derived cells produce and
respond to interleukin 6. Proc Natl Acad Sci U S
A. 1990;87:4068-4072.
46. Laor Y, Schwartz RA. Epidemiologic aspects of
American Kaposi’s sarcoma. J Surg Oncol. 1979;
12:299-303.
47. Friedman-Birnbaum R, Weltfriend S, Katz I. Kaposi’s sarcoma: retrospective study of 67 cases
with the classical form. Dermatologica. 1990;180:
13-17.
48. Cook PM, Whitby D, Calabro ML, et al. Variability
and evolution of Kaposi’s sarcoma-associated
herpesvirus in Europe and Africa. International
Collaborative Group. AIDS. 1999;13:1165-1176.
49. Simpson GR, Schulz TF, Whitby D, et al. Prevalence of Kaposi’s sarcoma associated herpesvirus infection measured by antibodies to recombinant capsid protein and latent immunofluorescence antigen. Lancet. 1996;348:1133-1138.
50. Sarid R, Olsen SJ, Moore PS. Kaposi’s sarcomaassociated herpesvirus: epidemiology, virology,
and molecular biology. Adv Virus Res. 1999;52:
139-232.
51. Franceschi S, Geddes M. Epidemiology of classic
Kaposi’s sarcoma, with special reference to Mediterranean population. Tumori. 1995;81:308-314.
From www.bloodjournal.org by guest on June 17, 2017. For personal use only.
2001 97: 3244-3250
doi:10.1182/blood.V97.10.3244
Hypoxia induces lytic replication of Kaposi sarcoma−associated herpesvirus
David A. Davis, Andrea S. Rinderknecht, J. Paul Zoeteweij, Yoshiyasu Aoki, Elizabeth L. Read-Connole,
Giovanna Tosato, Andrew Blauvelt and Robert Yarchoan
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