A Proapoptotic Peptide Derived from
Reovirus Outer Capsid Protein µ1 Has
Membrane-Destabilizing Activity
Jae-Won Kim, Sangbom M. Lyi, Colin R. Parrish and John
S. L. Parker
J. Virol. 2011, 85(4):1507. DOI: 10.1128/JVI.01876-10.
Published Ahead of Print 24 November 2010.
These include:
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JOURNAL OF VIROLOGY, Feb. 2011, p. 1507–1516
0022-538X/11/$12.00 doi:10.1128/JVI.01876-10
Copyright © 2011, American Society for Microbiology. All Rights Reserved.
Vol. 85, No. 4
A Proapoptotic Peptide Derived from Reovirus Outer Capsid Protein
␮1 Has Membrane-Destabilizing Activity䌤
Jae-Won Kim, Sangbom M. Lyi, Colin R. Parrish, and John S. L. Parker*
Baker Institute for Animal Health, College of Veterinary Medicine, Cornell University, Ithaca, New York 14853
Received 3 September 2010/Accepted 10 November 2010
stabilize membranes, and this region of ␮1 was previously
suggested to be involved in membrane penetration during virus
entry (20). Based on these previous findings, we hypothesized
that one mechanism by which ␮1 induces apoptosis is by direct
destabilization of intracellular membranes. Such destabilization could induce a cellular stress response or lead to direct
leakage of proapoptotic molecules from mitochondria or the
endoplasmic reticulum (ER).
Here, we examined the capacity of peptides representing
residues 582 to 611 of ␮1 to perturb artificial and cellular
membranes.
Mammalian orthoreoviruses cause pathology in newborn
mice by inducing apoptosis in infected cells (2). In addition,
type 3 Dearing mammalian orthoreoviruses (Reolysin) are currently in phase II/III trials for treatment of a variety of cancers
(17, 28). Tumor killing by reovirus infection involves effects
that are both direct and indirect (e.g., activation of a tumorspecific immune response) (25). However, infected tumor cells
undergo virus-induced apoptosis (26). Although much is now
known about the specific apoptotic and signaling pathways that
are activated in reovirus-infected cells, the underlying mechanism of how the virus initiates apoptosis is not well understood.
The ␮1 protein is the major determinant of reovirus-induced
apoptosis, and ectopic expression of ␮1 in cells induces apoptosis (3, 5, 6). Apoptosis induced by ectopically expressed ␮1 is
abrogated by coexpression of outer capsid protein ␴3, with
which ␮1 assembles to form heterohexamers that form the
major portion of the outer capsid of the virus (3). Residues 582
to 611 of ␮1 are necessary and sufficient for apoptosis induction, and when fused to enhanced green fluorescent protein
(EGFP), this fragment of ␮1 predominantly localizes to mitochondrial membranes (3). In addition, recombinant T3D viruses with substitutions in this region of ␮1 (K594D or I595K)
are defective for induction of apoptosis and have attenuated
virulence in newborn mice (5). How this particular region of
␮1 functions to initiate apoptosis is unknown. In the atomic
resolution structure of the ␮1:␴3 heterohexamer, this region of
␮1 forms a short loop followed by an amphipathic ␣-helix;
residues 584 and 585 lie within the amphipathic ␣-helix (19).
Amphipathic peptides are known to target and sometimes de-
MATERIALS AND METHODS
Cells. L929 cells were maintained in suspension in Joklik’s modified minimal
essential medium supplemented with 4% fetal bovine serum, 2 mM glutamine,
100 U ml⫺1 penicillin, and 100 ␮g ml⫺1 streptomycin. CV-1 cells were grown in
Ham’s F-12 medium (CellGro) supplemented with 10% fetal bovine serum
(HyClone), 100 U ml⫺1 penicillin, 100 ␮g ml⫺1 streptomycin, 1 mM sodium
pyruvate, and nonessential amino acids (CellGro). CHO-K1 cells were grown in
Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% fetal
bovine serum, 100 U ml⫺1 penicillin, and 100 ␮g ml⫺1 streptomycin.
Peptide synthesis and CD. Peptides were synthesized by FlexPeptide technology, purified by high-performance liquid chromatography (HPLC) and confirmed by mass spectrometry at GenScript (Piscataway, NJ). Circular dichroism
(CD) was used to analyze the secondary structure of the synthetic peptides.
Initially, the peptides were analyzed at 50 ␮M in trifluorethanol or in the
presence of 10% phosphatidylcholine (PC) liposomes (prepared as described
below). Spectra were recorded with a model 400 AVIV CD spectrometer (Lakewood, NJ) in a 1-cm-path-length cuvette at room temperature (RT). Structural
analysis was performed using the Delta epsilon (DE) calculation method using
K2D CD spectra Deconvolution software (http://www.embl.de/⬃andrade/k2d
.html).
Cytotoxicity assay. We used a fluorescence-based cytotoxicity assay (CytoToxFluor; Promega, Madison, WI) to quantify cytotoxicity induced by incubation
with the synthetic peptides. L929 cells were seeded in 96-well plates and incubated with the indicated concentrations of each peptide for 1 h at 37°C in 100 ␮l
of DMEM, and then the levels of cytotoxicity were measured following incubation with the fluorescent substrate (100 ␮l) for an additional 30 min at 37°C
according to the manufacturer’s instructions. Fluorescence was measured by
* Corresponding author. Mailing address: Baker Institute for Animal Health, College of Veterinary Medicine, Cornell University, Hungerford Hill Road, Ithaca, NY 14853. Phone: (607) 256-5626. Fax:
(607) 256-5608. E-mail: [email protected].
䌤
Published ahead of print on 24 November 2010.
1507
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The reovirus outer capsid protein ␮1 is responsible for cell membrane penetration during virus entry and
contains determinants necessary for virus-induced apoptosis. Residues 582 to 611 of ␮1 are necessary and
sufficient for reovirus-induced apoptosis, and residues 594 and 595 independently regulate the efficiency of
viral entry and reovirus-induced cell apoptosis, respectively. Two of three ␣-helices within this region, helix 1
(residues 582 to 611) and helix 3 (residues 644 to 675), play a role in reovirus-induced apoptosis. Here, we
chemically synthesized peptides representing helix 1 (H1), H1:K594D, H1:I595K, and helix 3 (H3) and
examined their biological properties. We found that H1, but not H3, was able to cause concentration- and
size-dependent leakage of molecules from small unilamellar liposomes. We further found that direct application of H1, but not H1:K594D, H1:I595K, or H3, to cells resulted in cytotoxicity. Application of the H1 peptide
to L929 cells caused rapid elevations in intracellular calcium concentration that were independent of phospholipase C activation. Cytotoxicity of H1 was not restricted to eukaryotic cells, as the H1 peptide also had
bactericidal activity. Based on these findings, we propose that the proapoptotic function of the H1 region of ␮1
is dependent on its capacity to destabilize cellular membranes and cause release of molecules from intracellular organelles that ultimately induces cell necrosis or apoptosis, depending on the dose.
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KIM ET AL.
the coverslips were mounted on glass slides with Prolong Gold plus DAPI
(4⬘,6-diamidino-2-phenylindole; Invitrogen). Images were obtained with an inverted microscope (Nikon TE2000) equipped with fluorescence optics through a
60⫻ 1.4-numerical-aperture (NA) oil objective with 1.5⫻ optical zoom. Images
were collected digitally with a Coolsnap HQ charge-coupled-device camera
(Roper) and Openlab software (Improvision) and then prepared for publication
using Photoshop and Illustrator software (Adobe).
To assess the distribution of biotinylated H1 in cells, CHO-K1 cells seeded on
50-mm MatTek gridded dishes (MatTek, Ashland, MA) were preincubated with
500 nM Mitotracker CMXRos (Invitrogen) in growth medium for 45 min at
37°C. The labeling medium was replaced with medium lacking phenol red, and
the cells were microinjected with ⬃6 nl of sample using an InjectMan NI2
microinjector and a Femtoject micromanipulator (Eppendorf, Hauppauga, NY).
Micropipette needles were prepared with a P-97 micropipette puller (Sutter
Instrument Co., Novato, CA). Samples injected were 50 ␮M biotin-conjugated
H1 that was preincubated with Alexa488-conjugated streptavidin or, as a control,
Alexa488-conjugated streptavidin. Samples were prepared in PBS buffer and
injected into cells, and the cells were incubated at 37°C on a stage warmer
(World Precision Instruments, Sarasota, FL). Fluorescence and phase-contrast
images were collected using a Nikon TE300 microscope and a Hamamatsu Orca
ER charge-coupled-device camera. Images were collected using Simple PCI
software (Hamamatsu, Sewickley, PA) and contrast and brightness adjusted with
Photoshop (Adobe, San Jose, CA).
To assess the release of cytochrome c from cells microinjected with the H1,
H1:I595K, or H3 peptide, CHO-K1 cells maintained at 37°C were microinjected
as described above with a 1:1 mixture of Alexa594 bovine serum albumin (BSA)
(Invitrogen) and the indicated peptides (final concentration, 50 ␮M). Twenty
minutes after microinjection, the cells were fixed in 4% paraformaldehyde in
PBS, washed in PBS, permeabilized with 0.1% Triton X-100 in PBS plus 1%
BSA, and then immunostained with anti-cytochrome c monoclonal antibody
(MAb) (clone 6H2.B4; BD Biosciences) followed by Alexa594-conjugated goat
anti-mouse IgG. Fluorescence and phase-contrast images were collected and
processed as described above.
Peptide bactericidal activity. The bactericidal activity of peptides against Escherichia coli and Bacillus subtilis JH641 was assayed by colony inhibition. Bacterial
cultures were grown to early log phase at 37°C in Luria-Bertani (LB) broth and
then diluted to 2 ⫻ 104 CFU/ml in 10 mM phosphate buffer (pH 7.4) containing
0.03% LB broth. Equal volumes of peptide-containing solution and bacterial
culture (25 ␮l of each) were mixed and incubated at 37°C for 3 h, after which the
bacterium-peptide mixture was diluted 100-fold or 1,000-fold with cold phosphate buffer and inoculated on LB agar plates, which were incubated at 37°C
overnight. The number of bacterial colonies on each plate was counted the
following day and expressed as a percentage of the number of colonies obtained
when phosphate buffer was mixed with the bacteria instead of peptide.
Preparation of isolated mitochondria and cytochrome c release assay. Cytochrome c released from mitochondria purified from murine L929 cells was
assayed as described previously, with some modification (32). Cells in spinner
culture were harvested, washed with cold PBS, and then resuspended at 3 ⫻ 107
ml⫺1 in ice-cold mitochondrial buffer (20 mM HEPES, pH 7.5, 250 mM sucrose,
1 mM EGTA, 1 mM EDTA, 100 mM KCl, 1 mM dithiothreitol [DTT]) containing 0.1 mM phenylmethylsulfonyl fluoride (PMSF), 2 ␮g ml⫺1 pepstatin, and
complete protease inhibitor (Complete Mini; Roche). Cells were lysed by
Dounce homogenization (80 strokes), and nuclei and intact cells were removed
by centrifugation (800 ⫻ g for 15 min at 4°C). The supernatant was then centrifuged at 10,000 ⫻ g for 15 min at 4°C. The mitochondrial pellet was further
washed and resuspended in mitochondrial buffer before being used immediately
for further experiments. Both the mitochondrion-free supernatant and the mitochondrial pellet were saved for analysis. Isolated mitochondria were incubated
with 100 ␮M the H1 or H3 peptide, treated with 1% Triton X-100, or incubated
with an equal volume of PBS for 20 min at RT. After incubation, the samples
were returned to ice and the mitochondrial and supernatant fractions were
separated by centrifugation (16,000 ⫻ g for 15 min at 4°C). The pellet fraction
was resuspended in a volume of mitochondrial buffer equal to that of the
supernatant. Samples were then incubated in protein sample buffer with 200 mM
DTT on ice for 15 min and then at 80°C for 15 min, cooled on ice, and cleared
at 16,000 ⫻ g for 1 min, and then proteins were separated on 15% SDS-PAGE.
Proteins were transferred to nitrocellulose membranes by Semi-Dry Transfer
Cell (Bio-Rad). The membrane was blocked with 5% BSA in Tris-buffered
saline–Tween 20 (TBST) buffer, then incubated with anti-cytochrome c MAb
(1:1,000; 7H8.2C12; BD Biosciences) overnight at 4°C, washed, and then incubated with horseradish peroxidase (HRP)-conjugated donkey anti-mouse IgG
(1:10,000; Jackson Laboratories). Proteins were detected on autoradiograph film
with SuperSignal West Pico chemiluminescent substrate (Thermo Scientific).
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spectrofluorimetry (excitation wavelength [␭ex] ⫽ 485 nm; emission wavelength
[␭em] ⫽ 520 nm), and cytotoxicity was calculated by the following equation:
cytotoxicity ⫽ (mean for treated cells ⫺ mean for untreated cells)/standard
deviation (SD) for cell-free controls. Maximum cytotoxicity (100%) was set as
the fluorescence measured in cells treated with 30 ␮g ml⫺1 digitonin for 1 h at
37°C.
Hemolysis assay. Hemolysis assays were performed as described previously
(34). In brief, washed citrated chicken red blood cells (RBCs) in Alsevers solution (Colorado Serum Co., Denver, CO) were suspended in cold phosphatebuffered saline (PBS) containing 2 mM MgCl2 (PBS-Mg) at a concentration of
5% (vol/vol) with different concentrations of the indicated peptides. After incubation for 30 min at 37°C, the RBCs were pelleted by centrifugation (380 ⫻ g for
5 min) and released hemoglobin in the supernatant was measured by absorbance
at 415 nm. Supernatant collected from RBCs incubated with 0.1% Triton X-100
was used to assess complete release of hemoglobin, and RBCs incubated with
PBS were used as a negative control. Data shown are the means ⫾ SD of results
from a minimum of three independent experiments.
Preparation of liposomes and liposome leakage assays. Phosphatidylcholine
(PC) in chloroform (18:1; Avanti Lipids) was dried under nitrogen, rehydrated at
1 mg ml⫺1 in PBS, pH 7.4, containing 12.5 mM aminonaphthalene trisulfonic
acid (ANTS; Fluka) and 45 mM p-xylene-bis (N-pyridinium bromide) (DPX;
Sigma), and then frozen and thawed (10⫻) in liquid nitrogen. Unilamellar
liposomes (50-nm diameter) were prepared by extrusion (Lipex extruder; Northern lipids, Inc.). Similar PC liposomes (200-nm diameter) were prepared in PBS
containing 300 mg ml⫺1 fluorescein isothiocyanate (FITC)-conjugated dextran
(FD) with molecular weights of 40,000 (FD40) or 10,000 (FD10) (Sigma). Unincorporated ANTS-DPX or FD was separated from liposomes by mixing the
liposomes 1:1 (vol/vol) in 70% sucrose, overlaying them with 20% sucrose, and
then centrifuging them at 96,800 ⫻ g for 15 h at 4°C. Liposomes were harvested
from the top of the gradient and then reequilibrated in PBS by gel filtration on
a Sephadex G-25 PD10 column (Amersham Biosciences). Dequenching of
ANTS in response to liposome leakage was assayed by spectrofluorimetry (Cary
Eclipse spectrophotometer; Varian, Inc.) (␭ex ⫽ 355 nm; ␭em ⫽ 520 nm). The
percent leakage was calculated using the baseline fluorescence emission of
ANTS-DPX-loaded liposomes in buffer only (F0), the sample fluorescence (FS),
and the 100% leakage fluorescence (FT) (liposomes were lysed with 0.2% Triton
X-100) as follows: percent leakage ⫽ [(FS ⫺ F0)/(FT ⫺ F0)] ⫻ 100. Leakage of
FD40 and FD10 from liposomes was also assayed by dequenching (␭ex ⫽ 490 nm;
␭em ⫽ 500 to 540 nm). Maximum leakage of FD was assayed after addition of
0.2% Triton X-100; in control experiments, we found that 0.2% Triton X-100 was
required to fully lyse liposomes.
We prepared liposomes with the lipid composition of Xenopus mitochondria
using egg phosphatidylcholine (PC; 47 mol%), egg phosphatidylethanolamine
(PE; 28 mol%), soybean phosphatidylinositol (PI; 9 mol%), and brain phosphatidylserine (PS; 9 mol%) mixed with 7 mol% of heart cardiolipin (CL) (Avanti
Polar Lipids) as described in the text. For lower concentrations of cardiolipin, the
mol% of PC was increased. The lipid mixture was dried and resuspended in PBS
containing ANTS-DPX as described above. Liposome extrusion was as described
above, with final extrusion through a 50-nm filter membrane.
Measurement of intracellular free Ca2ⴙ concentration. We assessed changes
in intracellular free Ca2⫹ levels in L929 cells. Spinner culture L929 cells (approximately 3 ⫻ 107 cells) were harvested, washed in PBS buffer, incubated for
1 h at 37°C in the dark in 6 ml of serum-free Joklik’s modified minimal essential
medium containing 3 ␮M fura-2 acetoxymethyl ester (fura-2AM; Invitrogen) and
8 ␮M pluronic F-127 (Invitrogen), then washed twice with Hanks balanced salt
solution (HBSS) buffer (130 mM NaCl, 5 mM KCl, 1.8 mM CaCl2, 1.0 mM
MgCl2, 0.6 mM Na2HPO4, 0.4 mM KH2PO4, 5 mM glucose, 20 mM HEPES, pH
7.4), resuspended in HBSS containing 2.5 mM probenecid (Sigma), and further
incubated for 20 min at RT in the dark to allow esterification of fura-2AM. Following fura-2 loading, the cells were washed three times and resuspended in 3 ml of
HBSS containing 1 mM EGTA immediately prior to measurements. Cell suspensions (400 ␮l) were placed in a quartz cuvette with a magnetic stirrer. Experimental
reagents, including peptides, Triton X-100, carbachol, or phospholipase C (PLC)
inhibitor (U-73122; Sigma), were added at the time points indicated in Results.
Fluorescence was measured by spectrofluorimetry (␭ex ⫽ 340 and 380 nm; ␭em ⫽
510 nm). The ratio of the fura-2AM–Ca2⫹ complex (␭ex ⫽ 340 nm; ␭em ⫽ 510 nm)
to free fura-2AM (␭ex ⫽ 380 nm; ␭em ⫽ 510 nm) was generated using the Grynkiewicz equation to calculate internal calcium concentration (10).
Microscopy and microinjection. Biotin-conjugated H1 and H1:I595K peptides
were synthesized, dissolved in PBS, and incubated for 30 min at RT with CV-1
cells on 18-mm glass coverslips. The cells were washed extensively in PBS, then
fixed with 2% paraformaldehyde in PBS for 10 min, incubated with Alexa488
Fluor-conjugated streptavidin (Invitrogen), and washed again in PBS, and then
J. VIROL.
REOVIRUS ␮1 AND PEPTIDE MEMBRANE PERMEABILIZATION
VOL. 85, 2011
1509
Statistical analysis. All experiments were performed a minimum of three
times, and data are shown as mean ⫾ standard deviation (SD). Preparation of
graphs and statistical analysis was done using Prism GraphPad software.
RESULTS
FIG. 1. Linear representation and ribbon diagram of a monomer of
␮1 with sequence and CD spectra of the synthesized peptides. (A) The
different fragments of ␮1 generated during virus entry are illustrated in
a linear representation. The ␣-helical regions within the ␾ fragment of
␮1 are colored red (H1), yellow, and blue (H3). The amino acid
sequences of the H1, H1:K594D, H1:I595K, and H3 peptides are
shown. Numbers indicate amino acid positions within ␮1. (B) Circular
dichroism spectra of the synthetic peptides H1, H1:K594D, H1:I595K,
and H3. All of the peptides showed minima at 208 and 222 nm,
characteristic of alpha-helical secondary structure. Peptides (50 ␮M)
were solubilized in trifluoroethanol to prevent aggregation and to
stabilize peptide structure. Spectra were recorded at room temperature with a model 400 AVIV CD spectrometer (Lakewood, NJ).
type H1 peptide, H1:I595K did not induce liposome leakage.
The H1:K594D peptide did induce leakage, but ⬃2-fold-higher
concentrations than those of wild-type H1 were required to
induce 50% leakage (Fig. 2A). Thus, the capacity of ␮1, ␾, or
their I595K and K594D substitution mutants to induce apoptosis when expressed ectopically in cells correlates with the
capacities of peptides representing the region that is minimally
required for apoptosis induction to permeabilize artificial
membranes.
Synthetic H1 peptide causes size-selective leakage of FITCdextrans from liposomes. The ␮1N fragment of ␮1 (Fig. 1A) is
required for virus entry but is not involved in ␮1-mediated
Downloaded from http://jvi.asm.org/ on July 9, 2014 by Cornell University Library
A synthetic peptide representing residues 582 to 612 of ␮1 is
␣-helical in solution. We have previously shown that the Cterminal ␾ fragment (residues 582 to 708) of the reovirus outer
capsid ␮1 protein contains determinants necessary for virusinduced apoptosis. Residues 582 to 611 of ␮1 are necessary
and sufficient for apoptosis induction (3), and substitutions
K594D and I595K within this region in recombinant viruses
have reduced apoptosis-inducing capacity in vitro and in vivo
(5). In the structure of the ␮1:␴3 heterohexamer, residues 582
to 611 consist of a short loop and an amphipathic ␣-helix (Fig.
1A) (19).
Many amphipathic peptides interact with membranes and in
some cases destabilize membrane barriers (18). We previously
showed that residues 582 to 611 fused to EGFP localize to
mitochondria, lipid droplets, and the endoplasmic reticulum in
transfected cells (3). Destabilization of mitochondrial and/or
ER membranes could in part explain the mechanism by which
␮1 induces apoptosis. We therefore examined whether a synthetic peptide representing residues 582 to 611 could destabilize cellular and artificial membranes. Peptides representing
residues 582 to 612 of ␮1 (peptide H1) and, as a negative
control, a peptide representing residues 639 to 675 of ␮1 (peptide H3), which does not induce apoptosis when expressed in
cells as a fusion with EGFP (3), were chemically synthesized.
All of the peptides were soluble in aqueous solutions. We used
circular dichroism to identify the secondary structures of the
peptides in trifluorethanol (H1 and H3), which is known to
stabilize peptide structures. We found that the H1 and H3
peptides were ␣-helical in solution (100%) (Fig. 1B). The H1
and mutant peptides (K593D and I595K) had identical spectra,
while H3 had deeper and higher peaks, most likely because of
the greater length of H3 (Fig. 1B). We observed no significant
changes in the secondary structure conformers of the H1 peptide in the presence of liposomes (data not shown).
Peptide H1 (residues 582 to 611) directly permeabilizes liposomes. To examine if the H1 peptide could destabilize membranes, we prepared small unilamellar liposomes (⬃50-nm diameter) consisting of the neutral lipid phosphatidylcholine that
contained the fluorophore/quencher pair 8-aminonaphthalene
1,3,6-trisulfonic acid (ANTS)–p-xylene-bis-pyridinium (DPX).
We then assessed liposome leakage in response to increasing
concentrations of H1 by measuring dequenching of ANTS (1).
We found that H1 induced a concentration-dependent leakage
of ANTS from liposomes, with 50% leakage occurring at a
concentration of ⬃12.6 ␮M. In contrast, the H3 peptide had no
effect on liposome membrane permeability (Fig. 2A). We conclude that the H1 peptide has direct membrane-permeabilizing
properties.
Mutants of H1 lose the capacity to directly permeabilize
liposomes. We previously showed that recombinant viruses or
␮1 with the I595K or K594D substitution within the H1 helix
were defective in apoptosis induction (5). We therefore examined the capacity of peptides bearing these substitutions to
permeabilize PC liposomes. We found that unlike the wild-
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KIM ET AL.
apoptosis induction. Zhang et al. showed that a synthetic myristoylated ␮1N peptide likely forms size-selective pores in artificial membranes (34). To address the possibility that the H1
peptide also forms size-selective pores, we prepared liposomes
preloaded with self-quenching concentrations of 10- or 40-kDa
fluorescein isothiocyanate (FITC)-dextran polymers. Leakage
of the FITC-dextran molecules was assayed by fluorescence
dequenching (Fig. 2B). We found that concentrations of H1
that caused release of ANTS also caused release of 10 kDa
FITC-dextran; however, the release of 40 kDa FITC-dextran
was much less efficient (Fig. 2B). These findings indicate that
size-limited pores formed in liposomes treated with the H1
peptide.
The H1 peptide permeabilizes cell membranes. (i) H1 is
cytotoxic for L929 cells. Based on our finding that the H1
peptide directly permeabilized artificial membranes, we investigated whether incubation of H1 with cells was cytotoxic. Using an assay that quantifies release of protease activity from
dead cells (CytoTox-Fluor; Promega), we found that concentrations of H1 of 50 ␮M or greater were cytotoxic to L929 cells
after 1 h of incubation (Fig. 3A). In contrast, similar concentrations of the H3 peptide had no effect on the viability of L929
cells (Fig. 3A). The I595K and K594D mutant H1 peptides
were not cytotoxic, even at 200 ␮M (Fig. 3A).
(ii) H1 causes hemolysis of red blood cells. To further determine if the H1 peptide was directly permeabilizing the
plasma membranes of cells, we assayed for leakage of hemoglobin from chicken red blood cells. Unlike for the ␮1N peptide, where concentrations of 10 ␮g ml⫺1 led to 70% hemolysis,
we found that only 30% hemolysis occurred when chicken
RBCs were incubated with 20-fold-higher concentrations of
H1 peptide (Fig. 3B). This difference in efficiency of hemoglobin release (⬃68 kDa) likely reflects a lower size exclusion
limit of pores formed by the H1 peptide than of those formed
by ␮1N, which allow release of hemoglobin and 40-kDa FITCdextran (34).
(iii) H1 causes changes in cytosolic Ca2ⴙ levels. We hypothesized that the cytotoxicity that we observed in L929 cells was
caused by increases in cytosolic Ca2⫹ concentrations. In support of this hypothesis, we found that application of the H1
peptide to fura-2-loaded L929 cells induced substantial and
rapid increases in cytosolic [Ca2⫹], with 50 ␮M H1 causing a 3to 4-fold increase in cytosolic [Ca2⫹] by 2 min after addition of
the peptide. In contrast, application of 50 ␮M H3 had no
noticeable effect on cytosolic [Ca2⫹] (Fig. 3C). As these experiments were carried out with Ca2⫹-free buffer containing
EGTA, this finding suggested that in addition to having direct
membrane permeabilization effects, the H1 peptide might also
stimulate phospholipase C-mediated IP3 signaling pathways
with release of Ca2⫹ from intracellular stores. Addition of
purified rotavirus NSP4 or a peptide representing NSP4114-135
induces elevation of cytosolic calcium in Sf9 insect cells, which
is blocked by the phospholipase C inhibitor (U-73122) (29).
Treatment with U-73122 prevented the increase in Ca2⫹ induced by 100 ␮M carbachol, a PLC receptor agonist (Fig. 3D).
However, we found that pretreatment of cells with the PLC
inhibitor (U-73122) had no effect on H1-induced increases in
intracellular Ca2⫹ (Fig. 3D). Based on these findings, we conclude that the rapid changes in intracellular Ca2⫹ cannot be
attributed to direct permeabilization of the plasma membrane
alone. We speculate that the H1 peptide may access the intracellular membranes of the ER or mitochondria, causing the
release of calcium stores.
H1 peptide has antibacterial activity for Gram-negative and
Gram-positive bacteria. Based on the cytotoxicity of H1 for
L929 cells, the membrane-permeabilizing activity of H1, and
the propensity of cationic amphipathic helical peptides to have
antibacterial activity (33), we tested whether the H1 peptide
had antibacterial activity. We found dose-dependent inhibition
of the growth of Gram-positive (B. subtilis) and Gram-negative
(E. coli) bacteria by the H1 peptide (Fig. 4A), with complete
inhibition of colony formation in this assay at concentrations of
H1 greater than 25 ␮M. In contrast, 50 ␮M the H3 peptide had
little effect (Fig. 4B), and 50 ␮M concentrations of the K594D
and I595K substitution mutants had decreased antibacterial
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FIG. 2. Synthetic H1 but not mutant or H3 peptides cause release
of small molecules from phosphatidylcholine liposomes. (A) ANTSDPX liposome leakage assays of H1, H3, and mutant peptides. Peptides at the indicated concentrations were incubated with phosphatidylcholine (PC)-containing liposomes loaded with ANTS and DPX,
and leakage of ANTS was assayed by spectrofluorimetry (␭ex ⫽ 355
nm; ␭em ⫽ 520 nm). Means ⫾ SD of results from three independent
experiments are shown. The percent leakage was calculated using the
baseline fluorescence emission of ANTS-DPX-loaded liposomes in
buffer only (F0), the sample fluorescence (FS), and the 100% leakage
fluorescence (FT) (liposomes lysed with 0.2% Triton X-100) as follows:
percent leakage ⫽ [(FS ⫺ F0)/(FT ⫺ F0)] ⫻ 100. (B) Phosphatidylcholine (PC) liposomes containing self-quenching concentrations (300 mg
ml⫺1) of 10 or 40 kDa fluorescein isothiocyanate-conjugated dextran
(FD10 or FD40) were incubated with the indicated concentrations of
H1 peptide. Dequenching of FD was assayed by spectrofluorimetry
(␭ex ⫽ 490 nm; ␭em ⫽ 520 nm). The percent leakage was calculated
using the baseline fluorescence emission of FD-loaded liposomes in
buffer only (F0), the sample fluorescence (FS), and the 100% leakage
fluorescence (FT) (liposomes lysed with 0.2% Triton X-100) as follows:
percent leakage ⫽ [(FS ⫺ F0)/(FT ⫺ F0)] ⫻ 100. Means ⫾ SD of results
from three independent experiments are shown.
J. VIROL.
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1511
activity compared to the level for WT H1 (Fig. 4B). We conclude that the H1 peptide has antibacterial properties.
Effect of lipid composition on H1-induced ANTS leakage.
Green fluorescent protein (GFP)-fused H1 localizes prominently to mitochondria in transfected cells, causes release of
cytochrome c, and induces apoptosis (3, 31). Mitochondrial
membranes have increased concentrations of the neutral lipid
cardiolipin, and permeabilization of liposomes by the Bcl-2
homology domain 3 (BH3)-only peptides derived from proapoptotic proteins Bax and Bak is promoted by increased con-
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FIG. 3. The H1 peptide is cytotoxic and permeabilizes cellular membranes. (A) Different concentrations of H1, H1:K594D, H1:I595K, and H3
were incubated with L929 cells for 1 h at 37°C, and then the percentage cytotoxicity (maximum cytotoxicity was set as the fluorescence measured
in cells treated with 30 ␮g ml⫺1 digitonin for 1 h at 37°C) was assayed with Live/Dead fixable cell stain (Invitrogen) using a fluorescence plate
reader. Means ⫾ SD of results from 3 independent experiments are shown. (B) The indicated concentrations of peptides were incubated with
chicken RBCs at 37°C for 1 h, and hemolysis was assayed as described in Materials and Methods. (C) The indicated concentrations of H1 or H3
peptides were added (arrow) to L929 cells preloaded with 3 ␮M fura-2 acetoxymethyl ester (fura-2AM) in Ca2⫹-free Hanks buffered salt solution,
pH 7.4, in a stirred cuvette at 37°C in a spectrofluorimeter. Fluorescence measurements were collected before and after addition of peptides (␭ex ⫽
340 and 380 nm; ␭em ⫽ 510 nm). Change in cytosolic [Ca2⫹] (nM) was determined from emission data using the Grynkiewicz equation (10).
(D) The cytosolic calcium elevation in L929 induced by H1 is not mediated by a phospholipase C pathway. Fura-2AM-loaded L929 cells were
incubated with H1 (50 ␮M) or the M5 muscarinic agonist carbachol (100 ␮M) (left arrow; solid traces), and changes in cytosolic calcium levels
were monitored by spectrofluorimetry. Alternatively, H1 or carbachol was added to cells, together with 10 ␮M U-73122 (right arrow; dotted traces).
The traces shown are representative of three independent experiments.
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KIM ET AL.
centrations of cardiolipin (16). We therefore prepared liposomes with lipid contents that approximated those of Xenopus
mitochondria and examined the effect of the presence or absence of 7 mol% cardiolipin on the efficiency of H1 membrane
permeabilization. We found that the efficiency of H1 permeabilization of liposomes was increased when liposomes contained cardiolipin (Fig. 5A). Based on these findings, we tested
whether H1 could directly permeabilize the outer mitochondrial membranes of isolated mitochondria and cause release of
cytochrome c, which is normally retained in the mitochondrial
intermembrane space. We isolated mitochondria from L929
cells and then incubated them with different concentrations of
the H1, H3, H1:K594D, and H1:I595K peptides. To assess
permeabilization of the outer mitochondrial membrane, we
examined for leakage of cytochrome c from the intermembrane space. Although the H1 peptide localized to mitochon-
FIG. 5. H1 preferentially permeabilizes liposomes containing the mitochondrial lipid cardiolipin (A) but does not cause release of cytochrome
c from isolated mitochondria (B) or mitochondria in CHO-K1 cells microinjected with H1 (C). (A) ANTS-DPX-loaded liposomes containing a
mixture of lipids characteristic of mitochondria from Xenopus oocytes were prepared with or without 7 mol% cardiolipin. Leakage of ANTS was
quantified as before upon incubation with the indicated H1 concentrations. Means ⫾ SD of results from three independent experiments are shown.
(B) Mitochondria isolated from L929 cells were incubated with 100 ␮M the H1 or H3 peptide or with equal volumes of PBS or 1% Triton X-100
for 20 min at RT. After incubation, mitochondria were pelleted, and cytochrome c present in the pellet (P) and supernatant (S) fractions was
assayed by immunoblot analysis. A sample of purified cytochrome c (marked C) was loaded in the first lane. (C) CHO-K1 cells were microinjected
with a 1:1 mixture of Alexa594 BSA, and the H1, H1:I595K, or H3 peptide (final concentration, 50 ␮M). Cells were incubated for a further 20 min
at 37°C and then fixed and immunostained with mouse anti-cytochrome c MAb followed by Alexa488-conjugated goat anti-mouse IgG.
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FIG. 4. The peptide H1 has a bactericidal activity on Gram-positive
and Gram-negative bacteria. (A) The capacity of different concentrations of the H1 peptide to inhibit bacterial growth was assayed as
described in Materials and Methods. (B) The indicated peptides were
tested for their bactericidal activity at 50 ␮M. Results shown are the
means ⫾ SD for three independent experiments.
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REOVIRUS ␮1 AND PEPTIDE MEMBRANE PERMEABILIZATION
1513
drial membranes (Fig. 6B), we found that incubation of isolated mitochondria with 100 ␮M the H1 or H3 peptide did not
cause cytochrome c release, suggesting that any pores formed
by the H1 peptide in mitochondrial membranes do not allow
egress of cytochrome c from the mitochondrial intermembrane
space (Fig. 5B).
Microinjected H1 does not cause release of cytochrome c
from mitochondria. We have previously shown that expression
of GFP-␾ causes release of cytochrome c from the mitochondrial intermembrane space at 48 h posttransfection (31). However, we found that purified mitochondria did not release cytochrome c when they were incubated with the H1 peptide
(Fig. 5B). To test whether the H1 peptide could cause release
of cytochrome c from mitochondria in cells, we microinjected
H1 (final concentration, 50 ␮M) together with BSA fluorescently labeled with Alexa594 into CV-1 cells. After an incubation period of 20 to 30 min, the cells were fixed and immunostained for cytochrome c. We found that, similar to our
findings with purified mitochondria, the H1 peptide did not
cause release of cytochrome c from mitochondria in cells after
a 20-min incubation period (Fig. 5C). As expected, microinjection of the H1:I595K or H3 peptide also did not cause
release of cytochrome c. We conclude that although the H1
peptide can permeabilize the plasma membrane and artificial
membranes, it does not permeabilize mitochondrial membranes to an extent that will allow release of cytochrome c.
Biotinylated H1 localizes to mitochondria when microinjected into cells. A GFP-H1 fusion protein localizes to mi-
tochondria in transfected cells (3). We therefore assessed
the subcellular distribution of a biotinylated form of H1
complexed with streptavidin-Alexa488 after it was microinjected into CHO-K1 cells. Control microinjections of
streptavidin-Alexa488 alone were diffusely distributed
throughout the cell (Fig. 6A). However, the biotinylated H1
colocalized with the mitochondrial marker Mitotracker
CMXRos (Fig. 6B). In addition, 1 min following microinjection, the mitochondria in several H1-injected cells became punctate and aggregated and differed from the normal
tubulovesicular distribution of mitochondria in control injected cells (Fig. 6B, insets). These findings indicate that H1
localizes to mitochondrial membranes and disrupts normal
mitochondrial architecture.
Biotinylated H1 but not H1:I595K peptide forms a filamentous mosaic on the plasma membrane when incubated with
cells. To further investigate the mechanism of H1-induced
plasma membrane permeability, we assessed the cell surface
distribution of biotinylated H1 and H1:I595K. CV-1 monkey
kidney cells were incubated with 50 ␮M H1 for 30 min at RT
and then washed and fixed, and the distribution of biotinylated
H1 was detected by the addition of streptavidin-Alexa488. The
biotinylated H1 peptide formed a filamentous mosaic-like pattern on the plasma membrane of the CV-1 cells. In addition,
larger aggregates of peptide could be seen adherent to the
plasma membrane. In contrast, the mutant peptide H1:I595K
distributed as punctate spots on the plasma membrane and did
not form larger aggregates (Fig. 7). We conclude from these
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FIG. 6. Microinjected H1 localizes to mitochondrial membranes and disrupts normal mitochondrial architecture. CHO-K1 cells labeled with
Mitotracker CMXRos were microinjected with Alexa488-conjugated streptavidin (A) or 50 ␮M a biotinylated H1 peptide bound with Alexa488conjugated streptavidin (B). Phase-contrast and fluorescent images were collected before and 1 min after microinjection. Injected cells are marked
with red stars. Insets show an enlarged view of the region outlined in dotted lines. Bar, 10 ␮m.
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J. VIROL.
KIM ET AL.
findings that the H1 peptide can form filamentous arrays upon
interaction with cellular membranes perhaps similar to those
described for some cell-penetrating peptides upon interaction
with artificial membranes (22, 23).
DISCUSSION
The reovirus outer capsid protein ␮1 is the primary determinant of reovirus-induced apoptosis (3, 5–7). ␮1 also has an
additional critical role in membrane penetration during virus
entry. In this regard, a myristoylated N-terminal fragment of
␮1 (␮1N; residues 2 to 42) that is produced by autocatalytic
cleavage of ␮1 during virus entry has direct membrane-permeabilizing activity (21, 34). The C-terminal ␾ fragment of ␮1,
although not functionally required for entry, increases its efficiency and was suggested to have chaperone-like activity (13).
In previous work, we showed that ectopic expression of GFP-
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FIG. 7. The H1 peptide forms filaments on the plasma membrane. (A) CV-1 cells were incubated with 50 ␮M biotin-conjugated H1,
biotin-conjugated H1:I595K, or PBS. Biotinylated peptides were detected with Alexa488-conjugated streptavidin. Nuclei are stained with DAPI.
Bar, 10 ␮m. (B) Higher-magnification images of cells incubated with biotinylated H1 showing aggregates and filaments and biotinylated H1:I595K
showing punctate spots. Bar, 5 ␮M.
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REOVIRUS ␮1 AND PEPTIDE MEMBRANE PERMEABILIZATION
bilizing properties. The action of cationic amphipathic antibacterial peptides such as magainin on Gram-negative bacteria is
thought to be due to the capacity of these peptides to displace
Mg2⫹ and Ca2⫹ cations from lipopolysaccharide (LPS), causing localized destabilization of the bacterial outer membrane.
The peptides can then disrupt the cytoplasmic membrane in a
process called self-promoted uptake (11, 24).
The formation of filamentous structures on the plasma
membrane by H1 indicates that it can self-associate in the
presence of membranes to form higher-order oligomeric structures. In contrast, the mutant H1:I595K, which does not destabilize membranes, although able to associate with the
plasma membrane, did not form such structures, suggesting
that the capacity of H1 to self-associate or aggregate may be
required for membrane destabilization. The micellar aggregate
model of peptide-mediated membrane destabilization proposes that peptide aggregation within the membrane leads to
the formation of pores (12). Furthermore, collapse of such
aggregates is predicted to allow translocation of peptides into
the cytosol. In support of this mode of action for H1, we have
found that some cells incubated with biotinylated H1 have
peptide labeling on organellar membranes (data not shown).
We have proposed that when large numbers of viral particles
are added to cells, the ␾ fragment of ␮1 may gain access to the
cytosol of cells and induce apoptosis (3, 5). This hypothesis
would in part explain the capacity of UV-inactivated virions to
induced apoptosis (30) and the requirement for disassembly of
virions in order to induce apoptosis (4). Our current findings
now indicate that the minimal region of ␾ required for apoptosis induction has direct membrane-destabilizing activity and
can target mitochondrial membranes. However, we found that
neither incubation of the H1 peptide with purified mitochondria nor microinjection of H1 into cells led to release of cytochrome c from the mitochondrial intermembrane space. Taken
together, these findings support a model whereby C-terminal
fragments of ␮1 produced during virus entry and delivered into
the cytosol during membrane penetration or produced in the
cytosol as a consequence of the action of cytosolic proteases
can target and destabilize mitochondrial and other intracellular membranes. Such destabilization would be predicted to
lead to direct release of calcium and to activation of cellular
stress response pathways that would in aggregate promote
apoptosis. The recent finding that tBid is required for reovirusinduced apoptosis supports this model (8). We have previously
noted differences in the localization patterns of ␮1 and GFP-␾,
with GFP-␾ being much more strongly associated with mitochondrial membranes and ␮1 more often seen in association
with the ER and lipid droplets (3). We propose that targeting
of ␮1 to the ER leads to local release of ER calcium. Calcium
release from the ER could be responsible for activating calpain, which has been shown to be required for reovirus-induced apoptosis (9). Our current findings suggest that reovirus-induced apoptosis may initiate as a consequence of
␮1-induced intracellular membrane permeabilization.
ACKNOWLEDGMENTS
We thank Brian Ingel and Wendy Weichert for excellent technical
assistance. We thank Susan Daniel for technical advice.
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fused to residues 582 to 611 of ␮1 (within ␾) localized to
mitochondria and induced apoptosis (3). More recently, we
have found that ectopic expression of ␮1 or reovirus infection
causes release of cytochrome c and apoptosis in cells lacking
the proapoptotic BCL-2 family members Bax and Bak (31).
We now show that residues 582 to 611 of ␮1 also have direct
membrane-permeabilizing activity. These findings raise the
possibility that one mechanism by which ␮1 initiates apoptosis
in reovirus-infected cells is by directly permeabilizing cellular
membranes. The ␮1 protein localizes to mitochondria and the
ER in infected cells (3), and we speculate that direct permeabilization of these membranes with release of proapoptotic
molecules into the cytosol initiates the apoptotic process in
infected cells. Our data show that the H1 peptide directly
permeabilizes artificial membranes, causing release of small
molecules up to 10 kDa in size, but cannot itself cause release
of cytochrome c from the intermembrane space of purified
mitochondria. In contrast, ectopic expression of GFP-␾ causes
release of cytochrome c by 48 h posttransfection (31). It is
possible that the release of cytochrome c and smac/DIABLO,
which occurs during reovirus infection (14, 15), is a consequence of direct permeabilization of the outer mitochondrial
membrane by ␮1, as the full-length ␮1 protein may be able to
oligomerize and form larger pores in intracellular membranes.
However, cleaved tBid is required for reovirus-induced apoptosis (8). Therefore, it is possible that ␮1 or a fragment of ␮1
acts together with cleaved tBid to permeabilize mitochondrial
membranes in the absence of Bax and Bak. The activation of
calpain during reovirus infection (9) might be explained by
leakage of calcium from the ER as a consequence of ␮1induced membrane damage.
Our data also indicate that the H1 peptide can directly
permeabilize the plasma membranes of cells, leading to cytotoxicity that is accompanied by increases in intracytosolic calcium. Although we initially expected that the increased calcium
resulted from influx from the extracellular environment, we
found that cytosolic calcium levels significantly increased even
when extracellular calcium was absent. We further showed that
this increase in cytosolic calcium was not a consequence of
activation of phospholipase C signaling. Our findings that direct application of biotinylated H1 to cells leads to localization
of the peptide to mitochondria suggest that in following plasma
membrane permeabilization, H1 can access the cytosol and
target mitochondrial and perhaps other intracellular membranes. A recombinant mutant virus bearing the I595K mutation in ␮1 has significantly decreased capacity to activate
NF-␬B and induce apoptosis (5). The efficiency of this mutant
in inducing hemolysis of red blood cells and in entering cells
was comparable to that of wild-type virus. However, the H1
peptide bearing this mutant was not able to permeabilize cellular membranes. Our findings suggest a strong correlation
between ⌯1-induced membrane permeabilization and apoptosis induction but clearly discriminate the capacity of ␾ to permeabilize membranes from its proposed chaperone role during
virus entry.
The H1 peptide had moderate antibacterial effects against
Gram-positive and Gram-negative bacteria. Peptides derived
from HIV-1 gp41 have also been shown to kill Gram-positive
and Gram-negative bacteria (27). The mechanism of the antibacterial action of H1 is likely related to its membrane-desta-
1515
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KIM ET AL.
J. VIROL.
This research was supported by Public Health Service awards R01
AI063036 (J.S.L.P.) and a Burroughs Wellcome Fund Investigators of
Pathogenesis of Infectious Disease award (J.S.L.P.).
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