Translocation of ?-Galactosidase Mediated by the Cell-Penetrating Peptide Pep-1
into Lipid Vesicles and Human HeLa Cells Is Driven by Membrane Electrostatic
So ´nia Troeira Henriques,‡Ju ´lia Costa,§and Miguel A. R. B. Castanho*,‡
Centro de Quı ´mica e Bioquı ´mica, Faculdade de Cie ˆncias da UniVersidade de Lisboa, Ed. C8, Campo Grande,
1749-016 Lisboa, Portugal, and Instituto de Tecnologia Quı ´mica e Biolo ´gica, Apartado 127, 2780 Oeiras, Portugal
ReceiVed February 14, 2005; ReVised Manuscript ReceiVed May 25, 2005
ABSTRACT: The cell-penetrating peptide (CPP) pep-1 is capable of introducing large proteins into different
cell lines, maintaining their biological activity. Two possible mechanisms have been proposed to explain
the entrance of other CPPs in cells, endosomal-dependent and independent types. In this work, we evaluated
the molecular mechanisms of pep-1-mediated cellular uptake of ?-galactosidase (?-Gal) from Escherichia
coli in large unilamellar vesicles (LUV) and HeLa cells. Fluorescence spectroscopy was used to evaluate
the translocation process in model systems (LUV). Immunofluorescence microscopy was used to study
the translocation in HeLa cells. Enzymatic activity detection enabled us to monitor the internalization of
?-Gal into LUV and the functionality of the protein in the interior of HeLa cells. ?-Gal translocated into
LUV in a transmembrane potential-dependent manner. Likewise, the extent of ?-Gal incorporation was
extensively decreased in depolarized cells. Furthermore, ?-Gal uptake efficiency and kinetics were
temperature-independent, and ?-Gal did not colocalize with endosomes, lysosomes, or caveosomes.
Therefore, ?-Gal translocation was not associated with the endosomal pathway. Although an excess of
pep-1 was mandatory for ?-Gal translocation in vivo, transmembrane pores were not formed as concluded
from the trypan blue exclusion method. These results altogether indicated that protein uptake both in
vitro with LUV and in vivo with HeLa cells was mainly, if not solely, dependent on negative transmembrane
potential across the bilayer, which suggests a physical mechanism governed by electrostatic interactions
between pep-1 (positively charged) and membranes (negatively charged).
The introduction of hydrophilic molecules into mammalian
cells has become a key strategy for the investigation of
intracellular processes and drug therapy. CPPs1are very
attractive for these purposes because of their ability to
mediate cellular uptake of proteins and nucleic acids, which
is otherwise impossible because of membrane selective
permeability (1-7). These peptides are small (9-33 amino
acid residues), and their only common feature is the presence
of basic amino acid residues (2).
The most widely used CPPs are derived from HIV-1 tat
(TAT) and Drosophila Antennapedia homoprotein (1, 4, 6).
Covalent linkage of CPP with cargo molecules leads to their
nontoxic import into cells, both rapidly and efficiently, while
maintaining functional activity inside the cell (2). The
process, however, is dependent on CPP, cargo, and cell type
(8). The translocation of these cationic peptides is not well
understood. A single general mechanism for all does not
seem reasonable, and there are examples of CPPs (e.g., TAT)
that are able to use both endosomal and nonendosomal
pathways (5, 9).
amine) is an artificial CPP that has the ability to establish
hydrophobic interactions with the cargo molecule, which may
render covalent links unnecessary and favor the native
structure (10). This sequence contains a Trp-rich hydrophobic
domain, KETWWETWWTEW, a hydrophilic sequence,
KKKRKV, that is the nuclear localization sequence of simian
virus 40 large T antigen, and a spacer, SQP, linking the two
previous ones (10). This peptide has been successfully used
to translocate different biomolecules into distinct cell lines,
for instance, proteins into protoplasts (11), antibodies into
†This work was funded by Grant POCTI/BCI/38631 from FCT,
Portugal, and Grant LSHG-CT-2004-503228 from the European
Commission. We thank FCT, Portugal, for Grant SFRH/BD/14337/
2003 under the program POCTI to S.T.H.
* To whom correspondence should be addressed. Telephone:
+351217500931. Fax: +351217500088. E-mail: firstname.lastname@example.org.
‡Faculdade de Cie ˆncias da Universidade de Lisboa.
§Instituto de Tecnologia Quı ´mica e Biolo ´gica.
1Abbreviations: CPP, cell-penetrating peptide; NLS, nuclear local-
ization sequence; SV-40, simian virus 40; TAT, CPP derived from
HIV-1 tat; ?-Gal, ?-galactosidase; LUV, large unilamellar vesicles;
MUG, 4-methylumbelliferyl galactoside; 4-MU, 4-methylumbelliferone;
PMSF, phenylmethanesulfonyl fluoride; phosphine, tris(2-cyanoethyl)-
phosphine; POPC, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine;
DPPC, 1,2-dipalmitoyl-sn-glycero-3-phosphocholine; DPPS, 1,2-di-
palmitoyl-sn-glycero-3-(phospho-L-serine); MEME, Minimum Essential
Medium Eagle; NEAA, nonessential amino acids; FBS, fetal bovine
serum; BSA, bovine serum albumin; PBS, phosphate-buffered saline
solution; TRITC, tetramethylrhodamine ?-isothiocyanate; FITC, fluo-
rescein isothiocyanate; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-
tetrazolium bromide; TX-100, Triton X-100; DMSO, dimethyl sulfox-
ide; DAPI, 4′,6-diaminidino-2-phenylindole.
Biochemistry 2005, 44, 10189-10198
10.1021/bi0502644 CCC: $30.25© 2005 American Chemical Society
Published on Web 07/07/2005
porcine renal epithelial cells (LLC-PK1) (12), and peptides
into rat pheochromocytoma cells (PC-12) (13). The unspeci-
ficity of this peptide is a potential advantage for ubiquitous
It has been shown that pep-1 translocates across lipidic
vesicles only when a negative transmembrane potential exists
(14). However, it is not known if endocytosis is also involved
in pep-1 translocation in vivo as described for CPPs Tat 48-
60 and (Arg)9(5, 9). Furthermore, it is necessary to determine
if the translocation of cargo proteins via pep-1 follows a
mechanism identical to that of the free peptide.
In this work, we used ?-galactosidase (?-Gal) from
Escherichia coli as the cargo protein. ?-Gal is a homo-
tetramere with an enzymatic activity (EC 220.127.116.11) that is
easy to assess and dependent on its quaternary structure (15).
Each subunit contains 1023 amino acid residues (116 kDa),
with 38 Trp residues that enable a characterization by
fluorescence spectroscopy. The formal global charge at pH
7.4 is approximately -38 (the estimated charged is based
on the pKa’s for the isolated amino acids and was determined
using the software available at www.scripps.edu/∼cdputnam/
In this work, we have studied formation of the pep-1-?-
Gal complex and its translocation across membranes. We
have found that ?-Gal translocation into LUV and human
HeLa cells depends on the negative transmembrane potential.
Alternative pathways, such as classical and caveolin-mediated
endosomal pathways, or possible pore formation induced by
pep-1, did not account for translocation of ?-Gal into the
Reagents. Pep-1 (Ac-KETWWETWWTEWSQPKKKRKV-
cysteamine) that was >95% pure was obtained from Gen-
Script Corp. (Piscataway, NJ). ?-Gal from E. coli, 4-methyl-
umbelliferyl galactoside (MUG), porcine pancreatic lyo-
philized trypsin, phenylmethanesulfonyl fluoride (PMSF),
cholesterol (chol), Triton X-100 (TX-100), and dextran (Mr
) 10 000) were obtained from Sigma-Aldrich (St. Louis,
MO). Tris(2-cyanoethyl)phosphine (phosphine) and dextran
tetramethylrhodamine ?-isothiocyanate (TRITC) conjugate
(Mr) 10 000) were from Molecular Probes (Eugene, OR).
erol)] (POPG), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine
(DPPC), and 1,2-dipalmitoyl-sn-glycero-3-(phospho-L-serine)
(DPPS) were from Avanti Polar-Lipids (Alabaster, AL);
Minimum essential medium Eagles with Earle’s salts (MEME),
L-glutamine, nonessential amino acids (NEAA), fetal bovine
serum (FBS), streptomycin and penicillin, and trypsin-EDTA
solution (0.05% trypsin and 0.53 mM EDTA‚4Na) were
obtained from Gibco Invitrogen Corp. (Carlsbad, CA). 4′,6-
Diaminidino-2-phenylindole (DAPI) was from Sigma. The
primary antibodies used were as follows: mouse monoclonal
anti-?-Gal (AB1; 1/500 dilution) from Promega (Madison,
WI), rabbit polyclonal anti-?-Gal (AB2; 1/500) from 5Prime
(Boulder, CO), mouse monoclonal anti-EEA1 (1/100) from
BD Biosciences (Palo Alto, CA), mouse monoclonal anti-
cathepsin D (1/20) from Sigma-Aldrich (St. Louis, MO), and
rabbit polyclonal anti-caveolin-1 (1/50) from Santa Cruz
Biotechnology (Santa Cruz, CA). The secondary antibodies
were goat polyclonal anti-mouse IgG TRITC conjugate
(1/60) and goat polyclonal anti-rabbit IgG fluorescein
isothiocyanate (FITC) conjugate (1/100) obtained from
Photophysics of ?-Gal in Aqueous Solution. The experi-
ments in aqueous solution and with LUV were performed
at room temperature with a UV-vis Jasco V-530 spectro-
photometer and a SLM Aminco 8100 spectrofluorometer
(equipped with a 450 W Xe lamp and double monochroma-
tors). The solutions were prepared in 10 mM HEPES buffer
(pH 7.4) containing 150 mM NaCl, at physiologic ionic
strength. Fluorescence intensities were corrected for the inner
filter effect with the equation Ic) I × 100.5A, where Icis the
corrected intensity, I the measured intensity, and A the
absorbance at the excitation wavelength.
Protein photophysical characterization was carried out by
means of Trp fluorescence emission (λexcitation) 280 nm).
Fluorescence emission characterization and determination of
quantum yield (16) were performed. Variation of fluores-
cence emission intensity with concentration (0-269 nM) and
fluorescence quenching by acrylamide (aqueous soluble Trp
quencher) were carried out in the absence and presence of a
reducing agent, 1 mM phosphine. The quenching assay was
performed by titration of ?-Gal with acrylamide (0-60 mM)
and followed by fluorescence emission with a λexcitation of
290 nm (to minimize the relative quencher/fluorophore light
absorption ratio). The Stern-Volmer equation (I0/I ) 1 +
KSV[Q], where I and I0are the fluorescence intensity in the
presence and absence of quencher, respectively, KSVis the
Stern-Volmer constant, and [Q] is the concentration of
quencher; for a revision, see ref 17) was applied to the data.
Data were corrected for simultaneous absorption of the
fluorophore and quencher (see eq 5 in ref 18).
Enzymatic Assay of ?-Gal. Enzyme activity of ?-Gal was
?-D-galactopyranoside (MUG), a nonfluorescent substrate,
to 4-methylumbelliferone (4-MU), a fluorescent product
(λexcitation) 360 nm, λemission) 440 nm) (19). Time progres-
sion curves were performed (0-60 min); briefly, enzyme
was added to 2.5 mM MUG (substrate at nonlimiting
concentrations), in 10 mM HEPES buffer (pH 7.4), contain-
ing 150 mM NaCl, to start the reaction. The reaction was
stopped by the addition of 0.2 M NaOH-containing buffer
(to a final substrate dilution of 1/40, pH 13.2). The assay
was followed by 4-MU fluorescence intensity. The concen-
tration was determined by A360[?360) 1.9 × 104M-1cm-1
Formation of the Pep-1-?-Galactosidase Complex. The
titration of 72 nM protein with peptide was performed up to
a pep-1/?-Gal molar ratio of 100. Trp fluorescence emission
spectra were monitored to follow complex formation. The
maximum of the fluorescence emission spectrum of ?-Gal
occurs at a wavelength significantly different from that of
pep-1 (329 nm vs 346 nm). The fluorescence emission
spectra of pep-1 in the absence of ?-Gal and vice versa were
followed simultaneously, under the same conditions. Enzy-
matic activity of ?-Gal and quenching of Trp fluorescence
by acrylamide at different pep-1/?-Gal ratios was followed
as mentioned above.
Interaction of ?-Gal and the Pep-1-?-Gal Complex with
Large Unilamellar Vesicles. LUV were prepared, in 10 mM
HEPES buffer (pH 7.4) containing 150 mM NaCl, by the
10190 Biochemistry, Vol. 44, No. 30, 2005
Henriques et al.
extrusion method (21). To evaluate the interaction of 72 nM
?-Gal (free or complexed with pep-1 at different concentra-
tions) with LUV, Trp fluorescence spectral shifts were
followed (λexcitation) 280 nm) by titration of samples with
lipidic suspensions (0-3.75 mM). POPC and POPC/POPG
(4/1) bilayers in vesicles are in liquid crystalline phases;
POPC/chol (2/1) bilayers in vesicles are in the liquid-ordered
phase, and DPPC and DPPC/DPPS (4/1) bilayers are in the
Uptake of the Pep-1-?-Gal Complex in LUV with Nega-
tiVe Transmembrane Potential. The pep-1-?-Gal complex
(molar ratio of 320) was incubated (30 min) with POPC/
POPG (4/1) (final lipid concentration of 0.5 mM) LUV in
the absence or presence of negative transmembrane potential
(see ref 14 for a description of production of LUV with a
negative transbilayer potential). Briefly, valinomycin was
added, at a 1/104molar ratio (moles per mole of lipid), to
K+-loaded LUV dispersed in Na+buffer. Afterward, a trypsin
solution (final concentration of 1.3 mM) was added, and the
mixture was allowed to digest the nonincorporated pep-1 and
?-Gal, incubated for 30 min at 37 °C. After that, incubation
with 4 mM PMSF (final concentration) was carried out for
15 min to inhibit trypsin. To induce LUV permeabilization
and leakage of the incorporated ?-Gal, 0.2% (w/v) TX-100
was added. Released ?-Gal was detected by means of its
enzymatic activity, namely, by MUG hydrolysis during 20
min at 37 °C followed by fluorescence spectroscopy, as
described above. Controls without pep-1 were performed,
and the slight contribution from nonincorporated ?-Gal
resistant to trypsin hydrolysis was discounted.
Cell Culture and Cell Viability Assays. Adherent human
negroid cervix epitheloid carcinoma cells (HeLa) were grown
in MEME supplemented with 2 mM Glu, 2 mM NEAA, 10%
(v/v) FBS, and 1% (v/v) streptomycin and penicillin, in a
5% CO2humidified atmosphere at 37 °C. Cells were split
in a 1/4 dilution every 3-4 days, after they reached
confluency, which was monitored using an inverted micro-
scope (Olympus CK30). Cell viability was determined by
the colorimetric assay with 3-(4,5-dimethylthiazol-2-yl)-2,5-
diphenyltetrazolium bromide (MTT). MTT is reduced by
mitochondrial dehydrogenases of viable cells (22). Briefly,
cells were grown in 96-well plates and washed with serum-
free medium; 10 µL of MTT (5 mg/mL) was added to each
well, and a 3 h incubation at 37 °C was performed. The
purple product was solubilized in dimethyl sulfoxide (DMSO),
and the absorbance at 540 nm was determined. Alternatively,
cell viability was determined by the trypan blue exclusion
assay. Briefly, after being detached, the cell suspension was
added to a trypan blue solution (1/10 dilution) and counted
in hemacytometer. Viable cells exclude trypan blue; non-
viable cells absorb the dye and appear blue.
Cellular Uptake of Proteins and Dextran Monitored by
Immunofluorescence Microscopy. For immunofluorescence
microscopy, cells were grown on 12 mm diameter glass
coverslips in 24-well plates to approximately 70% conflu-
ence. Prior to the uptake assays, the pep-1-?-Gal complex
was formed in serum-free medium (23) during 30 min at
room temperature following the instructions of the supplier
(ActiVe Motif; Rixensart) (24). Translocation efficiency was
evaluated with different pep-1/?-Gal ratios (4, 32, 100, 200,
320, 600, and 1000) at 10.8 nM ?-Gal. Coverslips were
inverted and placed over a drop of a macromolecular
complex solution (25) and incubated for 60 min at 37 °C.
Cells were washed three times in PBS containing 0.5 mM
MgCl2, fixed in 4% (w/v) paraformaldehyde for 20 min,
permeabilized with 0.1% (w/v) TX-100 for 15 min, and then
incubated in a blocking solution that consisted of PBS
containing 1% bovine serum albumin (BSA) for 1 h (26).
After that, incubations with primary and secondary antibodies
in blocking solution, for 2 and 1 h, respectively, were
performed. Washings were carried out in PBS. Coverslips
were mounted in Airvol and observed in the Leica DMRB
fluorescence microscope and/or in the Bio-Rad MRC1024
confocal microscope. The nucleus was visualized with DAPI
that was added to the secondary antibody mixture at a
dilution of 1/1000. The kinetics of the ?-Gal translocation
process was evaluated by incubation at a pep-1/?-Gal ratio
of 320 for 10, 20, 30, 40, 50, 60, and 120 min at 4 or 37 °C.
Enzymatic activity of internalized ?-Gal by HeLa cells was
monitored by the use of MUG (see Enzymatic Assay of
?-Gal). After incubation of the complex for 60 min at 4 or
37 °C, trypsin was added to the cells to hydrolyze non-
incorporated peptide and protein. Cells were centrifuged at
500g for 5 min, washed twice in PBS, and resuspended in
0.1% (w/v) TX-100 for 15 min for permeabilization. Then
2 mM substrate was added to cells, and product progression
curves were followed for 120 min at 37 °C. NaOH was
added, and product formation was monitored as described
above. A control without pep-1 was carried out.
The uptake of the anti-mouse IgG-TRITC conjugate
mediated by pep-1 was monitored by immunofluorescence
microscopy under live conditions, without the paraformal-
dehyde fixation step, in a 60 min incubation at 4 or 37 °C,
at a 1/320 protein/pep-1 ratio.
Cells were also incubated with the endocytic tracer
dextran-TRITC at 2 mg/mL associated with pep-1, at 37
and 4 °C for 60 min, in the presence or absence of 20 mg/
mL nonlabeled dextran.
Confocal Microscopy and Colocalization Analysis. For
each picture, laser intensities and amplifier gains were
adjusted to prevent pixel saturation. This was done using
GLOW LUT in the Leica Confocal software. Each fluoro-
phore that was used was excited and detected separately to
avoid any signal crossover. Each picture consisted of a
z-series of 14 images of 1024 × 1024 pixel resolution with
a pinhole Airy unit. Colocalization analysis was per-
formed using open source Image J version 1.30 (http://
rsb.info.nih.gov/ij/). The procedure was applied for a popula-
tion of 6-10 cells. Quantification of colocalization of ?-Gal
(AB1 or AB2) with endosomes (anti-EEA1), lysosomes (anti-
cathepsin D), and caveosomes (anti-caveolin-1) was based
on that previously described (27).
Effect of Transmembrane Potential on the Uptake of the
Pep-1/?-Gal Complex by HeLa Cells. To decrease the
transmembrane potential, cells were incubated, for 30 min
at 37 °C, with the pep-1-?-Gal complex preformed in PBS
buffer, containing different K+concentrations, with Na+
replaced by K+, at increasing concentrations (28). Quanti-
fication of internalized ?-Gal was carried out by using
enzymatic activity. Cell viability in the presence of different
K+concentrations was determined by the trypan blue
exclusion method. Absolute fluorescence intensity data were
divided by the number of viable cells.
Uptake Mechanism of the Pep-1-?-Galactosidase Complex
Biochemistry, Vol. 44, No. 30, 2005 10191
Formation of the Pep-1-?-Gal Complex in Aqueous
Solution. To investigate the suitability of ?-Gal for cellular
uptake studies mediated by pep-1, characterization of the
protein and the complex formed in aqueous solution has been
performed. Each subunit of ?-Gal contains 38 Trp, 35 Phe,
and 25 Tyr residues. Fluorescence emission with a λexcitation
of 280 nm is largely dominated by the Trp residues. Most
hydrophobic residues are not accessible to the aqueous
environment as concluded from the ?-Gal crystallographic
structure. In aqueous solution, fluorescence emission has a
quantum yield (Φ) of 0.099 ( 0.005 (constant up to 269
nM) with a band maximum at 329 nm which was signifi-
cantly different from the maximum of free Trp. This blue-
shifted emission is in agreement with most of the Trp
residues being inaccessible to the aqueous environment. The
low accessibility of Trp residues to the aqueous environment
was confirmed by acrylamide quenching; KSVwas signifi-
cantly lower from that obtained for free Trp (5.6 ( 0.3 and
18.9 ( 0.3 M-1, respectively). The initial velocity of MUG
hydrolysis, catalyzed by ?-Gal, was 17.5 µM min-1, and
linearity was maintained for 30 min.
Since the cytoplasm is a reducing environment, we have
tested the effect of the reducing agent phosphine (1 mM) on
?-Gal conformation and activity. The quantum yield was
slightly decreased (Φ ) 0.067 ( 0.007), but the accessibility
of Trp to aqueous solution (KSVby acrylamide is 5.4 ( 0.2
M-1) was not altered. A concomitant slight decrease in the
initial velocity of enzymatic activity occurred (12.0 µM
The initial velocity of the enzymatic reaction catalyzed
by ?-Gal was maintained up to a peptide/protein molar
ratio of 16. In this range, the peptide strongly interacts
with the protein (see the Supporting Information for experi-
mental results and discussion). The initial velocity de-
creased to 78% (V0) 13.4 µM min-1) and 54% (V0) 9.4
µM min-1) of the starting value at pep-1/?-Gal ratios of 40
and 400, respectively. In a reducing environment, a more
pronounced decrease was observed: for the 400 complex,
V0) 5.8 µM min-1, which corresponded to a reduction of
52% of the enzymatic activity in the absence of pep-1 (12.0
Transmembrane Potential Is Required for Translocation
of the Pep-1-?-Gal Complex into LUV. No spectral alter-
ations occurred when ?-Gal was in the presence of neutral
or negatively charged LUV [POPC, POPC/POPG (4/1),
POPC/chol (2/1), DPPC, and DPPC/DPPS (4/1)] (Table 1).
A slight blue shift in emission spectra was detected for pep-
1-?-Gal complexes (molar ratios of 4, 16, 38, 60, and 400)
in POPC, POPC/POPG (4/1), and POPC/chol (2/1) lipidic
systems (Table 1).
We investigated the uptake of ?-Gal mediated by pep-1
into LUV after the induction of an electrostatic gradient
across the membrane by valinomycin. Briefly, after a 30 min
incubation with the complex, hydrolysis of the pep-1-?-
Gal complex outside LUV was achieved with trypsin; then
the activity of ?-Gal enclosed in the LUV lumen was
determined after release with TX-100. Product formation
concentration was enhanced by a factor of 5 in the presence
of the negative transmembrane potential (when compared
with the situation without a potential). The fraction of
translocated protein is low, which is expected considering
the total volume of the vesicles in the total bulk solution
Therefore, these results indicate not only that pep-1
improves the affinity of the protein for the membrane but
also that the presence of a transmembrane potential induces
its translocation across a bilayer.
Pep-1 Does Not Induce Pore Formation in HeLa Cells. It
has been suggested that protein uptake mediated by pep-1
involves pore formation (29), but this is controversial in light
of biophysical studies with lipidic vesicles (14). To test this
hypothesis, we have determined the cell viability by the
trypan blue exclusion method. This method relies on the
inclusion of the dye trypan blue by dead cells once their
plasma membrane is permeable or damaged. If pep-1 would
induce pores on the plasma membrane of cells, a larger
amount of intracellular trypan blue would be observed.
However, this was not the case since viabilities were similar
for control cells (96.4 ( 0.6%) or cells incubated with pep-1
and the pep-1-?-Gal complex (95.5 ( 1.6 and 95.6 ( 0.1%,
respectively). These results indicated that pep-1 did not
induce permeability changes and hence pore formation in
the plasma membrane of HeLa cells.
To evaluate the possible toxic effect of pep-1 on the cells,
we have used an independent cell viability test, the MTT
assay. MTT is a very sensitive way of determining cytotox-
icity. In living cells, mitochondrial dehydrogenase enzymes
oxidize the yellow MTT and convert it into purple formazan
crystals (22). It was observed that the peptide induced a
reduction in cell viability of approximately 22%. Since
viability calculated by the trypan blue assay was not
decreased after addition of pep-1, these results suggested that
pep-1 might inhibit mitochondrial dehydrogenases.
Electrostatic Transmembrane Potential Is a Sine-Quanon
Requirement for Translocation of the Pep-1-?-Gal Complex
into HeLa cells. To monitor the uptake of the pep-1-?-Gal
complex by HeLa cells, increasing pep-1 concentrations and
Table 1: Partition of the Pep-1-?-Gal Complex into LUVa
aMaximum fluorescence emission of the pep-1-?-Gal complex, at
different ratios, in aqueous solution [10 mM HEPES buffer containing
150 mM NaCl (pH 7.4)] and in the presence of a 3.75 mM lipidic
suspension.bThe shift is the difference between the two conditions.
10192 Biochemistry, Vol. 44, No. 30, 2005
Henriques et al.
different incubation times have been tested. After a 60 min
incubation at a pep-1/?-Gal ratio of 320, a significant uptake
of ?-Gal has been observed (Figure 1A). ?-Gal was found
dispersed in aggregates within the cytosol. The presence of
protein inside the cell and not adsorbed on the cell surface
was confirmed by confocal microscopy. Apparently, the
translocation efficiency did not increase for complex ratios
between 320 and 1000. For lower ratios, between 4 and 200,
uptake of ?-Gal did not occur. Therefore, the chosen pep-
1/?-Gal ratio for all the experiments was 320. At this ratio,
there was an excess of soluble pep-1 in solution (see Figure
S1 of the Supporting Information).
Transfection was relatively fast; after 10 min, it was
already possible to identify a small quantity of protein in
some cells. The level of transfection increased until ap-
proximately 40 min; after this time, it seemed to stabilize
(data not shown). Translocation occurred with a similar
efficiency at 4 °C (Figure 1A). To test if ?-Gal was active
after translocation, MUG hydrolysis was monitored in cells
incubated at 37 and 4 °C with the complex. It was observed
that ?-Gal was indeed active after translocation at both
temperatures (Figure 1B).
For a typical animal cell characterized by dominant
potassium permeability, increasing the external potassium
concentration necessarily reduces the transmembrane poten-
tial due to the decrease in the electrochemical gradient of
K+across the cell membrane (28, 30). Negative transmem-
brane potential is dominated by potassium potential equi-
librium, which can be estimated by the potassium Nernst
potential (EK): EK) (RT/F) ln(K0/Ki), where R is the gas
constant, T is the absolute temperature, F is Faraday’s
constant, and K0and Kiare the extracellular and intracellular
potassium concentrations, respectively. So, increasing the
extracellular K+concentration leads to less negative Nernst
potentials (Figure 2).
To test if the electrostatic membrane potential would be
required for translocation of the pep-1-?-Gal complex, HeLa
cells have been incubated with the complex at increasing
external K+concentrations. The total ionic strength ([K+]
+ [Na+] ) 150 mM) was kept constant. The internalized
?-Gal concentration was estimated from enzymatic MUG
hydrolysis. This assay was performed for 30 min to guarantee
that the initial velocity of the reaction was maintained, and
that the concentration of ?-Gal uptake was directly related
FIGURE 1: Translocation of ?-Gal, mediated by pep-1, into HeLa cells. Panel A shows immunofluorescence microscopy detection of ?-Gal.
The pep-1-?-Gal complex (molar ratio of 320) was incubated with HeLa cells, at 37 or 4 °C, for 60 min. Cells were fixed with 4%
paraformaldehyde and permeabilized with 0.1% (w/v) TX-100. ?-Gal was detected with rabbit polyclonal anti-?-Gal and the secondary
anti-rabbit antibody coupled to FITC. DAPI was used to identify the nucleus. Internalized ?-Gal is in the cytosol. The scale bar is 10 µm.
Panel B shows enzymatic activity of ?-Gal internalized in a HeLa cell suspension. ?-Gal uptake, after incubation with the pep-1-?-Gal
complex (molar ratio of 320) for 60 min at 4 (9) or 37 °C (0), was followed by the progression curve of enzymatic MUG hydrolysis, at
37 °C. The control without pep-1 was subtracted. The formation of 4-MU monitored by fluorescence intensity at 440 nm with excitation
at 360 nm indicates that ?-Gal was efficiently translocated, at both temperatures, in an active form.
Uptake Mechanism of the Pep-1-?-Galactosidase Complex
Biochemistry, Vol. 44, No. 30, 2005 10193
with 4-MU production. A decrease in the absolute value of
the electrochemical K+gradient (calculated considering Ki
) 140 mM) led to a severe drop in the level of ?-Gal uptake
(Figure 2). When K0 ) 114.6 mM, uptake was almost
Although for low extracellular K+concentrations the
contribution of other conductors makes the membrane
potential less negative than that predicted by K+Nernst
potential (28), the dependence of ?-Gal uptake on trans-
membrane K+Nernst potential was clear.
The Pep-1-?-Gal Complex Is Not Internalized by HeLa
Cells Via the Endosomal Pathways. Other CPPs such as the
one derived from TAT and (Arg)9(5) seem to translocate
proteins across cells by two different mechanisms: one is
fast and physical in nature and the other is mediated by
endocytosis. In the case of an endosomal-dependent pathway
of ?-Gal uptake mediated by pep-1, colocalization with
endosomes or lysosomes at some extension would be
expected. To evaluate this possibility, colocalization of pep-
1-translocated ?-Gal with EEA1 (early endosomal marker),
caveolin-1 (caveosomes marker), and cathepsin D (lysosomal
marker) was performed. Monitoring localization of the
protein, instead of a fluorescence-labeled pep-1, is a better
choice. It prevents problems associated with the apparent
uptake of cationic peptides bound to negatively charged
membranes, causing an artifactual localization in cells (5)
and the possible influence of the fluorescent label in the
uptake and intracellular localization of the peptide (31). The
percentages of colocalization with each of the organelles from
the endocytic pathway determined after immunofluorescence
confocal microscopy were very low (shown on the right in
Figure 3), after incubation for 60 or 120 min. Similar results
were obtained at 4 °C. These results indicate that the uptake
of ?-Gal did not involve the endocytic pathway.
It has been suggested that fixation conditions could lead
to an artifactual uptake of cationic peptides associated with
the cell membrane at 4 °C (5, 9, 32). The uptake of a protein
FIGURE 2: Variation of uptake of ?-Gal into a HeLa cell suspension,
mediated by pep-1, with potassium Nernst potential. The pep-1-
?-Gal complex (molar ratio of 320) was incubated with HeLa cells,
for 30 min at 37 °C, in the presence of increasing external K+
concentrations, and a constant ionic strength ([K+] + [Na+] ) 150
mM). The relative level of ?-Gal uptake was determined from ?-Gal
enzymatic hydrolysis of MUG for 20 min at 37 °C. The potassium
Nernst potential was determined considering an internal K+
concentration of 140 mM; the external K+concentration ranged
from 5 to 114.6 mM, which corresponded to a range from -89 to
-5.4 mV, respectively (see the equation in the text). An increasing
external K+concentration severely reduces the level of ?-Gal
FIGURE 3: Immunofluorescence microscopy localization of pep-1-translocated ?-Gal, EEA1, caveolin-1, and cathepsin D from HeLa cells.
Cells were incubated with the pep-1-?-Gal complex (molar ratio of 320) for 60 min. Cells were fixed with 4% paraformaldehyde and
permeabilized with 0.1% (w/v) TX-100. ?-Gal, detected with rabbit polyclonal or mouse monoclonal antibodies, was colocalized with early
endosomal EEA1, caviosomal caveolin-1, and lysosomal cathepsin D. Secondary antibodies were the anti-rabbit antibody coupled to FITC
and the anti-mouse antibody coupled to TRITC. Pictures of a z-series of 14 images from the confocal microscope were analyzed with
Image J version 1.3 to perform colocalization analysis. Six to 10 cells were observed. The scale bar is 10 µm.
10194 Biochemistry, Vol. 44, No. 30, 2005
Henriques et al.
covalently marked with a fluorescent probe can be visualized
by fluorescence microscopy without the need to fixate or
permeabilize the cells. The uptake of anti-mouse TRITC at
4 and 37 °C was performed and visualized by immuno-
fluorescence microscopy without fixation (Figure 4). It was
possible to identify the presence of protein inside the cell at
FIGURE 4: Uptake into HeLa cells of anti-mouse antibody conjugated with TRITC, mediated by pep-1. Incubation of cells with a pep-1-
anti-mouse-TRITC complex was performed for 60 min at 4 or 37 °C. Cells were visualized under live conditions in a fluorescence microscope.
The scale bar is 10 µm.
FIGURE 5: Immunofluorescence microscopy localization of translocated ?-Gal and the dextran-TRITC marker in HeLa cells (incubation
for 60 min at 37 or 4 °C). Panel A shows incubation with the pep-1-?-Gal complex (molar ratio of 320) where the endocytic dextran-
TRITC marker (2 mg/mL) was added to the complex at the time of cell incubation. Panel B shows incubation with the preformed pep-
1-?-Gal-dextran-TRITC complex. Cells were fixed with 4% paraformaldehyde and permeabilized with 0.1% (w/v) TX-100. ?-Gal was
detected with rabbit polyclonal anti-?-Gal and anti-rabbit antibodies coupled to FITC. Pictures of a z-series of 14 images from the confocal
microscope were analyzed with Image J version 1.3 and were used to perform colocalization analysis. Six to 10 cells were used. The scale
bar is 10 µm.
Uptake Mechanism of the Pep-1-?-Galactosidase Complex
Biochemistry, Vol. 44, No. 30, 2005 10195
both temperatures. This indicated that internalized protein
was not an artifact associated with fixation.
The Polysaccharide Dextran Is Translocated into HeLa
Cells by Pep-1. Uptake of the dextran-TRITC complex by
HeLa cells occurs by a classical endosomal pathway. At
37 °C, ?-Gal did not colocalize with the dextran-TRITC
marker (1.0%) (Figure 5A), supporting the idea that ?-Gal
would be translocated into the cells by a mechanism
independent of the endocytic pathway. However, at 4 °C,
there was an extensive colocalization between ?-Gal and the
dextran-TRITC marker (51.3%) (Figure 5A). At 4 °C, the
classical endosomal pathway is inhibited, so the endocytic
dextran-TRITC marker did not enter the cells via this
pathway (see control in Figure 6, left image). The internal-
ization of the dextran-TRITC marker at 4 °C and its
colocalization with ?-Gal suggested that the uptake of
translocation of the polysaccharide was mediated by pep-1.
However, if the dextran-TRITC marker was preincubated
together with pep-1 and ?-Gal, a ternary complex seems to
be formed and ?-Gal-dextran-TRITC colocalization oc-
curred, even at 37 °C (Figure 5B).
To confirm that pep-1 was translocating the dextran-
TRITC marker via the polysaccharide without inter-
ference of artifacts from the fluorophore moiety, the HeLa
cells were incubated with pep-1-dextran-TRITC complex,
at 4 °C, in the presence of nonlabeled dextran (Figure 6). It
was observed that the presence of nonlabeled dextran
dramatically reduced the amount of dextran-TRITC marker
translocated into the cells, showing competition between
nonlabeled dextran and the dextran-TRITC marker. This
confirmed that pep-1 mediated the transport of the polysac-
charide into HeLa cells without any artifactual interference
?-Gal from E. coli was chosen for the study of the
interaction of pep-1 with a protein and evaluation of the
translocation efficiency inside human tumoral cells. Its
enzymatic activity is easy to access. Trp residues enable
fluorescence spectroscopy techniques to be used as analytical
tools. The absence of a signal sequence makes the protein
unable to address any specific organelle inside the cell;
therefore, this molecule is very useful in evaluating if the
pep-1-mediated translocation leads protein to a specific
organelle in human cells. HeLa cells are derived from a
human cervix epitheloid carcinoma; they are relatively large
(approximately 20 µm) and particularly suitable for organelle
visualization by immunofluorescence studies. Therefore, they
were chosen as the cellular model system.
?-Gal characterization in aqueous solution and in the
presence of LUV revealed that Trp residues are protected
from interaction with aqueous solution and no significant
interaction with membranes was observed. Enzymatic assays
revealed that ?-Gal maintains its quaternary structure, which
is required for enzymatic activity (15) in aqueous solution,
under reducing and nonreducing conditions. These results
showed that the reducing environment did not modify the
?-Gal conformation, so maintenance of enzyme activity
inside the cell was expected.
Fluorescence quenching upon the titration of the protein
with pep-1 suggests the existence of an excess of pep-1 in
solution not interacting with the protein above peptide/protein
molar ratios of 60 (data and detailed discussion in the
Supporting Information). Enzymatic activity is still detected
in the pep-1-?-Gal complex (a molar ratio of up to 400),
which eliminates the severe perturbation of ?-Gal structure
due to pep-1.
FIGURE 6: Uptake of the dextran-TRITC marker, mediated by pep-1, into HeLa cells. Comparison of dextran-TRITC (2 mg/mL) uptake,
mediated by pep-1, at 4 °C for 60 min when the dextran-TRITC-pep-1 complex was preformed in the presence or absence of dextran (20
mg/mL). Cells were fixed with 4% paraformaldehyde and permeabilized with 0.1% (w/v) TX-100. The dextran-TRITC marker at 4 °C
was detected only when pep-1 was present. In the presence of dextran (nonconjugated), the rate of uptake of the dextran-TRITC marker
is decreased, indicating a competition of pep-1 for nonconjugated dextran. The scale bar is 10 µm.
10196 Biochemistry, Vol. 44, No. 30, 2005
Henriques et al.
The wavelength of maximal fluorescence emission inten-
sity of pep-1 in lipidic membranes systems is higher than
that observed for the complex (see Table 1). The blue shift
observed for pep-1-?-Gal complexes relative to that of pep-1
alone together with its dependence on the peptide/protein
molar ratio showed that pep-1 is mediating the partitioning
of ?-Gal into lipidic membranes.
Pep-1 translocation in LUV was previously found to be
dependent on the transmembrane potential across bilayers
(14). In fact, in this work, the same was concluded for the
translocation of ?-Gal mediated by pep-1 in LUV. These
results suggested a common translocation mechanism for free
and complexed pep-1. This property has been shown
previously for other free peptides (28, 33) but is here
presented for the first time for a CPP-cargo protein complex,
to the best of our knowledge.
?-Gal from E. coli was efficiently transported by pep-1
into HeLa cells, maintaining its enzymatic activity. The
translocation of protein is dependent on peptide concentra-
tion; at a pep-1/?-Gal molar ratio of up to 200, translocation
was not detected. With a pep-1/?-Gal ratio of 320, the
process is very efficient. At this ratio, there is free pep-1 in
solution (see Figure S1 of the Supporting Information), which
seems to play a role in the translocation process. An excess
of pep-1 is probably necessary for membrane destabilization
(14), facilitating the uptake of the protein. If the translocation
was mediated by pore formation, a very large pore (≈104
Å × 140 Å, estimated from the ?-Gal crystallographic
structure) would be necessary for a protein as large as ?-Gal
(116 kDa) to pass across the bilayer. A pore with such a
diameter would probably induce leakage of the cellular
contents at some extension, compromising cellular viability.
Cell viability was maintained in the presence of pep-1 and
the complex, and pore formation was not detected. This is
in agreement with the study of interaction of pep-1 with LUV
(see ref 14), where pep-1 translocation occurred without pore
The high velocity of the translocation mechanism (10 min
was enough to detect protein inside the cell) and the
independence of temperature (see Figure 1A,B) suggested a
physical mechanism not dependent on complex cellular
biochemical processes. This hypothesis was confirmed by
the correlation found between cell depolarization and ?-Gal
translocation. Destroying the potassium electrochemical
gradient drastically reduced the level of ?-Gal uptake in HeLa
cells (see Figure 2). These results are in agreement with those
obtained with LUV. Therefore, in vivo the membrane charge
asymmetry (34, 35) and the combined effect of membrane
potentials (36) seemed to be the driving forces responsible
for the translocation process.
Inside the cell, the protein was found in the cytosol and
did not colocalize with endosomes, lysosomes, or caveo-
somes (see Figure 3). These results were consistent with a
translocation process independent of the classical endosomal
pathways or caveolin-mediated endosomal pathways. In the
case of endocytosis being a secondary mechanism for uptake,
at least a small colocalization with one of these organelles
would be expected. Monitoring localization of the protein,
instead of pep-1, prevents problems associated with the
apparent uptake of cationic peptides bound to negatively
charged membranes, causing an artifactual localization in
cells (5). Lebleu and co-workers found that TAT and
oligoarginine uptake was dependent on endocytosis (5), but
Norde ´n and co-workers (9) have proven that for arginine-
rich peptides both nonendocytic and endocytic uptake
pathways were involved in their cellular internalization. This
nonendocytic mechanism was fast and biologically relevant
(9). An endocytic pathway was not detected for pep-1, but
a physical transmembrane crossing mechanism was.
A translocation mechanism independent of endocytosis
was further confirmed under nonfixation conditions with anti-
mouse TRITC where protein uptake at 4 °C has been
observed (see Figure 4).
The transport of the dextran-TRITC marker mediated by
pep-1 under conditions where the endocytosis was inhibited
was also demonstrated (see Figures 5 and 6). It has been
suggested by Morris et al. (10) that pep-1 interacts with
macromolecules via hydrophobic interactions. Given the
hydrophilic nature of the molecule, the capacity of pep-1 to
translocate dextran demonstrated that hydrophobic pockets
are not essential for complex formation and uptake. Complex
formation of pep-1 and dextran is probably due to polar
interactions and hydrogen bonding.
In conclusion, the interaction of pep-1 with ?-Gal from
E. coli was extensively studied to gain insight into the
translocation mechanism at the molecular level. It has been
demonstrated that pep-1 can establish a variety of electro-
static and/or hydrophobic and/or hydrophilic interactions with
the cargo. The existence of a negative transmembrane
potential promotes uptake of ?-Gal, mediated by pep-1, in
vitro and in vivo, and the absence of a transmembrane
potential inhibits it. The charge asymmetry (negative inside)
seems to be the driving force for translocation to occur. There
was no evidence found for the involvement of the endocytic
pathway in the uptake of cargo mediated by pep-1. Further-
more, pep-1 did not induce the formation of pores in the
membrane. These results together suggested that the peptide
and the cargo translocate only by a physical process.
We thank Dr. Adriano Henriques (Instituto de Tecnologia
Quı ´mica e Biolo ´gica) for the anti-?-galactosidase antibodies.
We thank the Cell Imaging Service (IGC, Oeiras, Portugal)
for the use of the confocal microscope.
SUPPORTING INFORMATION AVAILABLE
Further experimental results and discussion. This material
is available free of charge via the Internet at http://
1. Wadia, J. S., Becker-Hapak, M., and Dowdy, S. F. (2002)
Interactions of cell-penetrating peptides with membranes, in Cell-
penetrating peptides, processes and applications (Langel, U ¨., Ed.)
pp 365-375, CRC Press Pharmacology and Toxicology Series,
CRC Press, New York.
2. Bogoyevitch, M. A., Kendrick, T. S., Dominic, C. H., and Barr,
R. K. (2002) Taking the cell by stealth or storm? Protein
transduction domain (PTDs) as versatile vectors for delivey, DNA
Cell Biol. 21, 879-894.
3. Bonetta, P. (2002) Getting protein into cells, Scientist 17, 38-40.
4. Eguchi, A., Akuta, T., Okuyama, H., Senda, T., Yokoi, H.,
Inokuchi, H., Fujita, S., Hayakama, T., Takeda, K., Hasegawa,
M., and Nakanisshi, M. (2001) Protein transduction domain of
HIV-1 Tat protein promotes efficient delivery of DNA into
mammalian cells, J. Biol. Chem. 247, 27205-27210.
Uptake Mechanism of the Pep-1-?-Galactosidase Complex
Biochemistry, Vol. 44, No. 30, 2005 10197
5. Richard, J. P., Melikov, K., Vives, E., Ramos, C., Verbeure, B., Download full-text
Gait, M. J., Chernomordik, L. V., and Lebleu, B. (2003) Cell-
penetrating peptides, a reevaluation of the mechanism of cellular
uptake, J. Biol. Chem. 278, 585-590.
6. Schwarze, S. R., Hruska, K. A., and Dowdy, S. F. (2000) Protein
transduction: Unrestricted delivery into all cells? Trends Cell Biol.
7. Chaloin, L., Mau, N. V., Divita, G., and Heitz, F. (2002)
Interactions of cell-penetrating peptides with membranes, in Cell-
penetrating peptides, processes and applications (Langel, U ¨., Ed.)
pp 23-51, CRC Press Pharmacology and Toxicology Series, CRC
Press, New York.
8. Joliot, A., and Prochiantz, A. (2004) Transduction peptides: From
technology to physiology, Nat. Cell Biol. 6, 189-196.
9. Thore ´n, P. E. G., Persson, D., Isakson, P., Gokso ¨r, M., O ¨nfelt,
A., and Norde ´n, B. (2003) Uptake of analogs of penetratin,
Tat(48-60) and oligoarginine in live cells, Biochem. Biophys. Res.
Commun. 307, 100-107.
10. Morris, M. C., Depollier, J., Mery, J., Heitz, F., and Divita, G.
(2001) A peptide carrier for the delivery of biologically active
proteins into mammalian cells, Nat. Biotechnol. 19, 1143-1147.
11. Wu, Y., Wood, M. D., and Katagiri, F. (2003) Direct delivery of
bacterial avirulance proteins intro resistant Arabidopsis protoplasts
lead to hypersensitive cell death, Plant J. 33, 131-137.
12. Ikari, A., Nakaro, M., Kawano, K., and Suketa, Y. (2002) Up-
regulation of sodium-dependent glucose transporter by interaction
with heat shock protein 70, J. Biol. Chem. 277, 33338-33343.
13. Zhou, J., and Hsieh, J.-T. (2001) The inhibitory role of DOC-2/
DAB2 in growth factor receptor-mediated signal cascade, J. Biol.
Chem. 276, 27793-27798.
14. Henriques, S. T., and Castanho, M. A. R. B. (2004) Consequences
of nonlytic membrane perturbation to the translocation of the cell
penetrating peptide pep-1 in lipidic vesicles, Biochemistry 43,
15. Nichtl, A., Buchner, J., Jaenicke, R., Rudolph, R., and Scheibel,
T. (1998) Folding and association of ?-galactosidase, J. Mol. Biol.
16. Fery-Forgues, S., and Lavabre, D. (1999) Are fluorescence
quantum yields so tricky to measure? A demonstration using
familiar stationery products, J. Chem. Educ. 76, 1260-1264.
17. Lakowics, J. R. (1999) Principles of fluorescence spectroscopy,
2nd ed., Kluwer Academic/Plenum, New York.
18. Coutinho, A., and Prieto, M. (1993) Ribonuclease T1 and alcohol
dehydrogenase fluorescence quenching by acrylamide. A labora-
tory experiment for undergraduate students, J. Chem. Educ. 70,
19. McGuire, J. B. J., James, T. J., Imber, C. J., St. Peter, S. D., Friend,
P. J., and Taylor, R. P. (2002) Optimisation of an enzymatic
method for ?-galactosidase, Clin. Chim. Acta 326, 123-129.
20. Haugland, R. P. (2002) Handbook of fluorescent probes and
research products, 9th ed., Molecular Probes, Eugene, OR.
21. Mayer, L. D., Hope, M. J., and Cullis, P. R. (1986)Vesicles of
variable sizes produced by a rapid extrusion method, Biochim.
Biophys. Acta 858, 161-168.
22. Mosmann, T. (1983) Rapid colorimetric assay for cellular growth
and survival: Application to proliferation and citotoxicity assays,
J. Immunol. Methods 65, 55-63.
23. Coulpier, M., Anders, J., and Iba ´n ˜ez, C. F. (2002) Coordinated
activation of autophosphorylation sites in the RET receptor
tyrosine kinase. Importance of tyrosine 1062 for GDNF mediated
neuronal differentiation and survival, J. Biol. Chem. 277, 1991-
24. Chariot, simple efficient protein delivery system. http://
www.activemotif.com/products/cell/chariot.php (accessed May
25. Tisdale, E. J. (2002) Glyceraldehyde-3-phosphate dehydrogenase
is phosphorylated by pyotein kinase C and plays a role in
microtubule dynamics in the early secretory pathway, J. Biol.
Chem. 277, 3334-3341.
26. Sousa, V. L., Brito, C., Costa, T., Lanoix, J., Nilsson, T., and
Costa, J. (2003) Importance of Cys, Gln, and Tyr from the
transmembrane domain of Human ?3/4 fucosyltransferase III for
its localization and sorting in the Golgi of baby hamster kidney
cells, J. Biol. Chem. 278, 7624-7629.
27. Sousa, V. L., Brito, C., and Costa, J. (2004) Delection of the
cytoplasmic domain of human R3/4 fucosyltransferase III causes
the shift of the enzyme to early golgi compartments, Biochim.
Biophys. Acta 1675, 95-104.
28. Rothbard, J. B., Jessop, T. C., Lewis, R. S., Murray, B. A., and
Wender, P. A. (2004) Role of membrane potential and hydrogen
bonding in the mechanism of translocation of guanidium-rich
peptides into cells, J. Am. Chem. Soc. 126, 9506-9507.
29. Deshayes, S., Heitz, A., Morris, M. C., Charnet, P., Divita, G.,
and Heitz, F. (2004) Insight into the mechanism of internalization
of the cell-penetrating carrier peptide pep-1 through conforma-
tional analysis, Biochemistry 43, 1449-1457.
30. Chifflet, S., Herna ´ndez, J. A., Grasso, S., and Cirillo, A. (2003)
Nonspecific depolarization of the plasma membrane potential
induces cytoskeletal modifications of bovine corneal endothelial
cells in culture, Exp. Cell Res. 282, 1-13.
31. Szeto, H. H., Schiller, P. W., Zhao, K., and Luo, G. (2005)
Fluorescent dyes alter intracellular targeting and function of cell-
penetrating tetrapeptides, FASEB J. (in press).
32. Lundberg, M., and Johansson, M. (2002) Positively charged DNA-
binding proteins cause apparent cell membrane translocation,
Biochem. Biophys. Res. Commun. 291, 367-371.
33. Terrone, D., Sang, S. L. W., Roudaia, L., and Silvius, J. R. (2003)
Penetratin and related cell-penetrating cationic peptides can
translocate across lipid bilayers in the presence of a transbilayer
potential, Biochemistry 42, 13787-13799.
34. Gennis, R. B. (1989) Molecular structure and functions, Springer-
Verlag, New York.
35. Manno, S., Takakuwa, Y., and Mohandas, N. (2002) Identification
of a functional role for lipid asymmetry in biological mem-
branes: Phosphatidylserine-skeletal protein interactions modulate
membrane stability, Proc. Natl. Acad. Sci. U.S.A. 99, 1943-1948.
36. O’Shea, P. (2003) Intermolecular interactions with/within cell
membranes and the trinity of membrane potentials: Kinetics and
imaging, Biochem. Soc. Trans. 31, 990-996.
10198 Biochemistry, Vol. 44, No. 30, 2005
Henriques et al.