Arginine-rich cell-penetrating peptides induce membrane multilamellarity and subsequently enter via formation of a fusion pore
- aInstitute of Organic Chemistry and Biochemistry, Czech Academy of Sciences, CZ-166 10 Prague 6, Czech Republic;
- bInstitute of Physical and Theoretical Chemistry, University of Regensburg, D-93040 Regensburg, Germany;
- cFritz Haber Research Center, The Hebrew University of Jerusalem, Jerusalem 9190401, Israel;
- dDepartment of Chemistry, The Hebrew University of Jerusalem, Jerusalem 9190401, Israel;
- eFaculty of Pharmacy, University of Helsinki, Helsinki 00014, Finland;
- fJ. Heyrovský Institute of Physical Chemistry, Czech Academy of Sciences, 182 23 Prague 8, Czech Republic;
- gInstitute of Biophysics and Biophysical Chemistry, University of Regensburg, D-93040 Regensburg, Germany;
- hMicrobiology and Archaea Centre, University of Regensburg, D-93040 Regensburg, Germany;
- iInstitute of Biophysics and Biophysical Chemistry, University of Regensburg, D-93040 Regensburg, Germany;
- jImaging Methods Core Facility at Biocev, Faculty of Sciences, Charles University, 242 50 Vestec, Czech Republic
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Edited by Michael L. Klein, Temple University, Philadelphia, PA, and approved October 10, 2018 (received for review July 6, 2018)

Significance
The passive translocation mechanism of arginine-rich cell-penetrating peptides has puzzled the scientific community for more than 20 y. In this study we propose a hitherto unrecognized mechanism of passive cell entry involving fusion of multilamellar structures generated by the cell-penetrating peptides. The geometry of entry for this mechanism is completely different from previously suggested direct translocation mechanisms, leading to another paradigm for designing molecular carriers for drug delivery to the cell.
Abstract
Arginine-rich cell-penetrating peptides do not enter cells by directly passing through a lipid membrane; they instead passively enter vesicles and live cells by inducing membrane multilamellarity and fusion. The molecular picture of this penetration mode, which differs qualitatively from the previously proposed direct mechanism, is provided by molecular dynamics simulations. The kinetics of vesicle agglomeration and fusion by an iconic cell-penetrating peptide—nonaarginine—are documented via real-time fluorescence techniques, while the induction of multilamellar phases in vesicles and live cells is demonstrated by a combination of electron and fluorescence microscopies. This concert of experiments and simulations reveals that the identified passive cell penetration mechanism bears analogy to vesicle fusion induced by calcium ions, indicating that the two processes may share a common mechanistic origin.
Cell-penetrating peptides have a unique potential for targeted drug delivery; therefore, mechanistic understanding of their membrane action has been sought since their discovery over 20 y ago (1). While ATP-driven endocytosis is known to play a major role in their internalization (2), there has been also ample evidence for the importance of passive translocation (3⇓–5) for which the direct mechanism, where the peptide is thought to directly pass through the membrane via a temporary pore, has been widely advocated (4, 6⇓–8). Here, we question this view and show that arginine-rich cell-penetrating peptides instead passively enter vesicles and live cells by inducing membrane multilamellarity and fusion.
Ions do not dissolve in oil. From this point of view the direct passive mechanism of cell penetration is intuitively problematic, as cationic peptides such as polyarginines or the transactivating transcriptional activator (TAT) are too highly charged to be able to pass through the “oily” interior of a lipid membrane. The concept of direct penetration was seen plausible due to the action of the related antimicrobial peptides, which are also charged, but in addition contain a large fraction of hydrophobic residues (9): These peptides are known to stabilize pores in membranes (10). At a close inspection, however, it becomes clear that their charged side chains do not interact directly with the aliphatic chains in the low dielectric interior of the phospholipid bilayer, but rather stabilize transient water channels or act as terminal residues anchoring the transmembrane helix (9). Taken together, the passive action of cell-penetrating peptides (CPPs) seems to be very different from direct translocation across an otherwise unperturbed cell membrane.
To make matters even more confusing, experimental facts and suggested mechanisms often seem contradictory to each other. For example, there are conflicting reports whether or not nonaarginine (
Another fundamental cellular process involving membranes and charged species is fusion of vesicles with the cell membrane during calcium-triggered exocytosis. In neuronal cells, vesicle–membrane fusion is mediated by the SNARE protein complex (17, 18) with synaptotagmins (19); nevertheless, it can also be induced in in vitro lipid vesicles without the need for the presence of the protein machinery (20, 21). It is experimentally well established that Ca2+ is a key player capable of promoting vesicle fusion (22) and there is general consensus about the fusion mechanism, which proceeds via a stalk intermediate, followed by formation of a hemifused structure and opening of a fusion pore (23, 24). In this context, it is worth mentioning that cationic CPPs, especially TAT and its derivatives, are known to aggregate at phospholipid membranes and occasionally fuse vesicles (2, 5, 20, 25). This brings up the idea, which is examined further in this study, that the processes of passive cell penetration and membrane fusion may be mechanistically more intimately connected than thought so far (25).
Results and Discussion
Exploring Vesicle Penetration by a Fluorescence Leakage Assay.
To explore the potential connection between cell penetration and membrane fusion, we start by investigating the abilities of
Fluorescence spectroscopy results. (Top Left and Top Center) Threshold concentrations for leakage induced by
Membrane Fusion Induced by Calcium As Well As by Cationic Cell-Penetrating Peptides.
The range of lipid compositions of vesicles capable of being leaked by
At high peptide content, the LUV leakage kinetics are described quantitatively by a second-order rate law in the vesicle concentration (for details see SI Appendix, Fig. S11 and the kinetic model in SI Appendix). This indicates that aggregation of vesicles and the double bilayer formed during this process is essential for vesicle leakage. It is revealing that for all lipid compositions at which significant leakage occurs the vesicles also exhibit
Ideal Fusion Topologically Precludes Cell Penetration.
The similarities in aggregation/fusion caused by
The schematic mechanisms of
Induced Multilamellarity as a Solution to the Topological Conundrum Observed by Cryoelectron Microscopy: Seeing Is Believing.
A tendency of GUVs (13) or cells (12) to become multilamellar upon addition of CPPs has been observed recently. To further explore this idea, we first conducted cryoelectron microscopy (cryo-EM) experiments on LUVs. The obtained cryo-EM images indeed reveal formation of multilamellar domains and lipid bilayer bifurcations after the addition of
Electron micrographs of LUVs in the presence of
Multilamellar structures can be formed via folding of a membrane or by stacking of deflated vesicles. Any process based on direct membrane stacking would, however, add an even number of bilayers in between the vesicles and, therefore, would not lead to leakage via fusion. It is thus a key finding that by counting the lipid bilayers we frequently find odd numbers (Fig. 3D). Moreover, a close inspection of the EM micrographs provides direct evidence for bilayer bifurcation at multiple positions (see Fig. 3C for an example). We conclude that
The proposed mechanism shares some similarities with the reverse micelle mechanism, proposed in the literature (38, 39). This mechanism also necessitates a small bifurcation, before the membrane edge is closed by forming the reverse micelle. The reverse micelle has negative curvature on the inside and is, therefore, stabilized by similar interactions to those of the bifurcations. We argue that the membrane edge energy can be compensated through extension of stable cross-linked multilamellar domains as seen in the EM pictures. In SI Appendix we show simulations, which indicate the stabilization of the bifurcation by
Fluorescence and EM on HeLa Cells.
To directly explore the mechanism behind cellular uptake of CPPs in the absence of endocytosis, we first observed penetration into living human HeLa cells by fluorescence confocal microscopy (SI Appendix, Figs. S9 and S10). HeLa cells incubated with 15 μM OG-labeled
These results suggest that
EM and fluorescence microscopy images of the same spot on a fixated HeLa cell in the presence of OG-
Molecular Dynamics Simulations: Atomistic Insights.
To gain atomistic insight into the fusion process and its connection to cell penetration we performed molecular dynamics simulations. Previous studies, based on continuum and coarse-grained models, agree that fusion proceeds via a stalk intermediate (23, 35). The stalk is strongly concave, explaining the observed lipid selectivity toward small (PE) headgroups as these stabilize negative curvature.
Our simulation setups involve strongly positively curved bilayer geometries, intended to lower the barriers for fusion (42, 43). The stress hereby induced in the PE-rich bilayers leads to spontaneous stalk formation in our simulations (see Materials and Methods and SI Appendix for full details). Snapshots from the
(A) Schematic drawing of vesicle fusion–lipid cross-linking, stalk initialization, and subsequent onset of stalk formation through lipid flip-flop. (B) Time evolution of the Ca2+ fusing bilayer system. (C) Time evolution of the same system with
In Fig. 5D we examine the action of
Conclusions
In summary, we unraveled here a passive entry mechanism of CPPs via branching and layering of membranes followed by fusion of the agglomerated system. The layering is induced by a cooperative bridging of bilayers via adsorbed
Future work will be directed toward unraveling further molecular details of the cell penetration mechanism suggested in this discovery study. In the next step, we need to understand the interaction of CPPs with biological cell surfaces. Increased experimental understanding of the specific binding will allow us to develop more realistic models and vice versa. This will not only allow us to firmly establish all of the details of this hithertho unrecognized mechanism of passive cell penetration, but also have a direct impact on development of smart cell delivery strategies for therapeutic molecules using CPPs. Should we get this passive cell penetration mechanism under full control, we may eventually be able to exploit it to directly deliver cargo into the cell without the need for releasing it from the transport vesicles, as is the case in active endocytosis.
Materials and Methods
Liposome Experiments.
Leakage.
Calcein-containing vesicles were stirred at room temperature with LUV buffer in a quartz cuvette to obtain 1.5 mL of solution. The calcein fluorescence was monitored at 520 nm, with excitation at 495 nm. After an initial stirring phase of no less than 200 s, 3–6 μL peptide in buffer solution was added. After the fluorescence intensity reached a plateau, 50 μL of TRITON-X was added. Fluorescence intensity measurements were performed on a Fluorolog-3 spectrofluorimeter (model FL3–11; JobinYvon Inc.) equipped with a xenon-arc lamp. See SI Appendix for further details.
Confocal microscopy.
GUVs labeled with DiD were prepared for confocal microscopy using electroformation in a 300-mOsm/L sucrose solution. Prepared GUVs were diluted with a glucose buffer [9 mM HEPES, pH 7.40 (KOH), 90 mM KCl, 90 mM EDTA, 120 mM glucose, 300 mOsm/L, filtrated] with 20 μL of 50 nM Atto 488 to a total volume of 300 μL. Images were recorded using an Olympus IX81 laser scanning confocal microscope. For further details see SI Appendix.
Cryo-EM.
For cryo-EM sample preparation, 4 μL of the sample was applied to plasma-cleaned EM grids [400-mesh copper grids, covered with Quantifoil film (R1.2/1.3)]. Samples were plunge frozen on the grids in liquid ethane in a Grid Plunger (Leica EM GP; Leica Microsystems GmbH) with the following parameters: preblotting exposure 5 s, blotting time 1.7 s, no postblotting exposure. Chamber humidity was set to 95% at 22 °C. The LUV solution was treated with
Cell Experiments.
Forty thousand HeLa cells were seeded to a well of μ-slide (ibiTreat; ibidi) 16–20 h before the experiment. Cells were washed with SF-DMEM and kept at 4 °C for 15 min to inhibit endocytic processes. For treatment, a precooled (4 °C) 15-μM solution of a peptide in SF-DMEM was added to cells via media exchange and incubated for indicated periods of time at 4 °C. In selected cases, cells were treated for 3 min with a peptide at 4 °C, washed with precooled SF-DMEM, and further incubated for an indicated period at 4 °C in fresh SF-DMEM. Cells were imaged using a scanning confocal microscope (FluoView 1000; Olympus) and the tomograms were acquired on a Titan Halo transmission electron microscope (see SI Appendix for details).
Computational Details.
We use all atom molecular dynamics (MD) for the fusion process: In a first setup, we created two curved membranes via lipid population imbalances at the two leaflets of each bilayer. In the second setup, we put a very small vesicle composed of DOPE (80%) and DOPS (20%) in the unit cell and let it fuse with its periodic image. Both of these approaches facilitate formation of the stalk without enforcing its shape. For calcium fusion we used optimized charge-scaled force fields for ions, to account effectively for electronic polarization effects. For vesicle aggregation and bifurcation calculations we used coarse-graining methods. See SI Appendix for full details.
Acknowledgments
We thank Aleš Bendar, Markéta Dalecká, Mario Vazdar, Daniel Harries, Šárka Pokorná, Uri Raviv, and Lea Fink for discussions and technical assistance. P. Jungwirth acknowledges support from the Czech Science Foundation (Grant 16-01074S). C.A. thanks the German Academic Exchange Service for support via a Prime fellowship and the Minerva foundation for a postdoctoral fellowship. A.M. acknowledges the Magnus Ehrnrooth Foundation, Finland for funding. R.S. and M.H. acknowledge the Czech Science Foundation (Grant 17-03160S). Allocation of computer time from the Finnish IT Center for Science (CSC) is appreciated. V.H., R.R., and C.M.Z. acknowledge the use of the cryoelectron microscope in the Department of Molecular Cell Anatomy, University of Regensburg, headed by Ralph Witzgall. We acknowledge the Imaging Methods Core Facility at BIOCEV, Faculty of Sciences, Charles University, an institution supported by the Czech-BioImaging large research infrastructure project (LM2015062 and CZ.02.1.01/0.0/0.0/16_013/0001775 funded by the Czech Ministry of Education), for their support with obtaining imaging data presented in this paper.
Footnotes
↵1C.A. and A.M. contributed equally to this work.
- ↵2To whom correspondence should be addressed. Email: pavel.jungwirth{at}uochb.cas.cz.
Author contributions: C.A., A.M., and P. Jungwirth designed research; C.A., A.M., P. Jurkiewicz, K.B., M.J., P.E.M., R.S., M.C., D.H., V.H., R.R., C.M.Z., and A.S. performed research; P.E.M., M.H., V.H., R.R., and P. Jungwirth analyzed data; and C.A. and P. Jungwirth wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1811520115/-/DCSupplemental.
- Copyright © 2018 the Author(s). Published by PNAS.
This open access article is distributed under Creative Commons Attribution-NonCommercial-NoDerivatives License 4.0 (CC BY-NC-ND).
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