Fusogenic Liposomes as Nanocarriers for the Delivery of Intracellular

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Fusogenic Liposomes as Nano Carriers for Delivery of Intracellular Proteins Sarah Kube, Nils Hersch, Elena Naumovska, Thomas Gensch, Johnny Hendriks, Arne Franzen, Lisa Landvogt, Jan-Peter Siebrasse, Ulrich Kubitscheck, Bernd Hoffmann, Rudolf Merkel, and Agnes Csiszár Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.6b04304 • Publication Date (Web): 06 Jan 2017 Downloaded from http://pubs.acs.org on January 7, 2017

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Fusogenic Liposomes as Nano Carriers for Delivery of Intracellular Proteins

AUTHORS Sarah Kube1, Nils Hersch1, Elena Naumovska1, Thomas Gensch2, Johnny Hendriks2, Arne Franzen2, Lisa Landvogt3, Jan-Peter Siebrasse3, Ulrich Kubitscheck3, Bernd Hoffmann1, Rudolf Merkel1, and Agnes Csiszár1*

AUTHORS ADDRESS 1

Forschungszentrum Jülich GmbH, Institute of Complex Systems: Biomechanics (ICS-7),

Germany 2

Forschungszentrum Jülich GmbH, Institute of Complex Systems: Cellular Biophysics (ICS-

4), Germany 3

Friedrich-Wilhelms-University Bonn, Biophysical Chemistry, Germany

ABSTRACT Direct delivery of proteins and peptides into living mammalian cells has been accomplished using phospholipid liposomes as carrier particles. Such liposomes are usually taken up via endocytosis where the main part of their cargo is degraded in lysosomes before reaching its destination. Here, fusogenic liposomes, a newly developed molecular carrier system, were used for protein delivery. When such liposomes were loaded with water-soluble proteins and brought into contact with mammalian cells, the liposomal membrane efficiently fused with the cellular plasma membrane delivering the liposomal content into the cytoplasm without degradation. To explore the key factors of proteofection processes the complex formation of

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fusogenic liposomes and proteins of interest, as well as size and zeta potential of the formed fusogenic proteoliposoms were monitored. Intracellular protein delivery was analyzed using fluorescence microscopy and flow cytometry. Proteins like EGFP, Dendra2, R-phycoerythrin or peptides like LifeAct-FITC or NTF2-AlexaFluor488 were successfully incorporated into mammalian cells with high efficiencies. Moreover, correct functionality and faithful transport to binding sites were also proven for the imported proteins.

KEYWORDS intracellular protein delivery, proteofection, fusogenic proteoliposomes, membrane fusion, lipid-protein interactions

* corresponding author


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INTRODUCTION

The transport of proteins into mammalian cells is an essential biomedical, pharmacological and research tool.1 Because most proteins are not able to cross the plasma membrane a delivery strategy is needed for successful and efficient transmembrane transport. A variety of methods have been established for this purpose during the last decades. They are all governed by two physicochemical interactions: first, the interaction of the cargo with the carrier tool, and second, the specific interaction of the cargo-carrier complex with the cell membrane.2 Unfortunately, proteins have a broad distribution of surface charge, hydrophobicity and size. This spread of properties usually necessitates a laborious adaptation of conditions to the proteins of interest.

Despite some limitations, liposomes have been shown to be powerful tools for intracellular protein delivery. Liposomes are spherical vesicles composed of one or more phospholipid bilayers enclosing an aqueous volume.3 The main advantages of liposomal microcapsules are their high biocompatibility and low toxicity due to their natural components.4 Moreover, depending on the chemical characteristics a broad range of cargos can be transported using liposomes. Lipophilic molecules can be incorporated into the lipid bilayers while hydrophilic molecules, like water- soluble proteins, can be enclosed in the inner aqueous core.5 Even targeted liposomal delivery with cell specific localization has already been established.6 A non-exhaustive list of possible protein cargos has been published by Pachioni-Vasconceros et al.7 The major drawback of liposomes for delivery is that they are mainly taken up by endocytosis where the cargo finally accumulates in lysosomes and immediately degrades (Figure 1).8 Newly developed strategies overcome this limitation by inducing endosomal leakage, in consequence they have to deal with a more complex liposomal composition and an intervention into cell functionality.9


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Figure 1 Comparison of putative protein delivery by fusogenic and endocytotic liposomes. When fusogenic liposomes are loaded with water-soluble proteins, the protein cargo is delivered into the cell cytoplasm by membrane fusion directly upon contact. Alternatively, free protein molecules penetrate into the cell through transient membrane pores formed by cationic lipids in the membrane. As a result functional proteins enter the cell cytoplasm. Compared to this, commercial phospholipid vesicles (with or without charge) can also be loaded with proteins but such liposomes are usually taken up via endocytosis accomplished by degradation of most biomolecules.

To overcome these drawbacks liposomes with special fusogenic character10 are proposed as protein delivery carriers. Such fusogenic liposomes (FLs) have been established in the last years as appropriate delivery systems for lipophilic compounds.11-15 These unilamellar liposomes rapidly fuse with the cellular plasma membrane whereupon the lipophilic cargo is directly delivered into the cellular membrane.16 The low liposomal concentration as well as the very short incubation time of some minutes prevents usual cell toxicity of the positively charged lipid component like DOTAP.14-16 Here, our working hypothesis is that hydrophilic molecules loaded into the liposomal lumen are transferred during fusion events into the cellular cytoplasm17 as shown in Figure 1. Alternatively, free 4 Environment ACS Paragon Plus

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cargo molecules enter the cell cytoplasm through transient pores formed by cationic lipids in the cell membranes.18 This delivery strategy has the potential to avoid the endosomal degradation route and to prevent cargo degradation. Therefore, in this project we have used fusogenic liposomes to deliver water-soluble proteins into living mammalian cells. We have systematically varied protein size and charge to elucidate the driving force of proteoliposome formation. Protein size ranged from 2.3 kDa (fluorescently tagged LifeAct) over 27 kDa (EGFP) to 240 kDa (R-phycoerythrin). The influence of surface charge, or more specific, zeta potential was tested by comparing the delivery of streptavidin, neutravidin and avidin with comparable size but very different zeta potentials. Fusogenic proteoliposome complexes were first characterized and the delivery efficiency into mammalian cells, optimal delivery concentration as well as protein function after delivery were monitored using fluorescence microscopy and flow cytometry techniques.


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EXPERIMENTAL SECTION Lipids 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine

(DOPE)

and

1,2-dioleoyl-3-

trimethylammonium-propane, chloride salt (DOTAP) were purchased from Avanti Polar Lipids,

Inc.

(Alabaster,

dihexadecyloxacarbocyanine

AL,

USA).

The

perchlorate

fluorescent

(DiO)

and

membrane

tracers

3,3'-

1,1'-dioctadecyl-3,3,3',3'-

tetramethylindotricarbocyanine iodide (DiR) were ordered from Thermo Fischer Scientific (Eugene, OR, USA).

Proteins R-phycoerythrin (R-PE), LifeAct-FITC and LifAct-TMR and all avidin derivatives like streptavidin-AlexaFluor488, neutravidin-OregonGreen, and avidin-AlexaFluor488 were ordered from Thermo Fischer Scientific (Waltham, MA, USA). Human NTF2 (nuclear transport factor 2) was expressed as Glutathione-S-transferase (GST) fusion protein in E.coli BL21 (pLysS) and purified using Glutathion agarose (SigmaAldrich,). The GST moiety was removed by thrombin digestion. To improve fluorescence labeling an additional tetra-cystein tag (Cys-Cys-Pro-Gly-Cys-Cys) was introduced at the Nterminus of the hNTF2 and the protein was labeled with AlexaFluor488 maleimide dye from Thermo Fischer Scientific. To obtain the prokaryotic expression constructs of the fluorescent proteins EGFP and Dendra2, full-length coding sequences were amplified and modified by PCR techniques and finally cloned into vectors for bacterial expression. For pET-EGFP the coding region of EGFP was amplified by PCR from pEGFP-C1 vector (Clontech, California, USA) with 5’ primer (AAACaagcttTGATGGTGAGCAAGGGCGAG)

and

3’primer

(AAActcgagCTTGTACAGCTCGTCCATGCC) adding HindIII at the N-terminus and a Cterminal XhoI restriction site. The PCR fragment was then cloned by those restriction sites


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into pET21a vector (Novagen/Merck Millipore, Temecula, CA, USA) in frame with the provided His-Tag. The pET-Dendra2 construct was generated by PCR amplification of the coding region of pDendra2-N

plasmid

(Clontech,

California,

USA)

with

5’primer

(CACGAATTcatatgAACACCCCGGGAATTAAC) containing a NdeI restriction site and 3’ primer (GAGcctaggATTACTACCACTACCACTACGGTGTGGACCGACCCGTC) adding His-Tag and BamHI site in the C-terminus and cloned via NdeI and BamHI into pET11a vector (Novagen/Merck Millipore). Overexpression of recombinant proteins was performed in the E. coli strain BL21 (DE3) Codon Plus RIL or RP that was transformed with the corresponding DNA. The starter cultures were grown in Luria-Bertani-Medium (bactotrypton 10 g/l, NaCl 10 g/l, yeast extract 5 g/l) with antibiotics (ampicillin 100 µg/ml, chloramphenicol 18 µg/ml), grown at 37°C overnight and then transferred into 500 ml to reach an OD600 of ~ 0.03. The bacteria were grown until they reached an OD600 of ~0.4 - 0.6 and protein expression was then induced by addition of isopropyl-D-thiogalactopyranoside (IPTG, 1 mM f. c.). Expression was further carried out at 20°C overnight (~20 h). The cells were harvested, processed and purified according to the semi-batch purification protocol of polyhistidine-tagged proteins under native conditions using Protino®Ni-NTA agarose (Macherey-Nagel, Dueren, Germany). Subsequently the samples were concentrated. Simultaneously, the buffer was exchanged to phosphate buffered saline (PBS) with a centrifugal filter unit. Integrity and function of the proteins was judged by coomassie-blue stained SDS-PAGE gels and spectrometric analysis.

Preparation of fusogenic proteoliposomes (FPLs) Stock solutions of DOPE/DOTAP were prepared in chloroform with a weight ratio of 1/1 in a total lipid concentration of 1 mg/mL. Stock solutions of the fluorescent lipid tracers were also prepared in chloroform at a final concentration of 1 mg/mL. Solutions were kept at -20°C until further usage. FPLs were prepared by mixing the lipid components (DOPE/DOTAP) and 7 Environment ACS Paragon Plus

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the fluorescent dye DiO or DiR, respectively, in a ratio of 1/1/0.1 w/w. After evaporation of chloroform under vacuum for 10-20 min, lipids were resuspended in 20 mM HEPES buffer containing proteins or peptides at different concentrations at pH 7.4. Total lipid concentration was adjusted to 4 mg/mL. The suspension was intensively stirred for 1-5 min and incubated subsequently in an ultrasonic bath (Sonocool from Bandeline electronic GmbH, Berlin, Germany) for 15 min at 4°C to obtain small fusogenic proteoliposomes. Free protein molecules and proteins encapsulated into FLs were not separated from each other.

Characterization of proteins and fusogenic proteoliposomes (FPLs) Zeta potential measurements were performed using a zetasizer (Zetasizer Nano ZS from Malvern Instruments, UK) equipped with a HeNe laser (633 nm). Scattered laser light was collected at a constant angle of 173°. Prior to measurements liposome stock solutions were diluted 1/50 with purified, degassed and filtrated water. Protein concentration was adjusted to 0.1 mg/ml. All measurements were performed at 20°C and repeated three times at 1 min intervals. Data were collected from three independently prepared samples and analyzed using the instrument software (DTS from Malvern Instruments). Average values of three independent measurements and standard deviations were reported.

Cell culture Chinese hamster ovary cells (CHOs) were purchased from American Type Culture Collection (ATCC, Manassas, VA, USA). They were maintained in DMEM-F12 (Sigma-Aldrich) supplemented with 10% fetal bovine serum and a 1/100 dilution of an antibiotic solution (10,000 units penicillin and 10 mg/mL streptomycin in 0.9% NaCl, (Sigma-Aldrich)). During culture as well as experimental steps, cells were kept at 37°C and 5% CO2 in a saturated humid atmosphere. Cell density never exceeded 80% confluencey. For liposome fusion and microscopy, 30,000 cells were seeded on fibronectin coated glass surfaces (10 µg/ml human plasma fibronectin (BD Biosciences)) one day prior to the experiment.


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Cardiac fibroblasts were isolated from 18-day old Wistar rat embryos as described earlier.19 Cells were maintained in F10 Ham's medium (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% fetal bovine serum, a 1/100 dilution of an antibiotic solution (10,000 units penicillin and 10 mg/mL streptomycin in 0.9% NaCl, (Sigma-Aldrich)) and a 1/200 dilution of solution containing insulin (1 mg/mL), transferrin (10 µg/mL) and sodium selenite (0.5 µg/mL) in Earle's balanced salt solution (EBSS) (Sigma-Aldrich). During culture as well as experimental steps, cells were kept at 37°C and 5% CO2 in a saturated humid atmosphere. Cell density never exceeded 80% confluence. For liposome fusion and microscopy, 40,000 cells were seeded on fibronectin coated glass surfaces, similar to experiments used by CHO cell culture, one day prior to the experiment.

Protein delivery using FPLs For protein delivery experiments, 5 µL of FPL solution were diluted 1/100 with PBS (SigmaAldrich) at pH 7.4 and vortexed for 1-2 min at room temperature. Cells in a Petri dish (Ø=3.5 cm), were incubated in 0.5 mL of fusogenic liposome solution for 10-15 min at 37°C. Before analysis cells were washed three times with PBS and the buffer solution was replaced by fresh cell culture medium.

Imaging For cell imaging a confocal laser scanning microscope (cLSM 710 from Carl Zeiss MicroImaging GmbH, Jena, Germany) equipped with an argon ion laser (488 nm) and a helium-neon laser (633 nm) was used. To detect the lipid tracer DiR (ex. 633 nm) a long pass filter LP650 nm was chosen while DiO, R-PE, AlexaFluor488, EGFP, and Dendra2 signals (ex. 488 nm) were collected using a band pass filter BP 500-550. The microscope was equipped with an oil immersion objective EC plan-Neofluar 40x/1.30 Ph3. Live cell imaging was performed under cell culture conditions (37°C and 5% CO2). Image acquisition and analysis were done with the manufacture´s software (ZEN, Carl Zeiss).


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Photoconversion experiments with Dendra2 in CHO cells after protein delivery by FLs were carried out on a home-built wide-field fluorescence microscope with single molecule detection efficiency.20 Low power (1-2 mW) of 488 nm and 561 nm laser light were used to excite the ground and photoconverted state of Dendra2, respectively. Much lower power of 405 nm laser light was applied for photoconversion. Signals of unconverted and converted Dendra2 were detected at 500-540 nm and 580-620 nm, respectively.

Determination of intracellular protein concentration delivered by FPLs R-PE, EGFP and NTF2-ALexaFluor488 proteins were dissolved in PBS buffer (pH 7.4) at an initial concentration of 1 mg/ml. Protein solutions were diluted 1/500, 1/1000, 1/5000, and 1/10000 and pipetted on cover glass previously passivated with avidin solution (1 mg/ml) to prevent protein adsorption on the glass surface. Fluorescent intensities of proteins of interest at different concentrations were recorded using the same microscopy setup at the same settings as used for cell analysis. Fluorescence intensity vs. protein concentration calibration data were fitted by a linear equation. The fluorescent intensities of CHO cells in the protein channel were compared with signal intensities of the calibration curves. Intracellular protein concentrations were converted into concentrations using the respective fit results.

Colocalization analysis FPLs were prepared as described above using FLs containing DiR as lipid tracer and streptavidin-AlexaFluor488,

neutravidin-OregonGreen,

and

avidin-AlexaFluor488

as

proteins. The liposomes were used either for cellular treatment (described above) or for further colocalization analysis. To avoid a quick sedimentation of liposomes and proteins on the glass surface FPLs were immobilized in PBS buffer diluting 20 µl of the solution with 20

µl of aqueous mounting medium (Sigma-Aldrich), and the solution was covered by a cover slide. For recording the fluorescence signals of proteins and lipid tracer identical microscopy setup was used as described above. Using this imaging setup only larger liposome, protein, or liposome/protein aggregates could be imaged. 10 Environment ACS Paragon Plus

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Colocalization analysis was carried out using the colocalization software package of ZEN (Carl Zeiss). Signal thresholds were determined using samples containing only proteins or protein treated cells and liposomes or liposomes treated cells, respectively. Measurements were repeated on three independent samples at least on three spots in each. Analysis was carried out on the whole micrographs. Results were shown as mean ± standard deviation. As characteristic values colocalisation coeffitients m1 and m2 were used and defined as following:

௉௜௫௘௟௦

݉ଵ = ௉௜௫௘௟௦಴೓భ,೎೚೗೚೎ ಴೓భ,೟೚೟ೌ೗

݉ଶ =

௉௜௫௘௟௦಴೓మ,೎೚೗೚೎ ௉௜௫௘௟௦಴೓మ,೟೚೟ೌ೗

where Ch1 and Ch2 denoted the protein and the lipid microscopy channels, respectively.

Flow Cytometry A flow cytometer (Guava easyCyte 8HT from Merck Millipore, Billerica, MA, USA) was used to simultaneously analyze fusion efficiency and protein delivery. Prior to analysis cells were trypsinized with trypsin-EDTA solution (Sigma-Aldrich) for 3 min. Cells were subsequently washed with PBS (pH 7.4) and centrifuged. Analyses were carried out on living cells in DMEM medium without phenol red indicator. The lipid tracer DiR was exited at 633 nm and its fluorescence was collected using the band bass filter 785/70 nm. The lipid tracer DiO as well as the fluorescence signals of R-PE, EGFP, Dendra2 and the protein label AlexaFluor488 (excitation 488 nm) were collected through a band pass emission filter BP 525/30 nm. Data analysis was performed on 10,000 cells/sample using the manufactures software (InCyte from Merck Millipore). The cell population was gated based on the forward scatter vs. side scatter signals. Subsequent analysis was carried out exclusively on these cells. Fusion efficiency was calculated based on the recorded fluorescent signal of the lipid tracer


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while protein delivery efficiency was determined as green positive cells compared to control. Here, completely untreated cells as well as only fused cell were used as control sample.


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RESULTS AND DISCUSSION Proteins for delivery were chosen to cover molecular size from small peptide (around 2 kDa) to large proteins (up to 240 kDa) as well as molecular surface charges from negative over neutral to positive. The surface charge of proteins was characterized by measuring their electrophoretic light scattering and calculating their zeta potential (ζ). Protein zeta potentials varied in a range from -30 mV to +15 mV while the zeta potential of fusogenic liposomes (FLs) used as delivery carriers was constant at +75 ± 5 mV. Furthermore, because of our fluorescence analytical techniques proteins either with intrinsic chromophores (e.g. GFP) or extrinsic synthetic fluorophores (e.g. AlexaFluor488 or OregonGreen) were used. For protein emission a broad spectral range from green (500 nm) across orange to red (up to 700 nm) was chosen. Physicochemical properties of all proteins tested in this study are summarized in Table 1.

Table 1. Fluorescent proteins and peptides delivered by FLs. Molecular weight (MW), zeta potential (ζ), their used (Cused) and their optimal delivery concentrations (Copt) are given. For zeta potential means and standard deviations of three independent measurements are listed. Optimal delivery concentrations are given as a

Protein/Peptide

MW

ζ

Cused

Copt

[kDa]

[mV]

[µM]

[µM]

range.


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LifeAct-FITC

2.3

-26 ± 6

0.01-2

0.5-1

LifeAct-TMR

2.4

-22 ± 4

0.01-2

0.5-1

Dendra2

26

-24 ± 5

0.05-2

1-2

EGFP

27

-26 ± 4

0.05-2

0.3-2

NTF2-AlexaFluor 488

32

-5 ± 5

0.05-2

0.5-1

Streptavidin-AlexaFluor 488

55-60

-27 ± 4

0.05-1

0.05-0.5

Neutravidin-OregonGreen

57-60

-9 ± 4

0.05-1

0.05-0.5

Avidin-AlexaFluor 488

57-65

+4 ± 4

0.05-1

-

240

-8 ± 6

0.05-1

0.1-0.5

R-PE

Protein delivery using fusogenic liposomes (FLs) To incorporate the protein of interest into the carrier liposomes a dried lipid film composed of the fusogenic lipid mixture was resuspended in the protein solution and subsequently homogenized (see Material and Methods). The fusogenic lipid mixture contained DiR, an infrared fluorescent membrane tracer enabling simultaneous detection of lipid carrier and its cargo.16 Due to the long wavelength of DiR emission in the near IR spectral range (Emmax = 780 nm) lipid and protein signals were clearly separated from each other without spectral overlap. First, R-phycoerythrin (R-PE), a large protein with 240 kDa was enclosed in FLs. R-PE, a multimeric protein complex that is part of phycobilisomes. It binds several naturally occurring tetrapyrrole chromophores absorbing and emitting across a broad spectral range with an emission maximum at 576 nm.21 Because of the very low non-specific binding of this protein in mammalian cells it is frequently used to label antibodies for immunoassays.22 This large molecule is not able to pass mammalian plasma membranes without microinjection or molecular carriers.23 Therefore we tested the applicability of FLs as molecular carriers for RPE. Liposomes were first swollen in the protein solution and subsequently fused to Chinese hamster ovary cells (CHOs). Intracellular protein delivery was monitored by fluorescence


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microscopy. As reference sample cells treated with R-PE at the same concentration of 0.1 µM without liposomal carriers were used. Representative results are shown on Figure 2. As expected CHO cells remained unstained upon simple incubation with R-PE without carrier particles (Fig. 2 upper panel) while they were successfully loaded with this protein using FPLs (Fig. 2 lower panel: R-PE channel). Here, R-PE homogenously distributed in the cell cytoplasm of CHO cells without entering the nucleus. This specific distribution pattern reflects the low physicochemical interaction of the protein with the cellular environment, its putative lack of binding sites and its high molecular weight above the limit of passive transport into the nucleus.24 Simultaneously with the cellular R-PE signal a homogenous plasma membrane staining was detected in the DiR microscopy channel. Similar signal distribution was also found in previous experiments using FLs without cargo.16 This signal was a clear proof for membrane fusion between the liposomal carrier membranes and the cellular plasma membrane (Fig. 2 lower panel: DiR channel). The simultaneous intracellular R-PE signal and plasma membrane signal of the lipid tracer DiR confirmed the ability of FLs for cellular protein delivery.


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Figure 2 R-Phycoerythrin (R-PE) delivery using fusogenic proteoliposomes (FPLs). CHO cells were incubated with a solution of R-PE (top row) and R-PE encapsulated in FLs (bottom row) at the same concentration. Optical microscopy was used to record the lipid tracer DiR (left column), the protein fluorescence (middle column) and the general appearance of the cells via phase contrast (right column). Images were recorded in cell culture medium within 1 h after treatments. Scale bar, 100 µm, applies to all.

Size independent protein delivery Searching for the parameters controlling the suitability of FLs as carriers for specific proteins we first explored the influence of protein size. As shown above the large R-PE (240 kDa) was delivered with high efficiency into living cells by FPLs. Similar to R-PE smaller proteins like EGFP (27 kDa) or Dendra2 (26 kDa) with cell staining functions were also successfully inserted into the cellular cytoplasm of CHO cells (Figure 3). In contrast to R-PE, EGFP fluorescence was also detected within the nuclei (Figure 3 middle panel). This was caused most likely by the small size of this protein enabling it to pass passively through the


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nuclear pore complexes. The intensity of the staining varied from cell to cell. Surprisingly, this variation was dependent on the molecule delivered and highest for EGFP.

Figure 3 Delivery of fluorescent proteins of different sizes by FPLs. R-PE was homogenously incorporated into the cytoplasm of CHO cells at a concentration of 0.01-0.02 µM. Due to its molecular size (240 kDa) it was excluded from the cell nuclei (white arrows). The delivery concentration of EGFP spanned between 0.1 and 0.5 µM. EGFP molecules were able to passively diffuse through the nucleic pore complexes resulting in green fluorescence in the nuclei as well. NTF2-AlexaFluor 488 (32 kDa) was mainly localized in the nuclear membrane at a high equivalent concentration of around 1.5 µM while its cytoplasmic concentration remained below 0.7 µM (white arrows). This localization is due to its functional binding to the nuclear pore complex. Primary rat embryonic myofibroblasts were treated with FPLs containing the fluorescently labeled LifeAct-FITC pepdite. This F-actin binding peptide was mainly localized on the actin-cytoskeleton of the cells 5 min after delivery. Scale bars, 20 µm.

When proteins or small peptides with distinct biological functions or intracellular binding sites were transferred into mammalian cells using FLs they were observed at their specific binding sites after a short time period of minutes (1-5 min). For example, the nuclear transport factor NTF2 is responsible for the import of Ran proteins through the nuclear pore


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complex into the nucleus.25 Its fluorescent derivative, NTF2-AlexaFluor488, was delivered by FLs into CHO cells and its distribution was detected by fluorescence microscopy (Figure 3 right panel). After direct delivery into the cellular cytoplasm it progressively enriched in the nucleus and the nuclear membranes. Concomitantly with this nuclear signal increase, the cytoplasmic signal decreased. Specific binding to an intracellular target was also observed after delivery of the fluorescently labeled actin-binding peptide LifeAct-FITC26 into primary rat cardiac myofibroblasts. This cell type is well known for its characteristic and pronounced actin cytoskeleton making it appropriate for such analysis.27 Immediately after treatment the fluorescent LifeAct-FITC signal appeared on the actin cytoskeleton of myofibroblasts (Figure 3) while the membrane tracer was homogenously distributed in the cellular plasma membrane (Figure S1).

Cellular concentration of the delivered proteins and peptides After protein incorporation and subsequent molecular distribution the cellular protein concentration was also analyzed based on the fluorescent emission of the coupled chromophores. Intracellular protein concentrations were estimated from fluorescence intensities. For the conversion calibration data were used (see Experimental Section and Supporting information Figure S2). Because they were collected on homogenous solutions we give here equivalent concentrations for molecules binding to filaments and membranes. Moreover, our analysis did not consider the different point spread functions that belong to the different molecular distributions (1D, 2D and 3D). Our results showed that the larger protein molecules are, the lower the cellular concentration is reached by delivery (Figure 3). The largest tested protein, R-PE, was homogenously incorporated into the cytoplasm of CHO cells at a concentration of 0.01-0.02

µM. Its strong fluorescence signal arose from the high chromophore concentration per 18 Environment ACS Paragon Plus

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molecule.21 The delivered concentration of the smaller EGFP spanned between 0.1 and 0.5

µM, and therefore 10 times higher compared to that of R-PE, while the smallest NTF2AlexaFluo488 was localized in the cytoplasm at even higher concentrations of around 0.7

µM. After delivery of proteins directly into the cytoplasm of living cells these molecules were recognized and sorted to their binding sites by cellular machineries. The local enrichments of the delivered proteins or peptides resulted in high protein concentrations and clearly visible staining. For example, the local concentration of NTF2-AlexaFluo488 at sites of nuclear pore complexes was two times higher (ca. 1.5 µM) compared to its cytoplasmic concentration (0.7 µM) (Figure 3 and Figure S2). A similar effect with even higher enrichment at the stained structure compared to cytosol was observed for the actin binding LifeAct-FITC peptide (Figure 3 and Figure S1). In both cases the elevated local concentrations of proteins on their binding sites, here on the nuclear pore complex and the actin cytoskeleton, respectively, prove the functional incorporation of the proteins into mammalian cells as proven earlier using other delivery strategies. 26, 28-29

Charge dependent protein delivery

Electrical charge of proteins to be encapsulated is another key parameter expected to govern protein transfer by FLs. The overall delivery efficiency comprises two contributions: complex formation of proteins with FLs and the delivery of cargo from these proteoliposomal complexes. The first process can be positively affected by attractive electrostatic interactions between the two partners or negatively influenced by repulsive forces. FLs were positively charged with a zeta potential of +75 ± 5 mV. Most of the proteins of interest are negatively charged at physiological pH enabling an effective protein-liposome complex formation. To 19 Environment ACS Paragon Plus

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analyze the influence of protein charge on the delivery efficiency proteins with similar size but different charges are required. The biotin binding proteins, streptavidin, neutravidin and avidin completely fulfill these criteria. All three proteins have a molecular weight around 5565 kDa and a similar tetrameric structure. The only marked difference between the three proteins is their surface characteristics.30 The avidin surface is covered by carbohydrate moieties and was found to be slightly positively charged at pH 7.4 (ζ = +4 mV) due to its high isoelectric point of 10.5. Neutravidin is a deglycolysated version of avidin with an accordingly lower and only slightly negative charge at the same pH (ζ = -9 mV) (pI = 7.4). Similar to neutravidin streptavidin lacks carbohydrate modifications but it is more negatively charged at physiological pH (ζ = -27 mV) because of its low pI (pI = 5.3).

All three proteins were commercially available with similar fluorescent dyes covalently attached. Their deliveries into the cytoplasm of CHO cells were monitored using confocal fluorescence microscopy. In all three cases initial protein concentration was kept at the same value of 0.1 mg/ml. As shown in Figure 4 homogenous and efficient membrane fusion occurred in all three cases. Due to simultaneous protein delivery the negatively charged streptavidin-AlexaFluor488 was homogenously distributed in the cytoplasm (top row, protein channel in Figure 4). To quantify the degree of protein and lipid signal overlaps after delivery signal colocalization was determined using a scatter diagram. This was generated by plotting each pixel in the fluorescence image based on its intensity level from each channel. The color in this scatter plot (Figure 4 right column) represented the number of pixels plotted in that region. To quantify the colocatization of protein and lipid signals 4 regions on the scatter plot were defined: 1 – background, 2 –lipid signal without protein signal (fusion without protein delivery), 3 – protein signal without lipid signal (protein uptake without fusion), and 4 – protein and lipid signals together (protein delivery due to membrane fusion). The calculated colocalization coefficients, m1 and m2, (summarized in Table 2) described the contribution of 20 Environment ACS Paragon Plus

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each one from the two selected channels to the pixels of interest.31 In the case of streptavidin the correlation of protein signal with the lipid signal raised over 90% indicating a very effective streptavidin delivery by FLs while only 60% of the membrane staining overlapped with the delivered protein signal.

Compared to this the chemically surface-modified neutravidin, neutravidinOregonGreen, was taken up by cells at an overall lower concentration and the calculated protein and membrane signal correlations were reduced to 83% and 18%, respectively. Furthermore, the positively charged avidin-AlexaFluor488 was completely excluded from CHO cells (bottom row, protein marker channel in Figure 4) and the free avidin molecules mainly adhered on the glass surface while FLs highly efficiency stained the cellular plasma membranes. Correspondingly, signal colocalizations remained around 20% (Table 2).

In order to understand our results on the different biotin-binding proteins we investigated protein incorporation into fusogenic liposomes by imaging proteoliposome complexes and subsequently analyzing the protein and lipid spatial colocalization. Colocalization plots are shown in supporting information Figure S3 and results are summarized in Table 3. The highest colocalization between protein and liposomes was found in the case of streptavidin where 44% of the green streptavidin pixels colocalized with the red liposomal pixels and 50% of liposomes interacted with the negatively charged streptavidin (see Table 3). Compared to this, only 11% of the neutral neutravidin signal overlapped with the liposomal signal while similar part of liposomes (56%) bound neutravidin. The positively charged avidin showed the lowest colocalization with the similarly charged liposomes (12%) and only 26% of the liposomes interacted with the avidin.


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Figure 4. Cellular delivery of fluorescent proteins with different isoelectric points. Fusogenic proteoliposomes were formed in presence of fluorescently labeled streptavidin-AlexaFluor488 (pI 5.3) (top row), neutravidin-OregonGreen (pI 7.4) (middle row), and avidin-AlexaFluor488 (pI 10.4) (bottom row), respectively, and incubated with CHO cells. Delivered protein fluorescence was detected in the green channel while membrane marker (DiR) signal was recorded in the red channel 0.5 h after treatment. Scale bars, 20 µm. Colocalization plots of the two fluorescence signals are shown in the right column. White crosshair divides the scatter diagrams into the following ranges:: 1 – background, 2 –fusion without protein delivery, 3 – protein uptake without fusion, and 4 –protein delivery due to membrane fusion. Color scale bar represents the relative frequency of detected signals.

Table 2. Colocalization of the delivered avidin- like proteins with the membrane staining signal in CHO cells. m1 represents the protein fraction colocalized with FLs while m2 shows the part of FLs containing protein. Results are listed as mean ±

sd. (N=12). 22 Environment ACS Paragon Plus

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Protein in Cells

m1±sd

m2±sd

Streptavidin-AlexaFluor488

0.93 ± 0.08

0.60 ± 0.17


0.83 ± 0.14

0.18 ± 0.14

Avidin-AlexaFluor488

0.21 ± 0.17

0.16 ± 0.09

Table 3. Colocalization of FPLs loaded with avidin- like proteins. m1 represents the protein fraction colocalized with FLs while m2 shows the part of FLs containing protein. Results are listed as mean ±

Protein in FLs

m1±sd

m2±sd

Streptavidin-AlexaFluor488

0.44 ± 0.09

0.50 ± 0.33


0.11 ± 0.10

0.56 ± 0.38

Avidin-AlexaFluor488

0.12 ± 0.13

0.26 ± 0.07

sd. (N=6).

These data show that the electrostatic interaction between cargo and carrier molecules plays a crucial role for successful protein delivery. Most likely, proteins first attach to the delivery vehicles driven by attractive electrostatic forces. Lacking such forces, for example in the case of avidin, FPLs complex formation was hampered and therefore significantly less avidin was incorporated into the FLs. The reduced amount of proteins in the proteoliposomes consequently resulted in missing protein delivery. Simultaneously, liposomes without cago still fused with cells.

Because the fusogenic carrier particles were positively charged only negatively charged proteins could be successfully delivered this way. Still, the surface charge or zeta potential of proteins strongly depends on the relation of the actual pH value and the pI of proteins. The pI of a protein is defined as the pH value, where the protein has no net charge. At pH values below or above their pI proteins are negatively or positively charged, respectively. The pH inside the fusogenic liposomes is chosen as 7.4 to match the cytosolic pH of mammalian cells. In case of protein delivery to cells with proteins that have a modestly 23 Environment ACS Paragon Plus

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basic pI (pH 8-8.5), one may try to apply a loading solution with a pH slightly larger than the pI to allow attractive electrostatic interaction, hence improve protein incorporation into FLs and subsequently protein delivery.

Protein delivery efficiency

In our experiments we observed an upper limit for the useful protein concentration during FL loading. Above this concentration the fusogenic potential is hampered. To analyze this effect in more detail we performed flow cytometry experiments enabling fast and statistically significant quantification of membrane fusion and protein delivery at the same time. Moreover, electrophoretic light scattering was employed to characterize the zeta potential of protein loaded liposomes. Due to its small size (26 kDa) and negative zeta potential at physiological pH (ζ =-24 ± 5 mV) we selected the photoswitchable fluorescent protein Dendra2 for these experiments.32 We first systematically varied the Dendra2 concentration during complexation with a constant amount of liposomes. Complex formation was followed by zeta potential measurements (ζDendra2/FL). As shown in Figure 5A FLs without protein cargo had a zeta potential of +75 mV. With increasing Dendra2 concentrations this value continuously decreased because of the adhesion of Dendra2 to the liposomal surface. At a protein concentration of 3.7 µM the surface charge of the FPLs dropped to zero.

Subsequently, the same liposomes were used for Dendra2 delivery into the cytoplasm of CHO cells and liposomal fusion efficiency as well as Dendra2 delivery efficiency were determined using flow cytometry. Typical flow cytometric results are shown in Figure 5D. Here, the fluorescent liposomal tracer DiR was used to monitor successful liposomal fusion with CHO cells while protein delivery efficiency was followed in the Dendra2 channel. The 24 Environment ACS Paragon Plus

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two channels were plotted against each other, where each dot represented one single cell. When no delivery happened both signals remained in the lower left quadrant of the plot (untreated cells in Figure 5D-I.). In the case of efficient fusion without protein delivery a high DiR signal and an unchanged low Dendra2 signal were recorded and the cell population appeared in the lower right quadrant of the plot (Fused cells in Figure 5D-II.). Efficient protein delivery upon membrane fusion resulted in increased red as well as green signals, and the cell population was shifted into the upper right quadrant of the plot (Deandra2 loaded cells in Figure 5D-III.). This analysis enabled the simultaneous quantification of fusion and protein delivery efficiencies, i.e. the fraction of cells detected positive in DiR and Dendra2 channels, respectively.

As shown in Figure 5B fusion efficiency continuously decreased with increasing protein concentration and decreasing zeta potential (Figure 5A). As consequence, fusion efficiency was recorded more then 50% if zeta potential of FLs reached slightly negative values. This decrease in fusogenity went along with significantly reduced amount of transferred Dendra2 proteins (Figure 5C). The data therefore point to a crucial role of electrostatic interaction for the docking of protein-loaded FLs to the cell membrane. Only if a certain positive surface potential of the particles was left (here +25 mV) they could effectively attach to the negatively charged cell surface to subsequently fuse and deliver their cargo.


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Figure 5 Analysis of Dendra2 delivery by FPLs. A) Zeta potential of FPLs, B) fusion efficiency, and C) Dendra2 delivery efficiencies at different concentration of Dendra2. Error bars represent standard deviation of at least five independent measurements. D) To determine fusion and protein delivery efficiencies untreated, FL and Dendra2-PFL treated CHO cells were analyzed using flow cytometry and the fusion signals vs. protein signals were plotted. Red lines separate quadrants used for frequency counts: I. - untreated cells, II. - fused cells, and III. -fused and Dendra2 loaded cells. E) Fluorescence and phase contrast micrographs of fused and Dendra2- loaded CHO cells. F) CHO cells imaged during photoactivation of Dendra2 molecules from the inactive to the active state using 405 nm laser 0.5 h after cellular treatment. The average intensity change of both cells of the orange channel (580-620 nm) vs. time was plotted. Blue arrow indicates the conversion starting point at 405 nm laser line. Scale bars, 20 µm.


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At high protein concentrations the efficiency of protein delivery falls in parallel with fusion efficiency as expected (Figure 5B and C). However, it also decreased at low protein concentrations despite close to 100% fusion efficiency. This apparent contradiction could be explained by the low intensity cutoff of flow cytometric detection. In other words, we assume that at low Dendra2 concentrations the protein molecules were transferred to cells but their fluorescent signal was simple too low to shift the green signal above the detection limit. Therefore, cells loaded with small amounts of protein and without protein were not effectively distinguished from each other. However, using fluorescence techniques for protein signal detection the fluorescence detection limit is unavoidable and has to be considered.

Dendra2 delivery upon fusion was also visualized using fluorescence microscopy as shown in Figure 5E. Similar to our previous experiments lipid tracer and protein signals were not colocalized in the cells after successful delivery. The membrane tracer DiR was mainly localized in the plasma membrane of CHO cells while the Dendra2 signal was distributed in the cell cytoplasm. Phase contrast micrographs confirmed healthy cell morphology.

To determine protein functionality after successful cellular incorporation we analyzed the photoconversion of Dendra2 in CHO cells. Dendra2 undergoes irreversible photoconversion by 405 nm light irradiation resulting in an increase in orange fluorescence3233

as shown in Figure 5F. Our observation of successful photoconversion indicates that the

protein function is not impaired by delivery via fusogenic liposomes. Therefore the full potential of photoswichable proteins can be used in untransfected cells.34 35


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CONCLUSIONS Here we demonstrated that fusogenic liposomes are efficient carriers for protein transport into living mammalian cells. Small peptides as well as large proteins were used without detectable limitation of molecular size. However, protein charge strongly influenced proteoliposome formation. Attractive electrostatic interactions between the positively charged carriers and the negatively charged cargo resulted in an effective complex formation while repulsive interactions between positively charged proteins and the similarly charged liposomes completely prevented this process. When living mammalian cells were treated with fusogenic proteoliposomes the carrier membrane first fused with the cellular plasma membrane and the protein cargos were released into the cellular cytoplasm. Proteins with biological functions in cells were subsequently transported to their binding sites by cellular machineries without any functional changes. Here we tested fluorescent or fluorescently labeled proteins. However, as formation of proteoliposomes and subsequent fusion with cellular membranes do not depend on protein fluorescence, it is obvious that this method can be used for the intracellular delivery of non-fluorescent proteins as well.

SUPPORTING INFORMATION Figure S1: Imaging of the actin cytoskeleton by LifeAct delivered by FPLs. Figure S2 Determination of intracellular peptide concentration delivered by FPLs. Figure S3 Colocalization analysis of FPLs.


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REFERENCES (1.) Gregoriadis, G. Liposome technology; 3 ed.; Informa Healthcare: New York, 2007; Vol. 2. (2.) Hapala, I. Breaking the barrier: methods for reversible permeabilization of cellular membranes. Crit Rev Biotechnol 1997, 17 (2), 105-22. (3.) Cevc, G. Phospholipids handbook; 1 ed.; Marcel Dekker, Inc.: New Yourk, Basel, 1993. (4.) Sercombe, L.; Veerati, T.; Moheimani, F.; Wu, S. Y.; Sood, A. K.; Hua, S. Advances and Challenges of Liposome Assisted Drug Delivery. Front Pharmacol 2015, 6, 286. (5.) Martins, S.; Sarmento, B.; Ferreira, D. C.; Souto, E. B. Lipid-based colloidal carriers for peptide and protein delivery--liposomes versus lipid nanoparticles. Int J Nanomedicine 2007, 2 (4), 595-607. (6.) Lasic, D. D.; Vallner, J. J.; Working, P. K. Sterically stabilized liposomes in cancer therapy and gene delivery. Curr Opin Mol Ther 1999, 1 (2), 177-85. (7.) Pachioni-Vasconcelos Jde, A.; Lopes, A. M.; Apolinario, A. C.; Valenzuela-Oses, J. K.; Costa, J. S.; Nascimento Lde, O.; Pessoa, A.; Barbosa, L. R.; Rangel-Yagui Cde, O. Nanostructures for protein drug delivery. Biomater Sci 2016, 4 (2), 205-18. (8.) Straubinger, R. M.; Hong, K.; Friend, D. S.; Papahadjopoulos, D. Endocytosis of liposomes and intracellular fate of encapsulated molecules: encounter with a low pH compartment after internalization in coated vesicles. Cell 1983, 32 (4), 1069-79. (9.) Erazo-Oliveras, A.; Najjar, K.; Dayani, L.; Wang, T. Y.; Johnson, G. A.; Pellois, J. P. Protein delivery into live cells by incubation with an endosomolytic agent. Nat Methods 2014, 11 (8), 861-7. (10.) Csiszar, A.; Hersch, N.; Dieluweit, S.; Biehl, R.; Merkel, R.; Hoffmann, B. Novel Fusogenic Liposomes for Fluorescent Cell Labeling and Membrane Modification. Bioconjugate Chem 2010, 21 (3), 537-543. (11.) Csiszar, A.; Csiszar, A.; Pinto, J. T.; Gautam, T.; Kleusch, C.; Hoffmann, B.; Tucsek, Z.; Toth, P.; Sonntag, W. E.; Ungvari, Z. Resveratrol encapsulated in novel fusogenic liposomes activates Nrf2 and attenuates oxidative stress in cerebromicrovascular endothelial cells from aged rats. J Gerontol A Biol Sci Med Sci 2015, 70 (3), 303-13. (12.) Dutta, D.; Pulsipher, A.; Luo, W.; Mak, H.; Yousaf, M. N. Engineering cell surfaces via liposome fusion. Bioconjug Chem 2011, 22 (12), 2423-33. (13.) Dutta, D.; Pulsipher, A.; Luo, W.; Yousaf, M. N. Synthetic chemoselective rewiring of cell surfaces: generation of three-dimensional tissue structures. J Am Chem Soc 2011, 133 (22), 8704-13. (14.) Hersch, N.; Wolters, B.; Ungvari, Z.; Gautam, T.; Deshpande, D.; Merkel, R.; Csiszar, A.; Hoffmann, B.; Csiszar, A. Biotin-conjugated fusogenic liposomes for high-quality cell purification. J Biomater Appl 2016, 30 (6), 846-856. (15.) Naumovska, E.; Ludwanowski, S.; Hersch, N.; Braun, T.; Merkel, R.; Hoffmann, B.; Csiszar, A. Plasma membrane functionalization using highly fusogenic immune activator liposomes. Acta Biomater 2014, 10 (3), 1403-1411. (16.) Kleusch, C.; Hersch, N.; Hoffmann, B.; Merkel, R.; Csiszar, A. Fluorescent Lipids: Functional Parts of Fusogenic Liposomes and Tools for Cell Membrane Labeling and Visualization. Molecules 2012, 17 (1), 1055-1073. (17.) Lira, R. B.; Seabra, M. A.; Matos, A. L. L.; Vasconselos, J. V.; Bezerra, D. P.; de Paula, E.; Santos, B. S.; Fontes, A. Studies on intracellular delivery of carboxyl-coated CdTe quantum dots mediated by fusogenic liposomes. . Journal of Materials Chemistry B 2013, 1, 4297.


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(18.) Simberg, D.; Weisman, S.; Talmon, Y.; Barenholz, Y. DOTAP (and other cationic lipids): chemistry, biophysics, and transfection. Crit Rev Ther Drug Carrier Syst 2004, 21 (4), 257-317. (19.) Hersch, N.; Wolters, B.; Dreissen, G.; Springer, R.; Kirchgessner, N.; Merkel, R.; Hoffmann, B. The constant beat: cardiomyocytes adapt their forces by equal contraction upon environmental stiffening. Biol Open 2013, 2 (3), 351-61. (20.) Tang, Y.; Dai, L.; Zhang, X.; Li, J.; Hendriks, J.; Fan, X.; Gruteser, N.; Meisenberg, A.; Baumann, A.; Katranidis, A.; Gensch, T. SNSMIL, a real-time single molecule identification and localization algorithm for super-resolution fluorescence microscopy. Sci Rep 2015, 5, 11073. (21.) Glazer, A. N. Light harvesting by phycobilisomes. Annu Rev Biophys Biophys Chem 1985, 14, 47-77. (22.) Oi, V. T.; Glazer, A. N.; Stryer, L. Fluorescent phycobiliprotein conjugates for analyses of cells and molecules. J Cell Biol 1982, 93 (3), 981-6. (23.) Goulian, M.; Simon, S. M. Tracking single proteins within cells. Biophys J 2000, 79 (4), 2188-98. (24.) Mohr, D.; Frey, S.; Fischer, T.; Guttler, T.; Gorlich, D. Characterisation of the passive permeability barrier of nuclear pore complexes. EMBO J 2009, 28 (17), 2541-53. (25.) Ribbeck, K.; Lipowsky, G.; Kent, H. M.; Stewart, M.; Gorlich, D. NTF2 mediates nuclear import of Ran. EMBO J 1998, 17 (22), 6587-98. (26.) Riedl, J.; Crevenna, A. H.; Kessenbrock, K.; Yu, J. H.; Neukirchen, D.; Bista, M.; Bradke, F.; Jenne, D.; Holak, T. A.; Werb, Z.; Sixt, M.; Wedlich-Soldner, R. Lifeact: a versatile marker to visualize F-actin. Nat Methods 2008, 5 (7), 605-7. (27.) Smith-Clerc, J.; Hinz, B. Immunofluorescence detection of the cytoskeleton and extracellular matrix in tissue and cultured cells. Methods Mol Biol 2010, 611, 43-57. (28.) Siebrasse, J. P.; Kaminski, T.; Kubitscheck, U. Nuclear export of single native mRNA molecules observed by light sheet fluorescence microscopy. Proc Natl Acad Sci U S A 2012, 109 (24), 9426-31. (29.) Kubitscheck, U.; Grunwald, D.; Hoekstra, A.; Rohleder, D.; Kues, T.; Siebrasse, J. P.; Peters, R. Nuclear transport of single molecules: dwell times at the nuclear pore complex. J Cell Biol 2005, 168 (2), 233-43. (30.) Green, N. M. Avidin and streptavidin. Methods Enzymol 1990, 184, 51-67. (31.) Zinchuk, V.; Zinchuk, O.; Okada, T. Quantitative colocalization analysis of multicolor confocal immunofluorescence microscopy images: pushing pixels to explore biological phenomena. Acta Histochem Cytochem 2007, 40 (4), 101-11. (32.) Gurskaya, N. G.; Verkhusha, V. V.; Shcheglov, A. S.; Staroverov, D. B.; Chepurnykh, T. V.; Fradkov, A. F.; Lukyanov, S.; Lukyanov, K. A. Engineering of a monomeric green-tored photoactivatable fluorescent protein induced by blue light. Nat Biotechnol 2006, 24 (4), 461-5. (33.) Berardozzi, R.; Adam, V.; Martins, A.; Bourgeois, D. Arginine 66 Controls DarkState Formation in Green-to-Red Photoconvertible Fluorescent Proteins. J Am Chem Soc 2016, 138 (2), 558-65. (34.) Jasik, J.; Boggetti, B.; Baluska, F.; Volkmann, D.; Gensch, T.; Rutten, T.; Altmann, T.; Schmelzer, E. PIN2 turnover in Arabidopsis root epidermal cells explored by the photoconvertible protein Dendra2. PLoS One 2013, 8 (4), e61403. (35.) Chudakov, D. M.; Lukyanov, S.; Lukyanov, K. A. Tracking intracellular protein movements using photoswitchable fluorescent proteins PS-CFP2 and Dendra2. Nat Protoc 2007, 2 (8), 2024-32.


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TABLE OF CONTENTS GRAPHIC


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Figure 1 Comparison of putative protein delivery by fusogenic and endocytotic liposomes. When fusogenic liposomes are loaded with water-soluble proteins, the protein cargo is delivered into the cell cytoplasm by membrane fusion directly upon contact. Alternatively, free protein molecules penetrate into the cell through transient membrane pores formed by cationic lipids in the membrane. As a result functional proteins enter the cell cytoplasm. Compared to this, commercial phospholipid vesicles (with or without charge) can also be loaded with proteins but such liposomes are usually taken up via endocytosis accomplished by degradation of most biomolecules. 298x171mm (300 x 300 DPI)

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Figure 2 R-Phycoerythrin (R-PE) delivery using fusogenic proteoliposomes (FPLs). CHO cells were incubated with a solution of R-PE (top row) and R-PE encapsulated in FLs (bottom row) at the same concentration. Optical microscopy was used to record the lipid tracer DiR (left column), the protein fluorescence (middle column) and the general appearance of the cells via phase contrast (right column). Images were recorded in cell culture medium within 1 h after treatments. Scale bar, 100 µm, applies to all. 710x480mm (72 x 72 DPI)


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Figure 3 Delivery of fluorescent proteins of different sizes by FPLs. R-PE was homogenously incorporated into the cytoplasm of CHO cells at a concentration of 0.01-0.02 µM. Due to its molecular size (240 kDa) it was excluded from the cell nuclei (white arrows). The delivery concentration of EGFP spanned between 0.1 and 0.5 µM. EGFP molecules were able to passively diffuse through the nucleic pore complexes resulting in green fluorescence in the nuclei as well. NTF2-AlexaFluor 488 (32 kDa) was mainly localized in the nuclear membrane at a high equivalent concentration of around 1.5 µM while its cytoplasmic concentration remained below 0.7 µM (white arrows). This localization is due to its functional binding to the nuclear pore complex. Primary rat embryonic myofibroblasts were treated with FPLs containing the fluorescently labeled LifeAct-FITC pepdite. This F-actin binding peptide was mainly localized on the actin-cytoskeleton of the cells 5 min after delivery. Scale bars, 20 µm. 750x440mm (72 x 72 DPI)


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Figure 4. Cellular delivery of fluorescent proteins with different isoelectric points. Fusogenic proteoliposomes were formed in presence of fluorescently labeled streptavidin-AlexaFluor488 (pI 5.3) (top row), neutravidinOregonGreen (pI 7.4) (middle row), and avidin-AlexaFluor488 (pI 10.4) (bottom row), respectively, and incubated with CHO cells. Delivered protein fluorescence was detected in the green channel while membrane marker (DiR) signal was recorded in the red channel 0.5 h after treatment. Scale bars, 20 µm. Colocalization plots of the two fluorescence signals are shown in the right column. White crosshair divides the scatter diagrams into the following ranges: 1 – background, 2 –fusion without protein delivery, 3 – protein uptake without fusion, and 4 –protein delivery due to membrane fusion. Color scale bar represents the relative frequency of detected signals. 383x420mm (72 x 72 DPI)


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Figure 5 Analysis of Dendra2 delivery by FPLs. A) Zeta potential of FPLs, B) fusion efficiency, and C) Dendra2 delivery efficiencies at different concentration of Dendra2. Error bars represent standard deviation of at least five independent measurements. D) To determine fusion and protein delivery efficiencies untreated, FL and Dendra2-PFL treated CHO cells were analyzed using flow cytometry and the fusion signals vs. protein signals were plotted. Red lines separate quadrants used for frequency counts: I. - untreated cells, II. - fused cells, and III. -fused and Dendra2 loaded cells. E) Fluorescence and phase contrast micrographs of fused and Dendra2- loaded CHO cells. F) CHO cells imaged during photoactivation of Dendra2 molecules from the inactive to the active state using 405 nm laser 0.5 h after cellular treatment. The average intensity change of both cells of the orange channel (580-620 nm) vs. time was plotted. Blue arrow indicates the conversion starting point at 405 nm laser line. Scale bars, 20 µm. 904x899mm (72 x 72 DPI)


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Langmuir

TOC Graphic 298x171mm (300 x 300 DPI)


Fusogenic Liposomes as Nanocarriers for the Delivery of Intracellular

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