Name: delivery of proteins, peptides or cell-impermeable small molecules into live cells by incubation with the endosomolytic reagent dftat
Uploaded: 2015-09-02T15:00:00.000+00:00
Duration: 10 min 30 s
Description: Watch this Scientific Journal Video about Delivery of Proteins, Peptides or Cell-impermeable Small Molecules into Live Cells by Incubation with the Endosomolytic Reagent dfTAT at JoVE.com

This article was supported by Award Number R01GM087227 from the National Institute of General Medical Sciences, the Norman Ackerman Advanced Research Program, and the Robert A. Welch foundation (Grant A-1769).

1. SPPS: CK(TMR)TATG (fTAT) Synthesis
Note: dfTAT is produced in two steps: synthesis of the monomer fTAT by solid phase peptide synthesis followed by disulfide bond dimerization to form dfTAT.
<ol>
	<li>Swell 500 mg of rink amide MBHA resin in dimethylformamide (DMF) in a standard 50 ml SPPS vessel for 1 hr.</li>
	<li>Perform Fmoc deprotection by incubating the Fmoc protected resin in a 20% piperidine solution (20% piperidine in DMF, 10 ml (0.30 mmol). Carry out deprotection steps twice (1 x 5 min and 1 x 15 min) with a washing step using DMF (the washing step includes rinsing the resin multiple times with a total volume of approximately 150 ml of DMF and removing the solvent by vacuum filtration) in between reactions.</li>
	<li>Synthesize CK(TMR)RKKRRQRRRG (fTAT) on the rink amide MBHA resin. Use the following Fmoc protected amino acids: Fmoc-Lys(Mtt)-OH (only on N-terminus), Fmoc-Lys(Boc)-OH, Fmoc-Gly-OH, Fmoc-Arg(Pbf)-OH, Fmoc-Gln(Trt)-OH, and Fmoc-Cys(Trt)-OH. Carry out the reaction using N2 (20 PSI) to agitate the reaction. Perform all reactions at RT.
	<ol>
		<li>Carry out each amino acid coupling reaction for 4 hr. For each coupling, use the Fmoc protected amino acid (1.2 mmol), N,N,N &#8242; ,N &#8242; -Tetramethyl- O- (1 H -benzotriazol-1-yl)uronium hexafluorophosphate (HBTU) (0.44 g, 1.1 mmol), and diisopropylethylamine (DIEA) (0.51 ml, 3.0 mmol) dissolved in DMF. Following the 4 hr coupling, wash the resin with DMF and perform the Fmoc deprotection step as described in step 1.2.</li>
		<li>Repeat until the linear peptide chain of fTAT (Linear peptide chain: CKRKKRRQRRRG no TMR) was synthesized.</li>
		<li>Then wash resin thoroughly using first DMF and following that with dichloromethane (DCM). Rinse the resin multiple times with a total volume of approximately 150 ml of DMF or DCM and remove the solvent by vacuum filtration.</li>
		<li>Use MALDI-TOF to confirm that the peptide obtained has the correct molecular weight.
		<ol>
			<li>Make a matrix mix by adding 1 mg of matrix to a solution composed of 50 &#181;l of 0.01% TFA in water solution and 50 &#181;l of acetonitrile.</li>
			<li>To confirm linear peptide chain mass, use &#945;-Cyano-4-hydroxycinnamic acid. Run an analytical HPLC of the crude peptide and collect 50 &#181;l from the top of the pure peak.</li>
			<li>To prepare the sample to plate on the MALDI plate: Add 8 &#181;l of the peptide to 2 &#181;l of the matrix solution and place on the MALDI plate to air dry or on a 37 &#176;C heat plate. 
			Note: Arginine is known to be an amino acid difficult to couple in SPPS. Successful coupling is established by performing a Kaiser test14 (e.g., ninhydrin test). This test allows for the detection of free amines that might remain uncoupled on the resin. In case a blue color is detected (indicative of the presence of free amines), coupling steps are repeated.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>For CK(-NH-TMR)TATG (fTAT), cleave the Mtt protecting group at the -amino group of Lys on CK(-NH-Mtt)TATG with a solution composed of 1% trifluoroacetic acid (TFA) and 2% triisopropylsilane (TIS) in DCM.
	<ol>
		<li>As the removal of MTT protecting group would result in the appearance of a yellow color, incubate the resin with 20 ml of the above solution for 5 min, repeat this until no yellow color is observed. Wash the resin with DCM and DMF in between. Additionally since TIS is a scavenger that would scavenge the MTT, once no yellow color appears in the solution in the presence of MTT.</li>
		<li>Add an additional solution with 1% TFA in DCM (no TIS) to the resin to insure no more MTT is removed and the solution remains clear.</li>
	</ol>
	</li>
	<li>Dissolve the three components: Carboxytetramethylrhodamine (TMR), HBTU, and DIEA (4, 3.9 and 10 equivalence in respect to the amount of resin (in mg)) in DMF and add this mixture to the resin and carry out the reaction O/N using dry N2 to provide agitation.</li>
	<li>Following Fmoc-deprotection and amino acid conjugation, wash the resin with DCM and allow it to dry. For complete cleavage of the peptide from the resin, add a solution containing 92.5% TFA, 2.5% H2O, 2.5%, TIS, and 2.5% ethanedithiol (EDT) (total volume = 4 ml) to the peptidyl-resin for 3 hr at RT to achieve global deprotection and cleavage from the resin.
	<ol>
		<li>Precipitate the crude peptide products using cold anhydrous ethyl ether (Et2O) by draining the solution from step 1.6 into 40 ml of Et2O, spin this down at 4 &#176;C and 4,000 x g for 20 - 25 min. Repeat this step to allow washing of the precipitate with cold anhydrous Et2O.</li>
		<li>Resuspend the precipitates in water (5 ml or until precipitate is completely dissolved) and lyophilize. Then resuspend the products obtained in 0.1% aqueous TFA/acetonitrile.</li>
	</ol>
	</li>
	<li>Perform HPLC analysis with an analytical C18 column (5 &#181;m, 4 x 150 mm) to analyze each peptide. Use a flow rate of 1 ml/min, and detection at 214 nm and 550 nm.</li>
	<li>Perform semi-preparative HPLC on a C18 10 x 250 mm column for peptide purification. Use a flow rate of 4 ml/min, and detection at 214 nm and 550 nm. For all runs, use linear gradients of 0.1% aqueous TFA (solvent A) and 90% acetonitrile, 9.9% water, and 0.1% TFA (solvent B).</li>
	<li>Confirm the correct identity of the peptides by MALDI-TOF according to manufacturer&#8217;s protocol: fTAT, expected mass: 2,041.17, observed mass: 2040.66. DEAC-K9, expected mass: 1,412.97, observed mass: 1,415.59. Use an &#945;- cyano-4-hydroxycinnamic acid matrix for the MALDI-TOF.</li>
</ol>
2. Oxidation Reaction: dfTAT Generation
<ol>
	<li>Aerate phosphate buffered saline (PBS) pH 7.4 for 1 hr (at least 5 ml for the reaction below).</li>
	<li>Dissolve fTAT (0.3 mg, 1.5 x 10-4 mmol) in aerated PBS pH 7.4 (5 ml). Make sure that the pH is between 7.0 - 7.5 after the addition of the peptide. If not add sodium hydroxide (1 M, in small increments 1 - 5 &#181;l) to bring the pH back up to 7.4.</li>
	<li>Note: Oxygen dissolved in PBS acts to oxidize the thiol groups on fTAT and form a disulfide bond.</li>
	<li>Nutate the reaction O/N to allow it to react until completion (100% yield based on HPLC analysis). Purify the product using reverse-phase HPLC and analyze by mass spectrometry (MALDI-TOF) as in step 1.9. Expected mass: 4,080.34, observed mass: 4,084.21.</li>
	<li>Lyophilize pure dfTAT (0.71 x10-4 mmol) and resuspend in 200 &#181;l water (concentration pure dfTAT = 356.3 &#181;M).</li>
</ol>
3. Measuring dfTAT Concentration
<ol>
	<li>Resuspend an aliquot of the purified dfTAT (typically 1 &#181;l depending on the amount of peptide purified) in a 149 &#181;l of 50 mM TCEP solution.</li>
	<li>Note: In this step dfTAT is reduced to its monomer counterpart fTAT to eliminate the absorbance quenching that occurs due to the close proximity of the TMR fluorophore in dfTAT (the extinction coefficient &#949; of TMR in dfTAT is reduced in comparison to the &#949; of TMR in reduced fTAT).</li>
	<li>Allow the sample to react for approximately 20 min (analytical HPLC can be used to confirm formation of fTAT).</li>
	<li>Add all the solution to the quartz cuvette and measure the absorbance at 556 nm.</li>
	<li>Using Beer&#8217;s Law ( A= &#949;cl : &#949;= 91,500 M-1 cm-1) to calculate the concentration of fTAT, determine the concentration of dfTAT, and divide [fTAT] by two.</li>
</ol>
4. Cellular Delivery Experiments
<ol>
	<li>Seed the cells (HeLa, HDF, etc.) in a dish at a confluency of 80 - 90% (e.g., 8-well or 24-well dish). Grow cells in an appropriate medium (e.g., DMEM supplemented with 10% FBS and Pen/Strep) until 80 - 90% confluency in a 37 &#176;C humidified atmosphere containing 5% CO2.</li>
	<li>Wash the cells three times (3X) with PBS (by adding 200 &#181;l of PBS and then removing it three times).</li>
	<li>Make a 5 &#181;M working concentration of dfTAT by diluting a stock of dfTAT (in water) in nrL-15 media (for an 8 well dish the total volume should be 200 &#181;l). A concentration of 5 &#181;M dfTAT leads to efficient delivery (high level of cytosolic delivery in more than 90% of cells present in a dish) in most cell types tested to date (Figure 3). However, lower or higher concentrations might be more adequate for cell types with a high or low propensity for penetration. 
	NOTE ABOUT MEDIA: nrL-15 is used for the delivery of dfTAT since it does not contain cysteine which could reduce the disulfide bond in dfTAT. However, our data indicate that both regular L-15 (with cysteine) and DMEM can be used as the delivery media. L-15 contains the reducing amino acid cysteine but cysteine presumably oxidizes in the media to form cystine (DMEM is formulated with cystine). dfTAT therefore remains intact in these media and delivery works at the same efficiency as that obtained with nrL15.</li>
	<li>Incubate cells with dfTAT (5 &#181;M) with or without cargo (e.g., EGFP (10 &#181;M)) and keep at 37 &#176;C for 1 hr (incubation time can be reduced but dfTAT typically requires approximately 30 to 45 min to induce endosomal leakage).</li>
	<li>Wash cells with heparin (1 mg/ml) in L-15 (3 washes are recommended) to remove dfTAT bound to the plasma membrane of cells.</li>
	<li>Incubate cells with cell-impermeable nuclear stain, e.g., Sytox Blue, Sytox Green (2 &#181;M in nrL-15), to determine whether the plasma membrane of cells is compromised (dead cells will be stained while live cells will not) according to manufacturer&#8217;s protocol.</li>
	<li>Image cells using a fluorescence microscope (100X oil immersion or 20X objective). Image dfTAT using a RFP filter (Ex = 560 &#177; 20 nm/Em = 630 &#177; 35 nm). 
	Note: Successful delivery leads to a diffuse fluorescence of dfTAT throughout the cell (assessing the delivery of protein or peptide of interest will depend on application). Staining of nucleoli by fTAT (the reduced product of dfTAT upon cytosolic entry) can be used as an indication that the fluorescence detected is intracellular. fTAT will degrade within few hours. At this point, the fluorescence of the degradation fragments will appear as punctate. This should not be confused for the punctate distribution that is also seen if dfTAT remains unsuccessfully trapped inside endosomes (this can happen if dfTAT is present at too low of a concentration).</li>
</ol>
5. Controling Concentration of MOI Delivered
<ol>
	<li>Identify the &#8220;optimal delivery&#8221; concentration required to achieve efficient cytosolic release of dfTAT in the cell type used. Perform fluorescence microscopy on cells incubated with increasing concentrations of dfTAT. The optimal dfTAT concentration is defined as the minimal concentration that leads to clear cytosolic &#8220;diffuse&#8221; TMR fluorescence in approximately 100% of cells in a culture.</li>
	<li>Vary the concentration of MOI used in co-incubation protocol while keeping the concentration of dfTAT constant (e.g., using dfTAT&#8217;s optimal delivery concentration). Changing the incubation time to less than 1 hr could also be an option to vary the amount of MOI delivered.</li>
</ol>

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The authors have nothing to disclose.

The cells used for dfTAT delivery should not be overly confluent (&#62;90% confluency) since this might affect the delivery efficiency. The cells should also be healthy: dead cells in the culture can release apoptotic fragments with which dfTAT can interact (e.g., DNA from degraded nuclei). This in turn can interfere with delivery efficiency and the quality of imaging. Cells should be washed thoroughly to remove FBS before adding dfTAT. BSA present in FBS can bind to dfTAT and this can lower the delivery efficie...

The delivery of proteins, peptides or cell-impermeable small molecules into live cells is often desirable in many biological or biotechnological applications (cellular imaging, functional assays, cellular reprogramming, etc.)1-4. Many delivery approaches have already been reported, including microinjection, electroporation, or use of carrier agents (e.g., cell-penetrating peptides such as TAT, lipids)5-7. Each technique typically has specific pros and cons that might make these approaches adequate for certain applications but not for others. Common issues involve poor delivery efficiencies and/or lack of control of how much material is delivered8,9, toxicity or deleterious physiological impact10,11, lack of temporal control, delivery in few cells but not in a whole population (e.g., microinjection)12, and complex chemical conjugation or formulation schemes11.
Recently, we have developed a novel delivery strategy that circumvents these limitations. This strategy relies on a peptide named dfTAT (dimer fluorescent TAT)13. dfTAT is derived from the well-know cell&#8211;penetrating peptide (CPP) TAT. dfTAT contains two disulfide bonded copies of TAT labeled with the fluorophore tetramethylrhodamine. Despite their similarities, TAT and dfTAT differ significantly in activity. TAT is typically internalized into cells by endocytosis. This CPP remains however mostly trapped inside endosomes (this usually lead to a punctate distribution of the peptide inside cells when examined by fluorescence microscopy). Like TAT, dfTAT is efficiently endocytosed by cells. However, dfTAT does not stay trapped inside endosomes. Instead, it mediates endosomal leakage in a manner that is extremely efficient. The endosomolytic activity of dfTAT can then be exploited to deliver macromolecules by a simple incubation assay.
The current understanding of the delivery process is as follows. dfTAT induces macropinocytosis. As a result, cells incubated with dfTAT take up soluble proteins, peptides, or small molecules (molecules of interest, MOI) present in media by fluid-phase endocytosis (see Figure 1). Interactions between dfTAT and MOI are not necessary as long as both entities traffic together within the endocytic pathway. As dfTAT reaches a certain threshold within the lumen of endocytic organelles, it expresses its endosomal leakage activity (the molecular details remain to be fully characterized). The content of the lumen of leaky organelles, and therefore the MOI, is then released into the cells. This approach is therefore very convenient as no conjugation or formulation schemes with MOI are required. Moreover, because dfTAT is not directly modifying a MOI, it should also not interfere with MOIs function once intracellular delivery is achieved. In addition, the concentration of dfTAT used delivery is independent of that of MOI in media. For instance, dfTAT concentration can be kept constant between different experiments to guarantee reproducible efficiencies in endosomal leakage. In contrast, concentration of MOI in media can be gradually changed to achieve desired levels of MOI delivered in cytosol.
The high endosomal leakage efficiency achieved with dfTAT is remarkably innocuous to many of the cells tested to date. This is a surprise because endocytic organelles are important component of the cells and one would expect that the dramatic leakage mediated by dfTAT would be accompanied by deleterious cellular responses. Yet, treated cells proliferate at the same rate as untreated cells and do not display any significant changes in their transcriptome. Moreover, delivery can be repeated within minutes with reproducible delivery efficiencies, indicating that cells can either tolerate or recover from the delivery process without losing their capacity for endocytosis or endosomal leakage. Subtle cellular responses might take place during dfTAT delivery and the molecular details of what these responses might be remain to be explored. Yet, by combining high efficiency, convenience of protocols, and lack of toxicity, this delivery approach should prove immediately useful in many cell-based applications. The protocols presented herein are aimed at making this technology accessible to the research community.

<table><tbody><tr><td>Heparin&amp;nbsp;</td><td>Sigma</td><td>CAS 9041-08-1</td><td></td></tr><tr><td>SYTOX Blue</td><td>Invitrogen</td><td>S11348</td><td></td></tr><tr><td>SYTOX Green</td><td>Invitrogen</td><td>S7020</td><td></td></tr><tr><td>nrL15 L-15 (&amp;ndash;) cysteine</td><td>Hyclone</td><td></td><td>Special order</td></tr><tr><td>Fmoc-protected Amino acids</td><td>Novabiochem</td><td></td><td></td></tr><tr><td>dfTAT</td><td></td><td></td><td>Samples will be provided upon request (contact: pellois@tamu.edu)</td></tr><tr><td>Biosafety Cabinet</td><td></td><td></td><td></td></tr><tr><td>Inverted epifluorescence microscope</td><td>Olympus</td><td>model IXB1</td><td>equipped with a heating stage maintained at 37 &amp;deg;C and with a Rolera-MGI Plus back-illuminated electron-multiplying charge-coupled device (EMCCD) camera (Qimaging).&amp;nbsp;</td></tr><tr><td>37 &amp;deg;C humidified, 5% CO&lt;sub&gt;2&lt;/sub&gt; incubator</td><td></td><td></td><td></td></tr><tr><td>Peptide synthesizer or vessel to preform manual peptide synthesis</td><td></td><td></td><td></td></tr></tbody></table>

delivery of proteins, peptides or cell-impermeable small molecules into live cells by incubation with the endosomolytic reagent dftat

Macromolecular delivery strategies typically utilize the endocytic pathway as a route of cellular entry. However, endosomal entrapment severely limits the efficiency with which macromolecules penetrate the cytosolic space of cells. Recently, we have circumvented this problem by identifying the reagent dfTAT, a disulfide bond dimer of the peptide TAT labeled with the fluorophore tetramethylrhodamine. We have generated a fluorescently labeled dimer of the prototypical cell-penetrating peptide (CPP) TAT, dfTAT, which penetrates live cells and reaches the cytosolic space of cells with a particularly high efficiency. Cytosolic delivery of dfTAT is achieved in multiple cell lines, including primary cells. Moreover, delivery does not noticeably impact cell viability, proliferation or gene expression. dfTAT can deliver small molecules, peptides, antibodies, biologically active enzymes and a transcription factor. In this report, we describe the protocols involved in dfTAT synthesis and cellular delivery. The manuscript describes how to control the amount of protein delivered to the cytosolic space of cells by varying the amount of protein administered extracellularly. Finally, the current limitations of this new technology and steps involved in validating delivery are discussed. The described protocols should be extremely useful for cell-based assays as well as for the ex vivo manipulation and reprogramming of cells.

To assess the difference between fTAT and dfTAT, HeLa cells are incubated for 1 hr with each peptide to determine the difference in their cellular localization. The internalization of the two CPP&#8217;s was assessed using fluorescence microscopy. Figure 2 shows that fTAT (20 &#181;M) localizes in a punctate distribution. This distribution is consistent with the peptide remaining entrapped inside endosomes. In contrast, the fluorescence signal of dfTAT (5 &#181;M) displays a homogenous distribution throu...

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Delivery of Proteins, Peptides or Cell-impermeable Small Molecules into Live Cells by Incubation with the Endosomolytic Reagent dfTAT

We describe how to deliver proteins and cell-impermeable small molecules into cultured mammalian cells by a simple co-incubation protocol with a reagent that causes endocytic organelles to become leaky.

Macromolecular delivery strategies typically utilize the endocytic pathway as a route of cellular entry. However, endosomal entrapment severely limits the ...

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10.2K Views.  Texas A&M University. We describe how to deliver proteins and cell-impermeable small molecules into cultured mammalian cells by a simple co-incubation protocol with a reagent that causes endocytic organelles to become leaky. 

Video: Delivery of Proteins, Peptides or Cell-impermeable Small Molecules into Live Cells by Incubation with the Endosomolytic Reagent dfTAT

Cell Squeezing as a Robust, Microfluidic Intracellular Delivery Platform

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This technology has shown potential in addressing previously challenging applications; including, delivery to primary immune cells, cell reprogramming, carbon nanotube, and quantum dot delivery. This vector-free microfluidic platform relies on mechanical disruption of the cell membrane to facilitate cytosolic delivery of the target material. Herein, we describe the detailed method of use for these microfluidic devices including, device assembly, cell preparation, and system operation. This delivery approach requires a brief optimization of device type and operating conditions for previously unreported applications. The provided instructions are generalizable to most cell types and delivery materials as this system does not require specialized buffers or chemical modification/conjugation steps. This work also provides recommendations on how to improve device performance and trouble-shoot potential issues related to clogging, low delivery efficiencies, and cell viability.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This protocol provides detailed steps on how to use the system for a broad range of applications.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This ...

Detergent-assisted Reconstitution of Recombinant Drosophila Atlastin into Liposomes for Lipid-mixing Assays

Membrane fusion is a crucial process in the eukaryotic cell. Specialized proteins are necessary to catalyze fusion. Atlastins are endoplasmic reticulum (ER) resident proteins implicated in homotypic fusion of the ER. We detail here a method for purifying a glutathione S-transferase (GST) and poly-histidine tagged Drosophila atlastin by two rounds of affinity chromatography. Studying fusion reactions in vitro requires purified fusion proteins to be inserted into a lipid bilayer. Liposomes are ideal model membranes, as lipid composition and size may be adjusted. To this end, we describe a reconstitution method by detergent removal for Drosophila atlastin into preformed liposomes. While several reconstitution methods are available, reconstitution by detergent removal has several advantages that make it suitable for atlastins and other similar proteins. The advantage of this method includes a high reconstitution yield and correct orientation of the reconstituted protein. This method can be extended to other membrane proteins and for other applications that require proteoliposomes. Additionally, we describe a FRET based lipid mixing assay of proteoliposomes used as a measurement of membrane fusion.

Detergent-assisted Reconstitution of Recombinant Drosophila Atlastin into Liposomes for Lipid-mixing Assays

Biological membrane fusion is catalyzed by specialized fusion proteins. Measuring the fusogenic properties of proteins can be achieved by lipid mixing assays. We present a method for purifying recombinant Drosophila atlastin, a protein that mediates homotypic fusion of the ER, reconstituting it to preformed liposomes, and testing for fusion capacity.

Membrane fusion is a crucial process in the eukaryotic cell. Specialized proteins are necessary to catalyze fusion. Atlastins are endoplasmic reticulum ...

Biochemistry

Electroporation of Functional Bacterial Effectors into Mammalian Cells

The study of protein interactions in the context of living cells can generate critical information about localization, dynamics, and interacting partners. This information is particularly valuable in the context of host-pathogen interactions. Many pathogen proteins function within host cells in a variety of way such as, enabling evasion of the host immune system and survival within the intracellular environment. To study these pathogen-protein host-cell interactions, several approaches are commonly used, including: in vivo infection with a strain expressing a tagged or mutant protein, or introduction of pathogen genes via transfection or transduction. Each of these approaches has advantages and disadvantages. We sought a means to directly introduce exogenous proteins into cells. Electroporation is commonly used to introduce nucleic acids into cells, but has been more rarely applied to proteins although the biophysical basis is exactly the same. A standard electroporator was used to introduce affinity-tagged bacterial effectors into mammalian cells. Human epithelial and mouse macrophage cells were cultured by traditional methods, detached, and placed in 0.4 cm gap electroporation cuvettes with an exogenous bacterial pathogen protein of interest (e.g. Salmonella Typhimurium GtgE). After electroporation (0.3 kV) and a short (4 hr) recovery period, intracellular protein was verified by fluorescently labeling the protein via its affinity tag and examining spatial and temporal distribution by confocal microscopy. The electroporated protein was also shown to be functional inside the cell and capable of correct subcellular trafficking and protein-protein interaction. While the exogenous proteins tended to accumulate on the surface of the cells, the electroporated samples had large increases in intracellular effector concentration relative to incubation alone. The protocol is simple and fast enough to be done in a parallel fashion, allowing for high-throughput characterization of pathogen proteins in host cells including subcellular targeting and function of virulence proteins.

Electroporation was used to insert purified bacterial virulence effector proteins directly into living eukaryotic cells. Protein localization was monitored by confocal immunofluorescence microscopy. This method allows for studies on trafficking, function, and protein-protein interactions using active exogenous proteins, avoiding the need for heterologous expression in eukaryotic cells.

The study of protein interactions in the context of living cells can generate critical information about localization, dynamics, and interacting partners. ...

Immunology and Infection

Engineering Cell-permeable Protein

The protein transduction technique enables the direct delivery of biologically active material into mammalian cells [for review see 1,2]. For this one can make use of the translocating ability of so-called cell penetrating peptides (CPPs), also designated as protein transduction domains (PTDs). The TAT-CPP derived from the human immunodeficiency virus type 1 (HIV-1) Tat (trans-activator of transcription) protein has been widely used. The positively charged TAT promotes cell permeability thereby overcoming the barriers of the cellular membrane by endocytosis or/and direct membrane penetration2. In combination with a nuclear localization signal (NLS) fusion proteins are able to enter the nucleus exhibiting functionality. Our video presentation demonstrates, as an exemplification for the engineering of cell-permeable proteins, the construction, production and application of a cell-permeable version of the DNA-modifying enzyme Cre.
Cre is a site-specific recombinase that is able to recognize and recombine 34 base pair loxP sites in mammalian cells in vitro and in vivo. Therefore the Cre/loxP system is widely used to conditionally induce mutations in the genome of living cells3,4. The delivery of active Cre recombinase to cells, however, represents a limitation.
We describe the pSESAME vector system, which allows a direct insertion of the gene-of-interest and provides a platform to rapidly clone different domains and tags used within the vector in a convenient and standardized manner. Rearranging of the different tags has been shown to modify the biochemical properties of the fusion proteins providing a possibility to achieve higher yield and better solubility. We demonstrate how to express and purify recombinant cell-permeant proteins in and from E. coli. The functionality of the recombinant Cre protein is finally validated in cell culture by assessing its intracellular recombinase activity.

Protein transduction enables the direct delivery of biologically active proteins into cells. In contrast to conventional methods such as DNA transfection or viral transduction this non-invasive paradigm allows highly efficient cellular manipulation in a titratable manner circumventing cellular toxicity and the risk of oncogenic transformation by permanent genetic modification.

The protein transduction technique enables the direct delivery of biologically active material into mammalian cells [for review see 1,2]. For this one can ...

Biology

Measuring Peptide Translocation into Large Unilamellar Vesicles

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1. Many of these are referred to as cell-penetrating peptides, which are frequently noted for their potential as drug delivery vectors1-3. Moreover, there is increasing interest in antimicrobial peptides that operate via non-membrane lytic mechanisms4,5, particularly those that cross bacterial membranes without causing cell lysis and kill cells by interfering with intracellular processes6,7. In fact, authors have increasingly pointed out the relationship between cell-penetrating and antimicrobial peptides1,8. A firm understanding of the process of membrane translocation and the relationship between peptide structure and its ability to translocate requires effective, reproducible assays for translocation. Several groups have proposed methods to measure translocation into large unilamellar lipid vesicles (LUVs)9-13. LUVs serve as useful models for bacterial and eukaryotic cell membranes and are frequently used in peptide fluorescent studies14,15. Here, we describe our application of the method first developed by Matsuzaki and co-workers to consider antimicrobial peptides, such as magainin and buforin II16,17. In addition to providing our protocol for this method, we also present a straightforward approach to data analysis that quantifies translocation ability using this assay. The advantages of this translocation assay compared to others are that it has the potential to provide information about the rate of membrane translocation and does not require the addition of a fluorescent label, which can alter peptide properties18, to tryptophan-containing peptides. Briefly, translocation ability into lipid vesicles is measured as a function of the Foster Resonance Energy Transfer (FRET) between native tryptophan residues and dansyl phosphatidylethanolamine when proteins are associated with the external LUV membrane (Figure 1). Cell-penetrating peptides are cleaved as they encounter uninhibited trypsin encapsulated with the LUVs, leading to disassociation from the LUV membrane and a drop in FRET signal. The drop in FRET signal observed for a translocating peptide is significantly greater than that observed for the same peptide when the LUVs contain both trypsin and trypsin inhibitor, or when a peptide that does not spontaneously cross lipid membranes is exposed to trypsin-containing LUVs. This change in fluorescence provides a direct quantification of peptide translocation over time.

This protocol details a method for the quantitative measure of peptide translocation into large unilamellar lipid vesicles. This method also provides information about the rate of membrane translocation and can be used to identify peptides that efficiently and spontaneously cross lipid bilayers.

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1.  Many of these are referred ...

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

Direct Protein Delivery to Mammalian Cells Using Cell-permeable Cys2-His2 Zinc-finger Domains

Due to their modularity and ability to be reprogrammed to recognize a wide range of DNA sequences, Cys2-His2 zinc-finger DNA-binding domains have emerged as useful tools for targeted genome engineering. Like many other DNA-binding proteins, zinc-fingers also possess the innate ability to cross cell membranes. We recently demonstrated that this intrinsic cell-permeability could be leveraged for intracellular protein delivery. Genetic fusion of zinc-finger motifs leads to efficient transport of protein and enzyme cargo into a broad range of mammalian cell types. Unlike other protein transduction technologies, delivery via zinc-finger domains does not inhibit enzyme activity and leads to high levels of cytosolic delivery. Here a detailed step-by-step protocol is presented for the implementation of zinc-finger technology for protein delivery into mammalian cells. Key steps for achieving high levels of intracellular zinc-finger-mediated delivery are highlighted and strategies for maximizing the performance of this system are discussed.

Direct Protein Delivery to Mammalian Cells Using Cell-permeable Cys2-His2 Zinc-finger Domains

Zinc-finger domains are intrinsically cell-permeable and capable of mediating protein delivery into a broad range of mammalian cell types. Here, a detailed step-by-step protocol for implementing zinc-finger technology for intracellular protein delivery is presented.

Due to their modularity and ability to be reprogrammed to recognize a wide range of DNA sequences, Cys2-His2 zinc-finger DNA-binding domains have emerged ...

Imaging FITC-dextran as a Reporter for Regulated Exocytosis

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated exocytosis is fundamental for intercellular communication and is a key mechanism for the secretion of neurotransmitters, hormones, inflammatory mediators, and other compounds, by a variety of cells. At least three distinct mechanisms are known for regulated exocytosis: full exocytosis, where a single SG fully fuses with the plasma membrane, kiss-and-run exocytosis, where a single SG transiently fuses with the plasma membrane, and compound exocytosis, where several SGs fuse with each other, prior to or after SG fusion with the plasma membrane. The type of regulated exocytosis undertaken by a cell is often dictated by the type of secretory trigger. However, in many cells, a single secretory trigger can activate multiple modes of regulated exocytosis simultaneously. Despite their abundance and importance across cell types and species, the mechanisms that determine the different modes of secretion are largely unresolved. One of the main challenges in investigating the different modes of regulated exocytosis, is the difficulty in distinguishing between them as well as exploring them separately. Here we describe the use of fluorescein isothiocyanate (FITC)-dextran as an exocytosis reporter, and live cell imaging, to differentiate between the different pathways of regulated exocytosis, focusing on compound exocytosis, based on the robustness and duration of the exocytic events.

Here we detail a method for live cell imaging of regulated exocytosis. This method utilizes FITC-dextran, which accumulates in lysosome-related organelles, as a reporter. This simple method also allows distinguishing between different modes of regulated exocytosis in cells that are difficult to manipulate genetically.

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated ...

Bead Loading Proteins and Nucleic Acids into Adherent Human Cells

Many live-cell imaging experiments use exogenous particles (e.g., peptides, antibodies, beads) to label or function within cells. However, introducing proteins into a cell across its membrane is difficult. The limited selection of current methods struggles with low efficiency, requires expensive and technically demanding equipment, or functions within narrow parameters. Here, we describe a relatively simple and cost-effective technique for loading DNA, RNA, and proteins into live human cells. Bead loading induces a temporary mechanical disruption to the cell membrane, allowing macromolecules to enter adherent, live mammalian cells. At less than 0.01 USD per experiment, bead loading is the least expensive cell loading method available. Moreover, bead loading does not substantially stress cells or impact their viability or proliferation. This manuscript describes the steps of the bead loading procedure, adaptations, variations, and technical limitations. This methodology is especially suited for live-cell imaging but provides a practical solution for other applications requiring the introduction of proteins, beads, RNA, or plasmids into living, adherent mammalian cells.

Bead loading introduces proteins, plasmids, and particles into adherent mammalian cells. This cell loading technique is inexpensive, rapid, and does not substantially affect cell health. It is best suited for live-cell imaging.

Many live-cell imaging experiments use exogenous particles (e.g., peptides, antibodies, beads) to label or function within cells. However, introducing ...

Live Fluorescence, Inverse Imaging of Cell Ruffling, and Macropinocytosis

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase nutrients and other material in cells. The dramatic extension of cell surface ruffles, their closure to form macropinosomes, and the maturation of internalized macropinosomes are key events in this pathway that can be difficult to capture using conventional confocal imaging based on tracking a bolus of fluorescent cargo. Fluorescent dextrans are commonly used experimentally as fluid phase markers for macropinosomes and for other endocytic pathways. A method the lab has adopted to optimize the imaging of dextran uptake involves using live imaging of cells bathed in high concentrations of fluorescent dextran in the medium, with the unlabeled cells appearing in relief (as black). The cell ruffles are highlighted to visualize ruffle closure, and internalized macropinosomes appear as fluorescent vacuoles in the cell interior. This method is optimal for visualizing macropinosome features and allows for easy segmentation and quantification. This paper describes dual-labeling of pathways with different sized dextrans and the co-expression of lipid probes and fluorescent membrane proteins to demark macropinosomes and other endosomes. The detection of internalized dextran at an ultrastructural level using correlative light and electron microscopy (CLEM) is also demonstrated. These cell processes can be imaged using multiple live imaging modalities, including in 3D. Taken together, these approaches optimize macropinosome imaging for many different settings and experimental systems.

This protocol demonstrates dextran imaging in live cells using continuous uptake and inverse images to optimize visualization of ruffling, macropinosome maturation, and analysis of dextran and other cell labelings.

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase ...

Delivery of Proteins, Peptides or Cell-impermeable Small Molecules into Live Cells by Incubation with the Endosomolytic Reagent dfTAT

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Chapters in this video

Cell Squeezing as a Robust, Microfluidic Intracellular Delivery Platform

Detergent-assisted Reconstitution of Recombinant Drosophila Atlastin into Liposomes for Lipid-mixing Assays

Electroporation of Functional Bacterial Effectors into Mammalian Cells

Engineering Cell-permeable Protein

Measuring Peptide Translocation into Large Unilamellar Vesicles

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Direct Protein Delivery to Mammalian Cells Using Cell-permeable Cys₂-His₂ Zinc-finger Domains

Imaging FITC-dextran as a Reporter for Regulated Exocytosis

Bead Loading Proteins and Nucleic Acids into Adherent Human Cells

Live Fluorescence, Inverse Imaging of Cell Ruffling, and Macropinocytosis