Assessing the Cellular Uptake, Endosomal Escape, and Cytosolic Entry Efficiencies of Intracellular Biologics

Abstract

Intracellular biologics are an emerging class of macromolecular drugs that are either intrinsically cell-permeable or can be effectively delivered into the cell interior to modulate the activity of previously intractable drug targets. They generally enter the mammalian cell by endocytosis mechanisms and are initially localized inside the endosomes. They subsequently escape from the endosomes (and/or lysosomes) into the cytosol with varying efficiencies. In this chapter, we provide the detailed protocol for a flow cytometry-based assay method to quantitate the overall cellular uptake, endosomal escape, and cytosolic entry efficiencies of biomolecules (e.g., peptides, proteins, and nucleic acids), by using cell-penetrating peptides as an example. The scope of applicability, strengths, and weaknesses of this assay are also discussed.

1. Introduction

Intracellular biologics represent an emerging third major drug modality, in addition to small-molecule drugs and conventional biologics. Small-molecule drugs are, by definition, small in size (molecular weight typically ≤500) and are usually membrane permeable by passive diffusion. As such, small-molecule drugs can act against targets that are either outside or inside the cell. However, because of their small size, small molecules are generally ineffective against targets devoid of well-defined binding pockets, such as the proteins involved in most protein-protein interactions [1, 2]. Conventional biologics are macromolecules, most frequently monoclonal antibodies and other recombinant proteins. Their large sizes endow them the capacity to bind essentially any target with exceptional affinity and specificity, with or without hydrophobic binding pockets. However, conventional biologics are membrane impermeable and thus limited to extracellular targets, e.g., cell-surface receptors and secreted proteins. It is estimated that the combination of small molecules and conventional biologics targets only ~20% of all disease relevant human proteins, leaving the remaining ~80% currently undruggable [2, 3]. It is clear that targeting the ~80% “undruggable” proteins will require novel modalities, including intracellular biologics [4].

We define intracellular biologics as any biologic agent that is intrinsically cell-permeable (usually by a mechanism other than passive diffusion) or can be effectively delivered into an intracellular space (e.g., the cytosol) to modulate the activity of an intracellular target (e.g., proteins, mRNAs, or chromosomal DNA). Notable examples of intracellular biologics include peptides, proteins, antisense oligonucleotides, siRNAs, mRNAs, and gene editing systems (e.g., CRISPR-Cas9) [4]. Among these molecules, cell-penetrating peptides [5], stapled peptides [6], and antisense oligonucleotides [7] have demonstrated intrinsic cell permeability, as they can reach the cell interior at therapeutically relevant levels without the help of any delivery agents, whereas proteins [8], mRNAs [9], and gene editing systems [10] usually require a separate delivery vehicle to enter the cell. These modalities, either by themselves or when conjugated/complexed with delivery vectors, generally enter the mammalian cell through one or more of the endocytosis mechanisms and are initially accumulated inside the endosomes and lysosomes [11]. How they escape from the endosomal/lysosomal pathway into the cytosol is largely unknown, except for cell-penetrating peptides, which exit the endosome by inducing budding and collapse of small vesicles from the endosomal membrane [12]. The efficiency of endosomal escape varies greatly among the different modalities or even within the same modality; while nearly quantitative endosomal release has been reported for some of the cell-penetrating peptides/molecules [13, 14], the endosomal escape efficiency of antisense oligonucleotides and siRNAs was thought to be <0.01% [14, 15]. Thus, for any intracellular biologic, a key parameter is its cytosolic entry efficiency, defined as the ratio of cytosolic over extracellular concentration (where 100% represents equal concentration in the two compartments). For modalities that enter the cell by endocytosis followed by endosomal escape, it is also important to determine their total cellular uptake efficiency and the efficiency of endosomal escape.

Determination of the total cellular uptake efficiency is relatively straightforward. One of the most commonly used methods involves covalent labeling of a biologic agent of interest (e.g., a peptide) with a fluorescent dye [e.g., fluorescein (Fl) or rhodamine (Rho)] and analyzing cells treated with the dye-labeled biologic by flow cytometry. However, this method does not differentiate molecules that have successfully reached the cytosol from those still entrapped inside the endosomal/lysosomal compartments. In recent years, several innovative methods have been developed to specifically quantify the cytosolic entry efficiency of biologic agents and were expertly reviewed [12, 16–18]. In this chapter, we provide a detailed protocol for a modified flow cytometry assay previously developed in our laboratory, which is operationally simple and yet provides quantitative assessment of the efficiencies for the total cellular uptake, endosomal escape, and cytosolic entry of a biologic agent [19]. This assay method employs a pair of fluorescent dyes, one of which is a pH sensitive fluorophore, naphthofluorescein (NF; pKa = 7.8), while the other is insensitive to pH, e.g., tetramethylrhodamine (TMR) (Figure 1). A biologic agent of interest is separately labeled with TMR and NF and cells treated with the dye-labeled agent are subjected to flow cytometry analysis as usual. The mean fluorescence intensity (MFI) value of cells treated with the TMR-labeled agent provides a quantitative assessment of the total cellular uptake, i.e., agent molecules inside all cellular compartments (including the cytosol, the nucleus, and endosomes and lysosomes). The MFI values of cells treated with the NF-labeled agent reflects the number of agent molecules that have successful reached the cytosol and beyond (e.g., the nucleus), because NF is intensely fluorescent inside the neutral environments of cytosol and nucleus (pH 7.4), but essentially non-fluorescent inside the acidic environments of the endosome (pH 5.5–6.5) and lysosome (pH 4.5–5.5). The MFI^NF/MFI^TMR ratio provides a quantitative assessment of the endosomal escape efficiency. Over the past few years, we have applied this assay method to compare the cytosolic entry and endosomal escape efficiencies of a large number of CPPs as well as CPPs-cargo conjugates to gain valuable insights into the molecular mechanism and structure-activity relationships of CPPs [13, 20–23]. The scope, advantages, and limitations of this assay are also discussed.

Figure 1.

Scheme illustrating the flow cytometry-based assay method. The fluorescence yield of a biomolecule labeled with a pH-insensitive dye (e.g., TMR) remains constant as the molecule travels through the endosomal/lysosomal pathway, whereas that of NF-labeled molecule greatly increases upon escaping from the endosome/lysosome (pH 4.5–6.5) into the cytosol (pH 7.4).

2. Materials

2.1. Materials for solid phase peptide synthesis

1. Rink Amide Resin LS (100–200 mesh, 0.2 mmol/g)

2. Solvents: N, N-dimethylformamide (DMF), dichloromethane (DCM)

3. Amino acids residues: 4 eq. of the desired N-Fmoc protected amino acids dissolved in DMF

4. Coupling reagents: 2-(7-aza-1H-benzotriazole-1-yl)-1,1,3,3- tetramethyluronium hexafluorophosphate (HATU), N-hydroxybenzotriazole (HOBt), and diisopropylethylamine (DIPEA)

5. Fmoc deprotection solution: 20% (v/v) piperidine in DMF

6. Synthesis vessel: Pierce unpacked chromatography columns

7. Methyltrityl (Mtt) deprotection solution: 2:1:97 (v/v/v) trifluoroacetic acid (TFA)/Triisopropylsilane (TIPS)/DCM

8. Allyl deprotection reagents: tetrakis(triphenylphosphine)palladium [Pd(PPh₃)₄], phenylsilane

9. Peptide cyclization reagents: (benzotriazol-1-yloxy)tripyrrolidinophosphonium hexafluorophosphate (PyBOP), HOBt and DIPEA, dissolved in 2:1 DMF/DCM (v/v).

10. Cleavage cocktail: 2.5:2.5:2.5:92.5 (v/v) water/1,3-dimethoxybenzene (DME)/TIPS/TFA

11. Inert gas (e.g. nitrogen or argon)

12. Diethyl ether, chilled to −20 °C

13. Reversed-phase HPLC: 0.05% (v/v) TFA in double distilled water, 0.05% (v/v) TFA in acetonitrile (ACN), C18 column.

2.2. Fluorescent labeling reagents

5(6)-Carboxytetramethylrhodamine-N-succinimidyl ester [5(6)-TMR-NHS], 5(6)- carboxyfluorescein-N-succinimidyl ester [5(6)-FAM-NHS], and 5(6)-carboxynaphthofluorescein-N-succinimidyl ester [5(6)-NF-NHS].

2.3. Flow cytometry instrumentation

BD FACS LSR II or Aria III flow cytometer using PE channel and 561-nm laser for excitation of peptides labeled with TMR, or APC channel and 633-nm laser for excitation of NF. FlowJo software (Tree Star) for data analysis.

3. Methods

3.1. Solid phase synthesis of cell penetrating peptides

The protocol below describes the detailed synthesis of fluorescently labeled CPP12, cyclo(FfΦRrRrQ) (where Φ is L-2-naphthylalanine (Nal), f is D-phenylalanine, and r is D-arginine), a cyclic CPP of high activity in cytosolic delivery [20]. All experiments are performed at ambient temperature unless otherwise stated. This protocol may be adapted to synthesize other cyclic or linear CPPs and assumes the user have access to basic organic laboratory equipment.

1. Transfer 100 mg of Rink amide resin LS (100–200 mesh, 0.2 mmol/g) into a 5-mL peptide synthesis vessel and add 3 mL of DMF. Allow to mix on a rotary wheel for 15–20 min (see Note 1).

2. Drain the DMF under vacuum and wash with DMF (3 mL).

3. Add 4 eq. of Fmoc-Lys(Mtt)-OH, 4 eq. of HATU, 4 eq. HOBt and 8 eq. of DIPEA in 2 mL of DMF to the resin and allow to mix for 1 h. (see Note 2)

4. Drain the reaction solution and wash the resin with DMF (3 mL).

5. Add 3 mL of deprotection solution - 20% (v/v) piperidine in DMF - and incubate for 10 min on the rotary wheel. Drain the piperidine solution under vacuum, then repeat the deprotection step once.

6. Drain and wash exhaustively with DMF, DCM and DMF (3 mL each) (see Note 3).

7. Repeat steps 3–6 to couple the remaining amino acids to obtain a resin-bound linear peptide Fmoc-Phe-phe-Nal-Arg(Pbf)-arg(Pbf)-Arg(Pbf)-arg(Pbf)-Glu-OAll-miniPEG-Lys(Mtt)-NH₂. In case of synthesizing a linear peptide, proceed directly to step 16 for on-bead labeling or to step 21 for solution phase labeling. For cyclic peptide synthesis, continue from step 8.

8. With the N-terminal Fmoc group still on, wash the resin with dry DCM (3 × 3 mL). Add a solution of 0.3 eq. of Pd(PPh₃)₄ and 10 eq. of PhSiH₃ in 2 mL of dry DCM to the resin and allow to mix for 15 min. Drain and repeat this step twice (see Note 4).

9. Drain and wash the resin exhaustively with DMF, DCM, DMF (3 mL each).

10. Add 3 mL of 0.5 M sodium dimethyldithiocarbamate (SDDC) in DMF to the resin and allow to incubate for 10 min. Drain and repeat this step once.

11. Wash the resin with DMF, DCM and DFM (3 mL each).

12. Repeat steps 5 and 6 to remove the N-terminal Fmoc group.

13. Add 3 mL of 1 M HOBt in DMF to the resin and incubate the mixture for 15 min. Drain the solution and repeat this step once.

14. Add 5 eq. PyBOP, 5 eq. HOBt and 10 eq. DIPEA in 3 mL of 2:1 (v/v) DMF/DCM to the resin and allow to incubate for 1.5 h. Drain and repeat this step once.

15. Drain the solution and wash the resin exhaustively with DMF and DCM (3 × 3 mL each). Ensure that the last washing step is carried out with DCM. For solution-phase peptide labeling, proceed directly to step 21.

16. For on-bead peptide labeling, remove the Mtt protecting group from the C-terminal lysine side chain by adding 3 mL of 2:1:97 (v/v) TFA/TIPS/DCM to the resin and allow the contents to mix for 5 min. Drain the solution and repeat this step five times (see Note 5).

17. Drain the solution and wash the resin with exhaustively with DCM and DMF (3 × 3 mL each).

18. Add 3 mL of 10% DIPEA in DMF to the resin and allow the contents to mix for 10 min.

19. Drain the solution and incubate the resin with 5 eq. of succinimidyl ester of the desired fluorescent dye and 5 eq. of DIPEA in DMF (3 mL) for ≥2 h in the dark.

20. Drain the solution and wash the resin with DMF (3 × 3 mL) and then DCM (3 × 3 mL).

21. Add 5 mL of 2.5:2.5:2.5:92.5 (v/v) H₂O/BME/TIPS/TFA to the resin and allow the contents to mix for 3 h to deprotect the side chains and release the peptide from the resin.

22. Drain the solution into a 15-mL Falcon tube and concentrate it by gently blowing an inert gas (e.g., argon or nitrogen) over the surface to obtain a semi-solid (see Note 6).

23. Add 10 mL of chilled diethyl ether to the concentrated solution to precipitate the peptide. Vortex briefly, and centrifuge at 6400 rpm for 5 min.

24. Carefully remove the supernatant with a Pasteur pipette.

25. Repeat steps 23 and 24 twice.

26. Dissolve ~5 mg of the crude peptide in a minimum volume of DMSO or DMF (20 µL) and dilute it into 400 µL of 3:1 (v/v) H₂O/ACN mixture. Centrifuge the mixture in a microcentrifuge at 15,000 rpm for 5 min. Crude peptide in the clear supernatant is purified by reversed-phase HPLC equipped with a C18 column on a linear gradient of 10 – 50% ACN over 40 min.

27. Collect HPLC fractions and check the authenticity of the peptide in each fraction by MALDI-TOF mass spectrometry. Combine the desired fractions and lyophilize the solution to obtain the pure peptide as a solid. Check the purity of the product by analytical reversed-phase HPLC to ensure ≥95% homogeneity.

3.2. Solution-phase labeling of peptides with NF and TMR

1. Weigh ~2 mg of purified peptide into a microcentrifuge tube.

2. Dissolve the peptide in a minimal volume of DMF, adjust the pH to ~8 by adding ~20 µL of 0.2 M NaHCO₃ solution.

3. Add a solution of 1.5 eq of the fluorescent dye (NF-NHS or TMR-NHS) in DMF (20 µL) to the peptide solution (see Note 7).

4. Wrap the tube in aluminum foil and allow to mix at room temperature for 2 h.

5. Purify the labeled peptide by reversed-phase HPLC equipped with a semi-preparative C18 column and elution with a linear gradient of 10 – 50% ACN over 40 min.

6. Collect HPLC fractions and check the authenticity of the peptide in each fraction by MALDI-TOF mass spectrometry. Combine the desired fractions and lyophilize the solution to obtain the pure peptide as a solid. Check the purity of the product by analytical reversed-phase HPLC to ensure ≥95% homogeneity.

3.3. Flow cytometry and data analysis

3.3.1. Flow cytometry

1. Culture HeLa cells as instructed by the supplier (see Note 8)

2. Seed 1 mL of HeLa cells (15 × 10⁴ cells/mL) into a 12-well plate and culture at 37 °C in the presence of 5% CO₂ for 24 h. (see Note 9)

3. Carefully remove the growth media and wash cells with 1 mL of warm Dulbecco’s phosphate buffer saline (DPBS). (see Note 10)

4. Add 1 mL of 5 µM TMR- or NF-labeled peptide in DMEM (with 1% FBS and 1% streptomycin/penicillin) to each well and incubate at 37 °C supplemented with 5% CO₂ for 2 h (see Notes 11-14).

5. Carefully remove the media, and gently wash cells with 1 mL of cold DPBS twice (see Note 10).

6. Detach cells by treating with 300 µL of 0.25% trypsin at 37 °C (supplemented with 5% CO₂) for 5 min.

7. Add 700 µL of DPBS to quench the trypsin and transfer the resuspended cells into a 15-mL Falcon tube (see Note 15).

8. Centrifuge the cells at 250 g and 4 ^oC for 5 min.

9. Carefully aspirate 900 µL of the trypsin/DPBS solution without disturbing the cell pellet.

10. Add 900 µL of fresh DPBS, gently resuspend the cells, centrifuge at 250 g and 4 ^oC for 5 min and carefully aspirate the solution, leaving the cell pellet at the bottom of the tube. Repeat this washing step once (see Note 16).

11. Resuspend the cells in 200 µL of DPBS and analyze the cells on a BD FACS LSR II or Aria III flow cytometer. For TMR-labeled peptides, use a 561-nm laser for excitation and analyze the emission fluorescence in the PE channel. For NF-labeled peptides, use a 633-nm laser for excitation and analyze the emission fluorescence in the APC channel (see Note 17).

12. Analyze the data with Flowjo software: Generate a single-parameter histogram where X is rhodamine intensity in case of TMR-labeled peptide or APC in case of NF-labeled peptides. Add properties for each histogram so that the program will estimate Mean Fluorescence intensity (MFI) as well as cell count (see Note 18).

13. Calculate the relative total cellular uptake efficiency by using the equation

    Total uptake efficiency% (relative to Tat) $=\left(\frac{{{\text{MFI}}_{\text{ID}}}^{TMR}}{{{\text{MFI}}_{\text{\_TAT}}}^{TMR}}\right)\times 100$

14. Calculate the relative cytosolic entry efficiency by using the equation

    Cytosolic entry efficiency% (relative to Tat) $=\left(\frac{{{\text{MFI}}_{\text{ID}}}^{NF}}{{{\text{MFI}}_{\text{Tat}}}^{NF}}\right)\times 100$

15. Calculate the relative endosomal escape efficiency by using the equation

$$\gamma \hspace{0.33em}\left(\text{relative}\hspace{0.33em}\text{to}\hspace{0.33em}\text{Tat}\right)\text{\%}=\left(\frac{{{\text{MFI}}_{\text{ID}}}^{NF}}{{{\text{MFI}}_{\text{ID}}}^{TMR}}\right)\text{/}\left(\frac{{{\text{MFI}}_{\text{Tat}}}^{NF}}{{{\text{MFI}}_{\text{Tat}}}^{TMR}}\right)\times 100$$

1. Calculate the absolute cytosolic entry efficiency of the biologic agent in question by using the cytosolic entry efficiency of Tat (2.0% [24]) and the equation

   Absolute cytosolic entry efficiency% = 2.0% x $\left(\frac{{{\text{MFI}}_{\text{ID}}}^{NF}}{{{\text{MFI}}_{\text{Tat}}}^{NF}}\right)$ (see Note 19).

3.3.2. Flow cytometry in acidic solution

Even after exhaustive washing, some NF-labeled biomolecules may remain tightly bound to the extracellular surface of cells, complicating the determination of cytosolic entry efficiency. The NF signal associated with any extracellular biomolecules can be readily quenched by suspending the cells in an acidic solution (e.g., pH 6.0) immediately before flow cytometry. However, prolonged incubation of cells in the acidic solution may reduce the cytosolic pH (and the fluorescence yield of cytosolic NF-labeled biomolecules) and cause cytotoxicity.

1. Perform steps 1–10 of section 3.3.1.

2. Suspend cells in 200 µL of a pH 6 buffer immediately before flow cytometry on a BD FACS LSR II or Aria III flow cytometer for NF-labeled peptides. The buffer may be prepared by the addition of 1.0 volume of 0.2 M glycine buffer (pH 3.0) into 5.0 volumes of DPBS. (see Note 20)

3. Perform steps 12–16 of section 3.3.1.

5. Discussion

Our method provides an operationally simple, high-throughput, and quantitative assay for the overall uptake, endosomal escape, and cytosolic entry efficiencies of biomolecules. We have applied this method to quantitate the cytosolic entry efficiencies of a large number of CPPs, CPP-cargo conjugates, and cell-permeable proteins [13, 19–23]. We envision that it should also be applicable to other modalities, e.g., nucleic acids and nanoparticles, provided that they can be covalently labeled with NF and a pH-insensitive dye. To the best of our knowledge, this is the only method currently available for assessing the endosomal escape efficiency of a biomolecule in a quantitative manner. Moreover, the increase of NF fluorescence is instantaneous upon cargo release from the endosome, thus allowing the cellular entry of a biomolecule to be monitored in real time [19]. However, it should be pointed out that our method offers a quantitative comparison of two or more biomolecules with respect to their relative cellular uptake, endosomal escape, and cytosolic entry efficiencies, but does not provide the absolute values of these parameters. Nevertheless, by comparing the relative cytosolic entry efficiencies of CPPs to that of Tat, whose absolute cytosolic entry efficiency (2.0%) had been determined by fluorescence correlation spectroscopy (FCS) [24], we were able to determine the absolute cytosolic entry efficiencies of a panel of CPPs [20]. NF is a relatively large, hydrophobic dye molecule; as such, our method suffers from some of the same drawbacks of any fluorescence-based method, e.g., the dye molecule may interact with the cell membranes and alter the overall cellular uptake and/or endosomal escape efficiencies of a biomolecule. Finally, the fluorescence of NF is weak but non-zero inside the acidic environments of the endosome and lysosome; for a biomolecule of very poor endosomal escape efficiency, the fluorescence contribution from the endosome/lysosome-entrapped molecules may not be negligible.

Other investigators have recently developed several innovative assays to quantitate the cytosolic entry of biomolecules [12, 16–18]. Some of the most widely used methods are briefly discussed below and compared against our method. Understanding the strengths and weaknesses of each method would allow a reader/user to choose the most appropriate method(s) to suit their individual needs. Schepartz, Rhoades, and their co-workers employed FCS to directly measure the cytosolic concentration of fluorescently labeled peptides and proteins [24]. To our knowledge, this was the first method available for determination of the absolute cytosolic concentration of a biomolecule. Unfortunately, the FCS method has relatively low throughput and requires special instrumentation, making it unavailable to most research laboratories. The cytosolic concentration as determined by FCS is rigorously true only if the biomolecule is unfirmly distributed throughout the cytosol, whereas biomolecules bound to intracellular membranes or organelles (e.g., inner leaflet of the plasma membrane, nuclear envelope, and mitochondria) would not be detected. Recently, quantitative fluorescence imaging based on photon counting was employed to measure the intracellular concentrations of antisense oligonucleotides, including those tightly bound to nuclear components [25].

Kritzer and colleagues reported a quantitative, high-throughput chloroalkane penetration assay (CAPA) [26]. Instead of a fluorescent dye, CAPA utilizes a relatively small chloroalkane tag, which is expected to have less effect on the physiochemical properties of a biomolecule. CAPA is a functional assay and therefore provides definitive evidence whether or not a biomolecule has reached the cytosol. It is high throughput and requires low sample volumes, making it an attractive option for profiling a large number of molecules. Like our method, CAPA provides the relative cytosolic entry efficiencies of different compounds, in the form of CP₅₀ values (i.e., concentration of the chloroalkane-tagged molecule at which the cellular fluorescence produced by a cell-permeable fluorescent probe is reduced by 50%), but does not reveal the absolute cytosolic concentration of a biomolecule.

Lastly, several protein complementation assays provide robust, convenient methods for assessing functional cargo delivery into the cytosol [27–31]. These methods involve splitting a protein (e.g., EGFP [27–29] and luciferase [30, 31]) into two fragments, neither of which is biologically active. One fragment (usually the larger fragment) is overexpressed inside the cytosol of target cells, whereas the small fragment (usually a short peptide) is chemically synthesized and fused to a delivery vehicle (e.g., a CPP). Successful delivery of the peptide results in reassembly of the two fragments into a functional protein, whose biological activity is readily quantified. Complementation assays provide definitive evidence for cytosolic entry (or the lack of), but is only semi-quantitative, as the final readout (e.g., EGFP fluorescence) is influenced by the proteolytic stability of the delivered peptide and the kinetics of protein reassembly. The peptidyl cargo, which is relatively large, may alter the intrinsic cellular entry efficiency of the delivery vehicle.

Figure 2.

Structure of NF-labeled cyclic CPP12.