Phase Change Dimethyldioctadecylammonium-Shelled Microdroplets as a Promising Drug Delivery System: Results on 3D Spheroids of Mammalian Tumor Cells

Summary

Decafluoropentane microdroplets developed with a shell of dimethyldioctadecylammonium bromide exhibited an exceptional colloidal stability and an actractive biointerface. DDAB-MDs proved to be efficient drug reservoirs characterized by a high affinity to plasma membranes together with enhanced uptake and antitumor activity of Doxorubicin against human triple-negative breast cancer (MDA-MB-231) 3D model.

Abstract

Significant improvement of phase-change perfluorocarbon microdroplets (MDs) in the vast theranostic scenario passes through the optimization of the MDs composition with respect to synthesis efficiency, stability, and drug delivery capability. To this aim, decafluoropentane (DFP) MDs stabilized by a shell of dimethyldioctadecylammonium bromide (DDAB) cationic surfactant were designed. A high concentration of DDAB-MDs was readily obtained within a few seconds by pulsed high-power insonation, resulting in low polydisperse 1 µm size droplets. Highly positive ζ-potential, together with a long, saturated hydrocarbon chains of the DDAB shell, are key factors to stabilize the droplet and the drug cargo therein. The high affinity of the DDAB shell with cell plasma membrane allows for localized chemotherapeutics delivery by increasing the drug concentration at the tumor cell interface and boosting the uptake. This would turn DDAB-MDs into a relevant drug delivery tool exhibiting high antitumor activity at very low drug doses.

In this work, the efficacy of such an approach is shown to dramatically improve the effect of doxorubicin against 3D spheroids of mammalian tumor cells, MDA-MB-231. The use of three-dimensional (3D) cell cultures developed in the form of multicellular tumor spheroids (i.e., densely packed cells in a spherical shape) has numerous advantages compared to 2D cell cultures: in addition to have the potential to bridge the gap between conventional in vitro studies and animal testing, it will improve the ability to perform more predictive in vitro screening assays for preclinical drug development or evaluate the potential of off-label drugs and new co-targeting strategies.

Introduction

Drug-delivery vectors capable of ensuring high antitumor efficacy and reducing side effects are primary goals while remaining a severe chemical-pharmaceutical challenge^1,2. To date, their progress is limited at first by the contrast of an insufficient in situ drug release and a critical level of nonspecific toxicity^3,4,5. In recent years, several drug delivery systems have been implemented to improve the administration of anticancer agents, including liposomes, polymeric micelles, polymersomes^6,7,8,9,10. These systems exhibit potential in increasing circulation time and selectivity of drugs, while reducing distribution and accumulation in healthy organs and tissues. Anyway, the encapsulated formulations of antineoplastic chemotherapy drugs, such as anthracyclines, led to a significantly reduced drug internalization efficiency. Recently, stimuli-responsive micron and submicron carriers such as microbubbles¹¹, microdroplets, hybrid gold nanoparticles¹², nano-hydrogels¹³, PLGA scaffolds, and mesoporous platforms¹⁴, have been gaining pharmacological interest for their high versatility in targeting and exerting tumor inhibitory effects using doxorubicin (Dox) and docetaxel. Pioneering experiments to turn these carriers into efficient anticancer soldiers for multimodal tasking (i.e., chemotherapeutic, photothermal, and gene synergistic approaches) and molecular imaging¹⁵ have paved the way for personalized theranostic nanomedicine.

In this scenario, phase-change perfluorocarbon microdroplets (MDs) have been evaluated through the key opportunity they offer to conjugate high drug cargo loading, chemical versatility of the MDs shell addressing biological barriers, colloidal stability and synthesis efficiency^11,12. As an additional asset, the echogenicity of the MDs promoted by acoustic or optical vaporization of the perfluorocarbon (PFC) core allows to gain in situ imaging and promising therapeutic efficacy. Moreover, MDs core vaporization obtained by the energy release of ionizing particle beams can be exploited for beam tracking and radiation dosimetry.

The present study is aimed to develop decafluoropentane (DFP) microdroplets stabilized by a multiple usable shell of dimethyldioctadecylammonium bromide (DDAB) cationic surfactant. DDAB shelled-MDs meet both physico-chemical and biological expectations. DFP based microdroplets have been demonstrated to be valuable phase-change contrast agents to achieve biocompatible and stable perfluorocarbon MDs¹⁶. DDAB crystalline gel saturates long-chains at physiological temperature, deeply penetrating the hydrophobic core, stabilizing the droplet and the drug cargo therein. Moreover, the high positive ζ-potential at the water interface enhances the colloidal stability of the MDs. Biological attractiveness of DDAB shell surface lies in the ability to cause the death of bacteria and fungi, at concentrations that barely affect mammalian cells, and to bind plasma membranes, negatively charged antigenic proteins, nucleotides, DNA, or nanoparticles. The above-mentioned features can be exploited to generate a remarkable immunoadjuvant, gene therapy and antitumor action within mammalian cells¹⁷.

Dox-loaded DDAB-MDs (Dox@DDAB-MDs) described herein promote the drug release against highly aggressive, invasive, and poorly differentiated triple-negative breast cancer cells. A simple and rapid protocol is described below based on high power probe insonation to obtain stable and high-density DDAB-MDs with a narrow size distribution with a high loading efficiency of Dox in a one-step formulation. Such characteristics are competitive even for other preparation methods like microfluidic devices and high shear homogenizers¹⁶.

The other major limiting issue in designing efficient drug delivery vectors is that the activity of a drug is a function of various parameters (e.g., absorption, distribution, concentrations) obtainable in an actual biological target, which cannot be considered by monolayer cell models¹⁸. For this reason, the history of the development of novel antitumor formulations is studded with in vitro studies that unfortunately have resulted to be ineffective already at the level of preclinical models in animals¹⁹.

Particularly, the need to move from cell cultures to a more complex and reliable system than in vivo and ex vivo studies is linked to the inherent limitations of pharmacological studies on 2D cultures. In this context, the in vitro 3D systems are included, such as spheroids, organoids, organ-on-chip, which simulate the morphology, activity, and physiological response of more complex structures than the 2D monolayers²⁰. In a preclinical view, 3D cell models mimicking the cellular microenvironment offer the possibility to better understand complex biology in a physiologically more pertinent frame in which traditional monolayer cultures are not effective^21,22.

After proving that DDAB-MDs can interact with the cell membrane of human breast cancer cells, favoring drug internalization and cell death at very low (nanomolar) Dox concentration, the efficacy of such methodology against 3D spheroids of mammalian tumor cells, MDA-MB-231, has been tested.

Protocol

NOTE: All the reagents and instruments are listed in the Table of Materials.

1. Fabricating and characterizing microdroplets

1. Preparing Dox-loaded DDAB-MDs
   1. Dissolve the DDAB powder in ethanol to obtain a final concentration of 10 mM and a final volume of 1 mL. Prepare 1 mL of Dox stock solution dissolving 2 mg Dox powder in ethanol.
      CAUTION: Dox is known to have acute oral toxicity, category 4 and carcinogenicity, category 1B. Use only under a fume hood with gloves and a health mask.
   2. Add 250 µL of DFP to 300 µL of DDAB solution (oil phase) in a 15 mL plastic graduated centrifuge tube.
      NOTE: The purity of DFP is 60% (GC), density 1.6 g/mL (20 °C), boiling point is 55 °C. Dox purity is 98, 0-102, 0% (HPLC). DDAB purity is ≥98% (TLC). Purity of ethanol is 97%.
   3. Gently add 2.15 mL of deionized water on the oil phase resulting in a two-phase water/oil mixture. Gently inject 10 µL of Dox solution directly into the oil phase immediately before the insonation to avoid partitioning of a significant amount of Dox into the water phase.
   4. Emulsify the biphasic mixture by probe insonation (with a 1/8 in titanium tapered microtip) using a high-intensity ultrasonic liquid processor equipment in a pulse mode (0.7 s On and 0.3 s Off) at 20 kHz, 100 W for 10 s. Immediately dilute freshly prepared MDs with ultrafiltered, deionized water (e.g., MilliQ) by a factor of 1.8.
      NOTE: The dilution factor is indicative, empirically chosen. MDs solution can be diluted moreas long as enough pellet is obtained from subsequent centrifugations.
   5. Take 1 mL of the obtained suspension and centrifuge 3x, resuspending in ultrafiltered, deionized water (e.g., MilliQ) to remove the excess of Dox and ethanol. Carry out the first centrifugation at 25 °C, 360 x g for 3 min, and the second and third one at 280 x g at the same temperature and time.
   6. After the last wash, remove the supernatant water and draw up 5 µL of the pellets and disperse them into 2 mL of the medium for cell treatment, obtaining an equivalent Dox concentration of 10 nM.Protocol steps for Dox@DDAB-MDs synthesis are shown in Figure 1.
      NOTE: Equivalent Dox concentration is defined as the amount of free Dox contained into the same volume of a suspension of Dox@DDAB-MDs.
2. Characterization of Dox-loaded DDAB-MDs
   1. Measure the size distribution using a Dynamic Light Scattering (DLS) photometer and analyze the obtained correlograms with the CONTIN algorithm to extrapolate the associated decay times. Then use the decay times to determine the distribution of the particles' diffusion coefficients (D) and, convert these in a distribution of hydrodynamic diameters (2R_H) using the Stokes-Einstein relationship R_H = k_BT/6πηD, where k_BT is the thermal energy of the system and η the solvent viscosity.
   2. Check the size and size distribution also using bright field microscopy with an image analysis software (e.g., Image J), measuring the size of at least 100 droplets per frame (for 3 or 4 frames), obtaining an average value and a standard deviation.
      NOTE: For DLS measurements dilute the MDs solution in ultrafiltered, deionized water (e.g., MilliQ), PBS and in cell culture medium to avoid backscattering to a fixed concentration of 1.5 x 10⁸ MDs /mL. For ultrafiltered, deionized water, and PBS in cell culture medium, we obtained mean size values of 1.1 ± 0.1 µm, 1.1 ± 0.25 µm, and 1.2 ± 0.2 µm, respectively. MDs recovered from the pellets, after centrifugation, do not show any size changes within the errors.
   3. Use a cell counting chamber slide for microscopy to assess MDs concentration. The chamber is composed of two different counting areas with a thickness of 0.10 mm. Place 10 µL of MDs suspension over the chamber. Count MDs inside the 0.25 x 0.25 nm² square of the chamber using an optical microscope with a 40x long distance objective and analyze with an image analysis freeware.
   4. Calculate the MDs concentration, expressed as number of MDs/mL, according to the equation:
      
   5. Check the internalized Dox by Confocal Laser Scanning Microscope (CLSM) images exploiting the Dox autofluorescence at 590 nm. Measure the Dox content using a fluorimetric assay, dispersing the particles in ultrafiltered, deionized water or PBS, centrifuging them at 25 °C, 360 x g for 3 min and then determining the amount of free drug by evaluating the fluorescence of the supernatant at 590 nm with a calibration curve (linearity range for Dox concentration: 2-20 µmol/mL, R² = 0.99).
      NOTE: Subtract the obtained value from the total concentration of Dox to obtain the encapsulated amount of Dox. Perform the experiment in triplicate. The Dox encapsulation efficiency results in 28% ± 2%, calculated by dividing the drug amount of Dox in the MDs with the total content of Dox deployed.
   6. Perform experiments of Dox release over time, by centrifuging and repeating the procedure as per step 1.2.3 to obtain the percentage of Dox released in the supernatant over to the number of encapsulated ones. After the first determination, repeat the procedure by decreasing the centrifugation speed to 280 x g to avoid MDs breaking. Plot a release curve over time. Check that the concentration of Dox released within 24 h in the supernatant reaches a maximum of 20%.
   7. Measure the ζ-potential at 37 °C using a dedicated apparatus (e.g., NanoZetaSizer) and verify that the obtained values are around 90-100 mV.

2. Fabrication of spheroids in micro-molded nonadhesive substrates

1. Casting micro-molds
   1. Place small 3D molds and 1 g of pure agarose powder in containers suitable for sterilization. Autoclave them for 30 min on a dry cycle (121 °C).
   2. In a biosafety cabinet, add 50 mL of 0.9% saline (NaCl) solution to the glass bottle containing sterilized agarose and place it into a microwave oven to boil and dissolve the powder. Let the molten agarose cool down to 60 °C and add 500 µL to each micro-mold avoiding creation of bubbles when pipetting.
      WARNING: Molten agarose can cause severe skin burns. Appropriate personal protective equipment is needed.
   3. Remove the gelled agarose by carefully flexing the mold and removing the newly formed substrate. Put the substrates in a 12-well plate and add 2 mL of fresh culture medium per well. Incubate for at least 15 min to equilibrate each substrate.
      NOTE: Prepare the culture medium in advance and pass it through a 0.22 µm filter to remove possible particles or contaminants.
2. Seeding the cells
   1. Culture MDA-MB 231 cells with complete medium (DMEM supplemented with 1% Pen/Strep, 10% FBS, and 1% L-Glu) in a cell culture incubator at 37 °C, 5% CO₂. When the cells reach about 80% confluence in a T75 flask, discard the medium, rinse with 4 mL of DPBS and add 3 mL of trypsin/EDTA. Place the flask in the incubator and wait for cells to detach. Inspect the detachment under an inverted microscope (at 20x) every 5 min.
   2. Collect the detached cells with 3 mL of DPBS with 10% of FBS in a 15 mL tube and centrifuge at 235 x g for 10 min. Discard the supernatant containing both DPBS and trypsin/EDTA and resuspend the cells pellet in 3 mL of fresh medium. Pipette 10 µL of cell suspension with 10 µL of trypanblue and count the cells using a Neubauer chamber.
      ​NOTE: When harvesting the cells, neutralize the trypsin/EDTA with DPBS containing 10% of FBS to prevent the trypsin from damaging the cells during the centrifugation step.
   3. To obtain a nominal spheroid diameter of 50 µm (~15 cells/spheroid), dilute the cell suspension to a final concentration of 3,840 cells/190 µL, resulting in 256 spheroids in the small mold. Alternatively, for a 200 µm spheroid diameter (~1,000 cells/spheroid) dilute to a final concentration of 81,000 cells/190 µL, resulting in 81 spheroids in the large mold, as per the manufacturer's instructions.
   4. Remove the culture medium from the 12-well plate and tilt the substrates to carefully remove the medium from the cell seeding chamber. Add 190 µL of cell suspension to each substrate (in a dropwise manner) and wait for cells to settle for 10 min in the tissue culture incubator.
   5. Add 2 mL of the surrounding medium per well and place the multiwell back to the incubator. Inspect for spheroid formation every 24 h. Replace with a fresh medium when needed.
3. Harvesting and processing spheroids
   1. Place a 35 mm Petri dish containing 2 mL of fresh medium in the incubator to equilibrate for 10-15 min.
   2. Remove the cell culture medium surrounding the substrate. With a sterile tweezer, remove the substrate from the well and invert it in the Petri dish. Gently tap the bottom of the substrate to make the spheroids fall by gravity.
   3. Place the Petri dish containing the spheroids back in the incubator for further processing. Protocol steps for spheroids fabrication are shown in Figure 2.

3. Spheroid treatment

1. Remove the supernatant from the mold and replace it with 200 µL of Dox@DDAB-MDs dispersion, prepared as described in step 1.1.6.
2. After 5 min, add 2 mL/well of surrounding medium to equilibrate. After the desired treatment time, proceed as per step 2.1.

4. Characterization of spheroid size and morphology

1. Check the spheroid size after the desired incubation time with a 40x objective combined with the microscope image processing software.
2. Measure the size of at least 10 spheroids to collect enough data for suitable statistics and analyze their volumes as reported in step 7.

5. Proliferation/viability assay: fluorescence microscopy with live cell staining

NOTE: Follow the instructions for spheroid fabrication until step 2.2.5.

1. Remove the supernatant from the mold and replace it with 200 µL of 4 µM calcein-AM in PBS and incubate for 3 h at room temperature in the dark. During the last 20 min of incubation, add 10 µg/mL of propidium iodide to stain the dead cells.
2. Harvest the spheroids by inverting the substrate into the Petri dish as per step 2.3.2.
3. Image the spheroids in staining solution using CLSM with 40x/60x objectives and an Ar+ laser, setting the gain and the pinhole in appropriate ways to obtain focused images.

6. Image analysis and acquisition

1. Transmission and confocal 2D images
   1. Open the confocal software (Supplementary File A) and select the objective (60x, 40x, etc). On the right panel click on Trans to select the transmission channel.
   2. Click on Live to visualize the image and search for the optimized focus. Click on Single to stop the acquisition.
   3. For the confocal images, click on Red or Green laser channels depending on the fluorescent dye used. On the pinhole section, select S-aperture on the drop-down menu and set the gain section at 6.00 B.
   4. Click on Live and set the pinhole aperture and gain for optimizing the contrast. Click on Single to stop the acquisition and save the image. Overlap the transmission and confocal image by selecting the correct channel on the top toolbar.
2. Confocal 3D images
   1. Click on Z on the right panel (Supplementary File B). Click on the red button to reset the settings.
   2. Select the Ref section, click on Live and search the median plan inside the object moving the focus. Select the Top section and move the focus up until the object is out of focus and disappears.
   3. Repeat step 6.2.2 for the bottom section moving down the focus.
   4. Set the step size to 0.75 µm. Click on Z-stack and then on Single to start the scanning. Save the images in .ics format.
3. Optical images and analysis
   1. Open the optical microscope software, capture the image with a 40x long focus or 20x objective by pressing Play on the top toolbar. Press Stop and save the image.
   2. On the top toolbar click on View, Analysis Control, Annotations and Measurements opening a panel with different options (Supplementary file C). In the Annotation and Measurements panel select Semiaxis and click on Ellipse.
   3. On the Image Search, search for the axis that better fits the object shape and press Enter on the keyboard to transfer the obtained value on the right panel table.
   4. Repeat step 6.3.3 for at least 10 objects to obtain sufficient data. Click on Export to Excel Data Sheet to export the data and save them.
4. 3D Z-Stack image visualization and volume numerical calculation
   1. Open the 3D image captured with the confocal microscopy in the optical microscope software (Supplementary file D).
   2. To see the 3D reconstruction, click on Show Volume View in the image toolbar; the object can be rotated in any direction by clicking the left mouse button. To select a portion of the volume, keep pressing ctrl + left mouse keys (Supplementary File E).
   3. Press the x button on the keyboard to take a snapshot of the 3D image and to save it.
   4. To calculate the object volume, click on Measure, 3D object measurements opening a panel (Supplementary File, Image F). Click on the panel toolbar Define 3D threshold and set the optimized threshold using also the smooth and clean filters (Supplementary File, Image G). Click on OK obtaining in the 3D object measurements panel different parameters, including the volume.
   5. Click on Export to Excel to export data and save them.

7. Spheroid data analysis

1. For the spheroids volume calculations, approximate the 3D structure with a prolate ellipsoid, estimating the major and minor axis through the 2D projection as shown in the insert of Figure 3.
2. Validate the prolate ellipsoid approximation through a comparison between the volume calculated numerically as shown in step 6.4 and the prolate ellipsoid volume formula with b>a=c where a, b, and c are the ellipsoid axis).
3. Calculate the mean volume of at least 10 spheroids and the respective standard deviations. If the volume distribution of spheroid is normal, apply the Dixon test to identify and reject the outlier value. Thereafter, calculate the volume ratio between the treated and samples together with the error propagation as reported in Figure 6A.

Results

Dox@DDAB-MDs were developed according to protocol (Section 1) as schematically described in Figure 1. The obtained MDs are made of a monolayer of DDAB encapsulating the DFP core (Figure 1A). The cationic charge of DDAB and the sonication procedure avoid the formation of DDAB multilamellar layers stacked at the DFP and water interface²³.

The CLSM micrograph (Figure 1B

Discussion

To improve the efficacy of anthracyclines as antitumor drugs, this work presents the formation of DDAB shelled PFC droplets encapsulating the chemotherapeutic drug doxorubicin (Dox) and the effect of such formulation interacting with the high aggressive triple-negative breast cancer cells, MDA-MB-231.

Building up of DOX@DDAB-MDs
Dox loaded MDs have been formulated by the insonation method with an extremely fast, well reproducible, user-friendly, and efficient protocol. T...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
µ-Petri dish	Ibidi	81156	35mm high, IbiTreat
1,1,1,2,3,4,4,5,5,5-Decafluoropentane	Sigma-Aldrich	138495-42-8	b.p. 55°C
12-well culture plate	Corning
15 ml centrifuge tube	Falcon	89039-664
3D-Petri dishes 12:256	Microtissues (Sigma-Aldrich)	Z764000-6EA	Small
3D-Petri dishes 12:81	Microtissues (Sigma-Aldrich)	Z764019-6EA	Large
5%CO2 culture incubator, 37°C	Thermo Scienific		HERAcell 150i
50 ml centrifuge tube	Falcon	352070
Biological safety cabinet, II level
Calcein	Sigma-Aldrich
Calcein-AM	Sigma-Aldrich	148504-34-1	4mM stock solution in DMSO
cam sCMOS Andor Zyla 4.2	Andor Instruments
Centrifuge Hettich Universal 320R	Hettich Lab. Technology
DAPI	SIgma-Aldrich
Dimethyldioctadecylammonium bromide powder	Sigma-Aldrich	3700-67-2
DMEM (Dulbecco's Modified Eagle Medium)	Corning	15-013-CV
Doxorubicin hydrochloride	Sigma-Aldrich	25316-40-9
DPBS (Dulbecco's Modified PBS)	Corning	21-030-CV	pH 7,4
Ethanol 70%	Sigma-Aldrich
EZ-C1 digital ecliplse	Nikon Instruments		Silver version 3.91
Fetal Bovine Serum (FBS)	Corning	35-079-CV
Goniometer BI-200SM	Brookhaven Instruments Corporations
Laser Ar+	Spectra Physics
Laser He-Ne	Melles-Griot
L-Glutammine	Corning	25-005-CI
Mcroscope Nikon Eclipse Ti	Nikon Instruments
MDA-MB 231 cell line	ATCC
Microsoft Excel	Microsoft
Microplates reader Spark	Tecan group
NanoZetaSizer ZS	Malvern Instruments LTD
Neubauer improved chamber		718605
NIS Elements software	Nikon Instruments		AR 4.30
Pen/Strepto	Corning	30-002-CI
Photocorrelator BI-9000 AT	Brookhaven Instruments Corporations	62927-1
Photometer HC120	Brookhaven Instruments Corporations	N° 1275
Pipettors and tips, various size	Gilson
Propidium Iodide	SIgma-Aldrich
Serological pipets, various size	Corning
Solid-state laser	Suwtech Laser	N° 22368
T25 Flasks	Sarstedt	83.3910.002
T75 Flasks	Sarstedt	83.3911.002
Trypsin/EDTA 0.05%	EuroClone	ECB3052D
Vibra-Cell VCX-400	Sonics & Materials, inc
Water bath			37°C