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. 2022 Aug 15;38(34):10351–10361. doi: 10.1021/acs.langmuir.1c03388

Deciphering the Emulsification Process to Create an Albumin-Perfluorocarbon-(o/w) Nanoemulsion with High Shelf Life and Bioresistivity

Johannes Jaegers †,‡,*, Sven Haferkamp §, Oliver Arnolds ∥, Daniel Moog ⊥, Anna Wrobeln †, Fabian Nocke †, Miriam Cantore †, Stefanie Pütz ∥, Anne Hartwig #, Rico Franzkoch ∇, Olympia Ekaterini Psathaki ∇, Holger Jastrow ○,◆, Carsten Schauerte §, Raphael Stoll ∥, Michael Kirsch , Katja Bettina Ferenz †,⋈,*
PMCID: PMC9435530  PMID: 35969658

Abstract

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This work aimed at the development of a stable albumin-perfluorocarbon (o/w) emulsion as an artificial oxygen carrier suitable for clinical application. So far, albumin-perfluorocarbon-(o/w) emulsions have been successfully applied in preclinical trials. Cross-linking a variety of different physical and chemical methods for the characterization of an albumin-perfluorocarbon (PFC)-(o/w) emulsion was necessary to gain a deep understanding of its specific emulsification processes during high-pressure homogenization. High-pressure homogenization is simple but incorporates complex physical reactions, with many factors influencing the formation of PFC droplets and their coating. This work describes and interprets the impact of albumin concentration, homogenization pressure, and repeated microfluidizer passages on PFC-droplet formation; its influence on storage stability; and the overcoming of obstacles in preparing stable nanoemulsions. The applied methods comprise dynamic light scattering, static light scattering, cryo- and non-cryo-scanning and transmission electron microscopies, nuclear magnetic resonance spectroscopy, light microscopy, amperometric oxygen measurements, and biochemical methods. The use of this wide range of methods provided a sufficiently comprehensive picture of this polydisperse emulsion. Optimization of PFC-droplet formation by means of temperature and pressure gradients results in an emulsion with improved storage stability (tested up to 5 months) that possibly qualifies for clinical applications. Adaptations in the manufacturing process strikingly changed the physical properties of the emulsion but did not affect its oxygen capacity.

Introduction

Substitution of hydrogen atoms in hydrocarbons either completely or partially with fluorine atoms tremendously changes their physical and chemical properties.1 These perfluorocarbons (PFCs) offer applicability as artificial oxygen carriers for blood substitution due to their gained oxygen capacity.2−4 In the case of a complete exchange, e.g., in perfluorodecalin (PFD), the ability to perform van der Waals interactions or accept any kind of hydrogen bond is diminished, forming rare species of hydrophobic and lipophobic substances.5 Therefore, emulsification is necessary for both avoiding life-threatening edema inside the body and confecting a physiological carrier solution.6 The clinical context asks for biocompatibility at all steps of the manufacturing process. Albumin as a physiological compartment of the human blood combines the advantage of biocompatibility with the ability to function as an emulsifier. On the downside, albumin is ponderous and, compared to other emulsifiers, a temperature-sensitive molecule.

The following work is dichotomous. (i) We performed a detailed and multifaceted characterization of the emulsification process, and the droplet appearance (shell thickness, droplet size distribution, stability) of an albumin-perfluorocarbon-(o/w) emulsion proved to be highly potent in supplying oxygen in ex vivo perfused kidneys for organ preservation.7 (ii) The gathered information was used to exploit the physical properties of albumin to create a stable O/W perfluorocarbon nanoemulsion with high shelf life and bioresistivity.

Emulsification is a thermodynamic process in which one liquid phase is fragmented inside another liquid phase forming small droplets. The process can be described by the Gibbs–Helmholtz equation, which is mainly influenced by the energy input and the interfacial tension.8 Of course, the stability of the formed emulsion essentially depends on kinetic stabilization.9 High-pressure homogenization (HPH) is a well-established technique to create stable nanoemulsions in the food, pharmaceutical, and cosmetic industries.10 In this physical process, pressures up to 30,000 PSI are used to accelerate a pre-emulsion, which is subsequently directed through microchannels with a diameter of 75–100 μm. Inside these microchannels, the droplets are compressed by pressure and disrupted either by collision with the channel walls/each other or by shear forces, plus eventually by decompression after leaving the channels (Scheme S1).11,12

During HPH, two forces counteract. On the one hand, the collision and shear forces transform bigger droplets into smaller ones. Noticeably, the newly generated smaller droplets fuse back again to bigger droplets (recoalescence).13−15 On the other hand, recoalescence emerges through holes in the interfacial surfactant shell that appear when smaller droplets are pinched off, leaving an area that is insufficiently kinetically stabilized.16 The closing of these holes, and therefore, the prevention of recoalescence, depends on the ability to gain high surface coverage of the surfactant in a short time by either Marangoni convection alongside the interface or diffusivity through the surrounding medium.17,18

However, PFC emulsions that entered clinical trials and applications were produced using either Pluronic F-68, egg yolk lecithin, or albumin as surfactants.19,20 Albumin as a surfactant seems advantageous since it allows for the creation of a biocompatible emulsion with anti-inflammatory properties.21−25 Further, compared to Pluronic F-68 and egg yolk lecithin, albumin has a greater molecular size and a more complex structure.19,26 Albumin contains three domains with 17 intramolecular disulfide bridges, rendering it extremely stable toward long-lasting high pressure and moderate temperature.27 The surface offers both charged and hydrophobic areas and pockets for the transport of ions and lipophilic hormones, lipids, and xenobiotics.28,29 These properties cause differences alongside the surface regarding its adhesive properties during HPH and require further investigation of its behavior during HPH.30,31

The formation of stable planar interfacial albumin layers is divided into three regimes.32

  • (i)

    Diffusivity-limited adhesion of albumin on the surface with random orientation. This takes minutes to hours and does not lead to a substantial reduction of surface tension.33 In addition, albumin consists of three subdomains, of which one domain is mostly hydrophilic, whereas the other two domains are mostly hydrophobic. This anisotropy of albumin allows for a quite unique ability to either adsorb reversibly, i.e., with the hydrophilic domain facing the hydrophobic droplet surface, or irreversible, where the hydrophobic domains touch the hydrophobic surface.32,34 The transition from reversible to irreversible adsorption can only be accomplished through desorption and readsorption.35

  • (ii)

    Further adhesion of albumin leads to a substantial reduction of the surface tension and, therefore, to increased stability of emulsions.

  • (iii)

    Generation of multilayers and a further increase of the emulsions’ kinetic stability by lowering surface tension to a minimum.

The characterization of PFC emulsions is difficult, and commonly, studies are limited to the size distribution determined via dynamic light scattering (DLS), static light scattering (SLS), or cryo-TEM (cTEM). To manipulate the process, a deeper understanding of the process of droplet formation is needed. Here, we describe the multifaceted analysis of droplet formation during HPH of an albumin-perfluorocarbon-(o/w) emulsion and a protocol to manufacture a stable nanoemulsion for use as an oxygen carrier.

Experimental Section

Ethical Approval

The Ethics Council of the Essen University Hospital authorized the collection of blood for the purpose of experimental studies (Docket number 19-8600-BO).

Emulsification via Microfluidics

For emulsification, 4 mL of PFD (F2, Lancashire, United Kingdom) was added to 20 mL of aqueous 5% BSA solution (5 mg/mL, 0.75 mM) (Applichem, Darmstadt, Germany). In the case of oxygen transfer, determination Ringer salts (Sigma Aldrich, Darmstadt, Germany) were added to avoid hemolysis. The mixture was predispersed for 1 min with the UltraTurrax dispersion tool (IKA, Staufen, Germany). Afterward, the predispersed mixture was further emulsified through a microfluidizer (Microfluidics, Westwood, MA) at, respectively, 10,000, 20,000, and 30,000 PSI either on ice or at room temperature.

Emulsification using a temperature and pressure gradient was performed under temperature control using a thermometer placed in the reservoir of the microfluidizer. The inner friction of the predispersed emulsion resulted in a 2 °C increase per passage starting at 4 °C and 20,000 PSI. After reaching exactly 40 °C, the EP was increased to 30,000 PSI and kept at this level until reaching a temperature of 50 °C. We detected that a temperature above 50 °C led to denaturation, thereby destructing the emulsion. Important for the durability of the emulsion was the rapid temperature decrease to 0–4 °C after the process.

Dynamic Light Scattering (DLS)

For droplet size distribution (DSD) determination via DLS measurements (Nano-flex, Droplet Metrix GmbH, Inning, Germany), the droplet’s refractive index was determined in advance. This was done by measuring a concentration series of the emulsion ranging between 2.5% (w/w) and 20% (w/w) droplets at 20 °C with a multiwavelength attenuated-total-reflection refractometer (DRS-λ, Schmidt + Haensch, Berlin, Germany) at different wavelengths between 403 and 938 nm. The measured refractive indices were extrapolated to the refractive index of the droplets using the Newton equation and afterward interpolated to the relevant wavelength of 780 nm by a Cauchy formula, resulting in a refractive index of 1.328. This refractive index could not be used for the DLS-based DSD because it does not differ sufficiently from the refractive index of the surrounding water of 1.33336 (data shown in Figure S1). The measurements of dynamic and static light scattering were carried out with a refractive index of 1.3. They were performed undiluted at 20 °C and in the intensity mode. The standard implementation parameters included the following: setzero time, 300 s; measurement duration, 300 s; automatic measurement repetitions, three; refractive index (droplets), 1.30; shape of the droplet, spherical; refractive index (medium), 1.35; viscosity (20 °C), 2.232; and viscosity (25 °C), 1.974.

Static Light Scattering (SLS)

For DSD determination, SLS measurements were performed (Beckman Coulter LS 13320 with Universal Liquid Module (ULM)/Brea/CA) incorporating polarization intensity differential scattering (PIDS) technology to provide a dynamic range of 0.017–20 μm. We tested various optical models (refractive indices) for undiluted, fully emulsified droplets, and the best “residual” value was obtained with a refractive index of 1.602. (This value was not applied to the DLS measurements because it did not improve the measurements (Figure S4).) This correlates with the refractive index of BSA, giving us reason to assume that BSA dominates the optical properties of the particles in SLS measurements.37 The standard implementation parameters included the following: pump speed, 35%; obscuration, 40–50%; runtime, 60 s; number of runs, 10; laser wavelength, 780 nm; PIDS wavelengths—450, 600, and 900 nm in an angle range of 0–146 °; refractive index (medium), 1.329; and refractive index undiluted droplets: 1.602.

Determination of the Electrokinetic Sonic Amplitude (ESA)

The method for determining ESA used in this study is an electroacoustic measurement technique to measure the electrostatic droplet charge. The ESA signal is the detected amplitude of the oscillating droplets and, thus, a raw data signal, which is picked up without optical parameter input of the droplets. Due to the proportionality with the ζ-potential, the ESA raw data signal can be used to reliably characterize the dispersion if detailed data of the dispersed phase are not available or are incomplete. Original formulations can be measured without dilution steps, and the stirring of the emulsion that prevents sedimentation has no disturbing influence on the quality of the measurement. For this reason, the ESA method described in this report is suitable for the characterization of the highly concentrated emulsion.38 Still, for the ESA determination, it was necessary to remove excess BSA, meaning all BSA that was not bound to the surface of the droplets. This was done by three-time centrifugation (first: 4 °C, 1000g, overnight; second and third: 4 °C, 1000g, 3 h) and resuspension in purified water. The ESA measurement was performed as previously described by Renner et al.39 In brief, 60 mL of the emulsion was transferred into a special measuring chamber that allowed for simultaneous probing of pH, conductivity, temperature, and ESA signal. The sample was stirred continuously at room temperature while 0.1 M NaOH was added dropwise during the measurement. To detect changes in the electrostatic stability of the droplets in dependence on the pH, an oscillating voltage was applied that forces charged droplets to oscillate dependent on the applied electric field. The amplitude of the sound waves generated by the droplets is the ESA signal and is proportional to the surface charge, the dynamic mobility, and, therefore, the ζ-potential of the droplet. ESA determination was performed by the commission and consulting company for powder and dispersion analysis “Pulveranalyse”; Dipl.-Ing. Daniel Moog, Euskirchen, Germany.

Cryo-Scanning Electron Microscopy (cSEM)

Excess BSA was removed by centrifugation as described above. cSEM was performed by “Labor Dr. Schäffner GmbH”, Solingen, Germany. The samples were jet frozen with a rate of 10,000 K/s and plunged in liquid propane at −196 °C. The cryo-fixated sample was transferred into a freeze corrosion unit where the top layer was ripped off to expose the fixated structures. Afterward, the temperature was increased up to −100 °C, and the surrounding pressure was decreased to 10–4 Pa to sublimate excess water. The exposed surface was sputtered with a layer of Pt/C with a thickness of 1.5 nm and further stabilized with a layer of 5 nm graphite. The surface imprint was then transferred onto a grid and immediately analyzed using a field emission electron microscope (Toshiba, Japan).

Cryo-TEM- (cTEM)

For cryo-transmission electron microscopy, 4 μL of each of the two following samples (i) single pass, 20,000 PSI, 4 °C and (ii) temperature gradient (4–50 °C) and pressure gradient (20,000–30,000 PSI), were placed on glow discharged holey copper grids (Quantifoil R2/2 200 Mesh). The grids were blotted for 3 s at 10 °C and 80% humidity and frozen in liquid ethane using a Leica GP2 plunge freezer (Leica, Germany). The grids were transferred into a JEOL JEM2100Plus transmission electron microscope (JEOL, Japan) operating at 200 kV. Digital micrographs were recorded manually with a XAROSA CMOS 20 Megapixel Camera (Emsis GmbH, Germany)

cTEM-Based Droplet Size Determination

The size of the droplets was determined using the Fiji/ImageJ freeware software. The size of all droplets observed in four images at a magnification of, respectively, 2000× and 3000× was determined using the straight-line tool, and the scale bar was used for calibration.

Transmission Electron Microscopy (TEM)

For TEM analysis, the cells were fixated in 4% paraformaldehyde + 2.5% glutaraldehyde in PHEM buffer (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, 2 mM MgCl2, 4% glutaraldehyde, 1% formaldehyde) and incubated for 3 h at room temperature.40 In a laboratory microwave (Pelco BioWave Pro+, Ted Pella, Redding), they were washed three times in PHEM buffer, incubated using 1% OsO4 in PHEM buffer, washed in water, contrasted with 1% uranyl acetate in water, washed in water and via an ethanol chain, and acetone-embedded in EponTM. After overnight polymerization, 55 nm-thick sections were cut on an EM UC7 ultramicrotome (Leica, Wetzlar, Germany) and mounted on 200 MESH copper grids. Post-contrastation in an automated grid stainer (EM AC20, Leica) and drying, ultrathin sections were analyzed and micrographed on a Jeol JEM 1400 Plus TEM equipped with a TemCam-F416 (TVIPS, Gauting, Germany).

Nuclear Magnetic Resonance (NMR)

The presence of free BSA in the emulsion was analyzed via one-dimensional (1D) 1H NMR spectroscopy. Ten percent D2O was added to the emulsion to yield a final volume of 600 μL, including sodium trimethylsilylpropanesulfonate as a standard compound. Spectra were recorded on a Bruker AVANCE III HD 700 MHz spectrometer at 25 °C using the “zggpw5” pulse program from the Bruker pulse scheme catalog. Spectra were processed with Bruker’s software suite Topspin 3.6.1.

To test the emulsion’s stability, emulsion droplets from a sample emulsified with a single pass, 20,000 PSI at 4 °C were purified as described above, aliquoted into 0.5 mL reaction tubes, sealed gas-tight, and stored at 4 °C. NMR spectra were recorded at the day of purification and after 7, 14, 24, 43, 77, and 119 days. A fresh tube was opened for each measurement. The first sample from the day of purification was stored after the measurement in the NMR tube at 4 °C and served as a control as it was remeasured at the same time as the other samples.

Oxygen Capacity

The oxygen capacity of the emulsion was determined using an Oxygraph-2k (Oroboros Instruments GmbH, Innsbruck, Austria). Chamber preparation: chambers were filled with 2 mL of yeast suspension (80 mg/mL) (Thermo Fisher, Waltham, MA) until reaching the oxygen baseline, indicating zero mL/mL O2. The emulsion was oxygenated prior to determination via a custom-made “Kugelrohr” oxygenator with 100% O2 (Air Liquide, Duesseldorf, Germany) and a flow of 0.5 L/min at 37 °C. Samples were obtained using a gas-tight 1.5 mL reaction tube (Sarstedt AG und Co. KG, Nümbrecht, Germany) with a puncture in the lid just about the size of a gas-tight Hamilton syringe (Hamilton Company, Reno, NEV). This gas-tight syringe then was used to inject 50 μL of the emulsion into the detection chamber.

Oxygen Transmission

To quantify the droplet-to-tissue oxygen transmission (DOT), the emulsion was oxygenated using 100% O2 as described above. Blood from three healthy male adults (age 25–35 years) was drawn, and 1 mL was transferred into 1.5 mL reaction tubes. Oxygen saturation (sO2) and oxygen partial pressure (pO2) of the blood were determined (ABL 800 flex, Radiometer, Copenhagen, Denmark) directly before the emulsion was injected into the reaction tube. Afterward, 50 μL of the freshly oxygenated emulsion was injected into the blood-containing reaction tube, and the sO2, pO2, and hemoglobin (Hb) concentrations were determined immediately. The oxygen concentration (cO2) before and after adding the emulsion was calculated according to Hüfner using the following equation41

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To calculate the amount of oxygen that was transferred into the erythrocytes, the concentrations after and before the addition of the emulsion were subtracted, and the absolute amount for 1 mL was calculated. The ratio between the absolute increase of oxygen bound to Hb and the maximal O2 capacity of the emulsion defined the DOT.

The total oxygen capacity of the emulsion was determined using an Oxygraph-2k.

Tissue Compatibility

THP-1 cells were grown at 37 °C in an atmosphere of 20% O2 and 5% CO2 in GibcoTM RPMI-1640 medium (Invitrogen, Waltham, USA) with 10% fetal bovine serum (Sigma Aldrich, Darmstadt, Germany) and a 1% penicillin/streptomycin mix (Invitrogen, Waltham).

Four days before the start of the experiment, the THP-1 cells were transferred to glass-bottom Petri dishes (IBIDI, Martinsried, Germany), and 10 nM phorbol-12-myristate-13-acetate (Sigma Aldrich, Darmstadt, Germany) was added to the medium to differentiate them into M0-like macrophages. The number of cells used was approx. 1 million cells per dish.

To avoid contamination and antigen recognition, the emulsion was made from a clinically approved human serum albumin (HSA) solution (STEEN solution) via a temperature gradient (4–50 °C) and pressure gradient (20,000–30,000 PSI) and stored at 4 °C for 2 weeks. At the beginning of the experiment, the cell medium was replaced by STEEN solution with a PFC droplet volume fraction of 4% and incubated for 4 h at 37 °C and 20% O2 and 5% CO2. The excess medium was taken off the cells using a pipette and fixated without further washing with glutaraldehyde and formaldehyde in PHEM buffer. After 4 h, the buffer was changed to PHEM buffer without glutaraldehyde.

Results and Discussion

Standard Conditions

HPH is a well-established technique used to create emulsions meeting industrial requirements.10 The simplest way of using this kind of emulsification is to apply a medium high pressure of about 2000 PSI once at low temperatures. In this study, we defined this condition as a standard condition, and it is used as a reference for all modifications to the emulsification process.

Recoalescence, Diffusivity, and Marangoni Convection

The imbalance of the two major processes during the formation of the emulsion, (i) the shear forces tearing big PFD droplets apart and (ii) the newly formed smaller droplets tending to fuse directly after formation (recoalescence), is a crucial factor that has to be considered when establishing a protocol for a stable emulsion.13−15 Recoalescence is caused by a density gradient of BSA on the shell of newly formed droplets at the exact same spot where the smaller droplets are pinched off. This gap on the surface exposes naked PFD that has an extreme surface tension difference toward the water phase, as well as diminished repulsive forces toward other droplets.16 Closing the gap depends on two speed-limiting factors: (i) Marangoni convection (degrees of freedom of BSA on the droplet surface) and (ii) the diffusivity of BSA through the surrounding medium (water).12,17,33,42 As compared to previous research dealing with the analysis of the behavior of proteins on interfaces,35,43 a very high concentration of 50 mg/mL BSA was used for emulsification in this study to mimic the physiological oncotic pressure of blood (mandatory for later use). This concentration might mislead us to the assumption that the diffusion distance is shortened in a way that allows for neglecting diffusion in this setup. In fact, it was evident that in the case of HPH, 50 mg/mL BSA was not sufficient to prevent recoalescence (Figure 1A). The increase in the BSA concentration led to a substantial decrease in droplet size. Furthermore, the convection of BSA alongside the droplet’s surface was indirectly measured via 1D 1H NMR measurements of the purified droplets. As the 1D 1H NMR spectra demonstrated (Figure 1B,C), the bands caused by BSA disappear as soon as BSA attaches to the hydrophobic interface. BSA loses the ability to tumble independently, resulting in faster relaxation of the signals.44,45 As purified PFC droplets in BSA-free distilled water are invisible for the 1D 1H NMR measurements, there is reason to assume that the Marangoni convection has only a minor impact on preventing recoalescence.

Figure 1.

Figure 1

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Analysis of the droplet formation: (A) Increase in BSA concentration leads to accelerated droplet formation resulting in decreased droplet size. (B) 1H NMR spectrum of the suspension. BSA gives rise to broad 1H resonances between 10 and 0 ppm. Removing excess BSA from the suspension leads to the disappearance of all BSA-1H signals. BSA bound to the emulsion droplets cannot be detected, presumably because of the much slower molecular tumbling due to the increased size of the droplets. (C) Increasing the emulsification pressure (EP) leads to initially smaller droplets that end up being the same size as droplets that form under lower EP. (D) Increasing the passage count (PC) from 1 to 15 led to initially smaller droplets that showed accelerated growth and ended up being bigger than those that formed during only one passage. (E) Increasing the PC led to a broader droplet size distribution (DSD) reflected by a higher polydispersity index (PDI) right after synthesis. (F) Interrelation between BSA consumption and the EP and PC during formation of the droplet. The average DSD (bars) determined by dynamic light scattering and the consumption of BSA (points) determined by bromocresol green staining and subsequent photometric determination are plotted as a function of EP and PC. With increasing EP and PC, the average droplet size decreased. The consumption of BSA increased with increasing PC and decreased with increasing EP. Mean values ± SD from n = 5; *p < 0.05, **p < 0.002, and ****p < 0.0001; one way analysis of variance (ANOVA) compared to the control followed by Dunnett′s post hoc test.

Pressure, Passages, and BSA Concentration

The variety of manipulation of the emulsifying process of PFC emulsions developed for intravenous application is limited to solely physical variables because chemical manipulation may lead to poisonous side effects.46 A major setscrew may be the alteration of pressure and the passage counts (PCs). But increasing the emulsification pressure (EP) leads only to initially smaller droplets (210 vs 240 vs 310 nm) that ripen faster with the effect that the size of droplets eventually becomes independent of the selected EP (Figure 1D). Increasing the PC from 1 to 15 also led to initially significantly smaller droplets (160 vs 210 nm), which grew faster and eventually generated bigger droplets than those that were formed with lower PC (Figure 1E). Also, the polydispersity index that gives information about the homogeneity of the DSD increases from passage to passage47 (Figure 1F).

To clarify what happens during the droplet formation that causes an effect on the droplets that lasts at least over a period of 25 days, BSA consumption during different HPH conditions was evaluated. Figure 1G shows the consumption of BSA during the formation process in dependence on the EP and the PC. These observations were made: (i) increasing the EP led to a reduced DSD and, therefore, a larger surface area, while lesser BSA was consumed during the formation and (ii) increasing the PC led to drastically reduced DSD but only a minor increase in BSA consumption.

How can this be explained? The droplets are reversibly compressed as they enter the microchannels of the homogenizer (Scheme S1), and therefore, with increasing EP, they offered less interfacial space for BSA to attach, which led to increased surface tension.48 Further, the increase of EP accelerated the droplets and thus decreased the dwelling time of the droplet and shortened the attachment time for BSA. This again has two consequences. (i) Lower BSA abundance at the interface and (ii) not enough time for the BSA to adhere to its kinetically preferable orientation, which again leads to destabilization of the surface.28,32,34,35,43,49 Repeating this procedure at low temperatures (4 °C) by increasing PC only amplified these effects. In addition, the curvature of the droplets increased, which hindered BSA from accessing the surface in the most stable conformation.50−52 The emulsification process experienced a transition from a diffusion-limited system (diffusion timescale ≫ adsorption timescale) to a kinetic-limited system (diffusion timescale ≪ adsorption timescale).53,54 This perspective explains why there is less BSA adhering to the surface (when using higher EP and increasing PC), which in turn leads to an inadequate BSA density on the surface of each droplet (Figure 1G). Therefore, the surface tension of the droplet is insufficiently reduced, and thereby, the droplet becomes more and more unstable, causing the ripening expressed by the droplet growth seen in the data presented in Figure 1D,E.

Surface Structure and Shell Thickness

Applying the standard conditions led to a surprisingly stable emulsion that showed no sign of decay for at least approximately 24 days and only a slight decay until the end of the study on day 119. The decay of an emulsion is defined by droplet growth by both Ostwald ripening and coalescence and is accompanied by the dissolution of surface-bound surfactant into the surrounding medium. For this reason, we measured the free BSA concentration of an emulsion created under standard conditions for 119 days (Figure 2). Inhibiting the evaporation led to the observation that BSA did not dissolve into the medium within the observed period. This allows for the conclusion that any considerable decay was absent, and neither Ostwald ripening nor coalescence was a major driving force for decay in this emulsion. Unfortunately, the droplet size distribution was unacceptably large for any intravenous application.

Figure 2.

Figure 2

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(A) Samples were stored in closed reaction vessels for 119 days at 4 °C. 1D 1H NMR spectra were recorded on days 0, 7, 14, 43, 77, and 119. (B) Sample was stored in an unsealed NMR tube at 4 °C for 119 days. 1D 1H NMR spectra were recorded on days 0, 7, 14, 43, 77, and 119.

Taking all of these observations into consideration, droplets derived from a single passage at moderate EP should have a thicker BSA shell than droplets derived from higher PC and higher EP.55 To get a glance at the shell thickness, the purified droplets of a single passage and 20,000 PSI were analyzed using cSEM. This technique includes a step in which, at −100 °C, water sublimates by decreasing the atmospheric pressure. These conditions also resulted in the sublimation of PFC, which in turn forced the frozen BSA shell to burst, giving us the opportunity to measure the breakage as done in Figure 4D. In the process of sputtering, gold application to the surface of the sample enables electrical conductivity of the sample, which is necessary to perform cSEM. Considering (i) a gold layer of around 10 nm and (ii) the assumption that the breakage is rolled-up and therefore double-layered, Figure 4D shows a wall thickness of about 5–10 nm. BSA is an anisotropic molecule with the dimensions of 4 × 4 × 7 nm3, i.e., the shell layer probably consists of a dual layer of BSA, which is in line with the literature.26,56

Figure 4.

Figure 4

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(A) Oxygen transfer: injection of 50 μL of preoxygenated PFC emulsion was able to increase the oxygen levels in 1 mL of blood by around 2.26 mL of O2 (n = 6, t-test, p = 0,006). (B) Schematic visualization of the DOT. (C) TEM image of a THP-1-derived macrophage with incorporated emulsion droplets emulsified using a temperature and pressure gradient with HSA as a surfactant. (D) TEM image of incorporated HSA–emulsion droplets. After 4 h of incubation, the droplets remain in their primal shape and size.

Problems with Size Analysis

As seen in Figure 1F, the dispersity of the emulsion increased with increasing PC and exceeded a PDI of 0.7, which is defined as the limit to generate precise DSD using DLS.47 Combined with a refractive index that is almost not distinguishable from that of water, it was almost impossible to obtain reliable results even after only one passage. Figure 4 shows a combination of DLS, SLS, and microscopic analysis of the droplets. The intensity-mode DLS determined that DSD shown in (A) followed a Gaussian distribution peaking at 223 nm, whereas the volume-mode SLS data in (C) showed an almost Gaussian distribution peaking at 100 nm with a second population appearing between 2 and 20 μm. Interference microscopy that allows for a resolution down to 300 nm confirmed the results (B). Using the advanced PIDS technology enables the observation of droplets in the lower nanometer range as well as the micrometer range, whereas DLS is not suitable for droplet diameters above 1 μm. The refractive index used for the measurements differed, as the refractive index is size-dependent, and therefore, the SLS measurements needed adjustments to the refractive index, whereas the DLS measurements showed the same droplet diameters independent of the refractive index (Figure S41). However, for clinical application, it is important to determine the DSD, and further, a broad DSD may lead to complications such as embolism or activation of the immune system.47

Increasing BSA’s Efficacy to Adsorb at the Interface

So far, we have discovered that applying higher EP and PC results in unsatisfactory short shelf life, mainly caused by both (i) the low efficacy and velocity of BSA to absorb and (ii) the decreased tension at the liquid–liquid interface.57 To increase the adsorption efficacy of BSA, a pressure and temperature gradient was applied, which is easier to control than alternatively starting with a higher temperature.17,58,59 To gradually increase the temperature, the cooling coil of HPH was simply not covered in ice. The inner friction of the emulsion resulted in a 2 °C increase per passage starting at 4 °C and 20,000 PSI. On reaching 40 °C, the EP was increased to 30,000 PSI and was kept at this level until reaching a temperature of 50 °C. We detected that a temperature above 50 °C led to denaturation, thereby destructing the emulsion. Important for the durability of the emulsion was the rapid temperature decrease to 0–4 °C after the process.

The gradients led to a clear emulsion that showed Rayleigh scattering at an angle of around 90 ° (Figure S2), which is part of the definition of a nanoemulsion.55 Normal microscopic analyses failed in identifying any visible droplets at any possible magnification. Thus, a determination of the DSD via SLS was validated using cTEM images.60 One day after emulsification, both analyzed emulsions neither sedimented nor flocculated, which would have been signs of agglomeration and coalescence.61 The cTEM images shown in Figure 4E,F validated this assumption: both emulsions display no sign of agglomeration. The SLS measurement shown in Figure 4G presents a DSD of the emulsion homogenized using the gradient method with a mean diameter of around 150 nm in volume mode with an upper limit of ∼350 nm. FIJI-based measurements of the SDS using cTEM images (Figure 4E,F) revealed mean diameters of 140 and 341 nm in the largest droplet. Therefore, the cTEM images confirmed the results of the SLS measurement. Storing the emulsions for 5 months at 4 °C resulted in an increased DSD with a mean diameter of 300 nm with an upper limit of around 600 nm.

Even though the pressure and temperature gradients led to a smaller DSD and a tremendously increased shelf life of the emulsion, the ESA measurement revealed no changes in the surface charge. The ESA signal cannot be accurately translated into ζ-potential because both emulsions are not monodisperse.62 However, this is where the advantage of the ESA method becomes apparent because, for both emulsions, the ESA signal can be addressed as a raw data signal. Both emulsions showed signals of about −2 to −2.5 (mPa (M/V)) at the physiological blood pH of 7.4, which shows a relatively high electrostatic stability (data shown in Figure S3).

Increasing the temperature led to a higher surface coverage of the droplets by either partial denaturation of adhered BSA or via mitigation of the transition from a diffusion-limited system to a kinetic-limited system due to the increased mobility of BSA.59,63 Both models may explain the substantial increase in stability observed in the NMR measurements shown in Figure 2.

This protocol was created using BSA but was transferable to HSA without any further adjustment of the protocol (Figure 3C,D).

Figure 3.

Figure 3

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(A) Intensity-mode DLS diagram of DSD showing a mean diameter of around 223 nm. (B) Interference microscopy image of an emulsion whose droplets are visible down to a diameter of 1 μm. The limit of detection for this microscopic method is about 300 nm; therefore, all visible droplets have diameters above the average diameter determined in DLS and SLS. (C) Volume-mode SLS diagram of the DSD showing a bimodal distribution. One population at around 150 nm and one population between 2 and 20 μm. (D) Size determination of albumin-PFC emulsion droplets using cSEM. Measurement of the shell thickness of BSA-PFC emulsion droplets after breakage during sublimation of PFC. (E)  cTEM image of the emulsion droplets emulsified with static temperature (4 °C) and a single passage at 20,000 PSI. (F)  cTEM image of emulsion droplets emulsified using a temperature and pressure gradient. (G) SLS diagram of repeated measurements presenting the DSD of two emulsion droplets emulsified using a temperature and pressure gradient on day 1 (white and light gray) and 5 months after emulsification (dark gray) in volume mode.

Oxygen Transfer and Biological Resilience

The most important property of the emulsion would be to transfer oxygen from the lungs to the tissues of the body. The oxygen capacity of a PFC emulsion is proportional to the PFC content and increases linearly with increasing PFC content. The most commonly used PFCs for oxygen carrier emulsions are perfluorodecalin (0.403 mLO2/mLPFC), perfluorooctylbromid (0.527 mLO2/mLPFC), and dodecafluoropentane (0.029 mLO2/mLPFC).2 This study investigated a perfluorodecalin-based emulsion, which proved to be highly potent in concentrations of 4–6% to maintain the oxygen supply in a model of ex vivo perfusion of a rat kidney and in a severe hemodilution model of the rat.7,21 To test the maximal capability to transfer the oxygen, this DOT was determined using human blood of three volunteers in which preoxygenated emulsion with a volume fraction of 17% was injected in gas-tight syringes. The theoretical amount of O2 in 50 μL of the emulsion is 3.34 μL. Amperometric measurements with an Oxygraph-2k revealed an actual oxygen capacity of around 3.5 to 3.7 μLO2/50 μL emulsion. This DOT is in the range of oxygen carriers that reached clinical trials and carry 3 μL of O2 in 50 μL emulsion. Blood can carry up to 10 μL of O2 in 50 μL, but because of the unique biochemical structure of hemoglobin, it can only release around 2.5 μL under physiological conditions.5 As shown in Figure 4A, the injection of 50 μL of emulsion at 37 °C increased the oxygen content of 1 mL of blood by around 2.26 μL of O2. This increase reflects ca. 60% of the amount of oxygen carried by 50 μL of the emulsion but mimics the amount of oxygen supplied by the blood. This result demonstrates that even under normoxic conditions, the emulsion droplets can transfer physiological amounts of oxygen into red blood cells and other tissues.

Another important point is to address the biological resilience, as this emulsion shall be used clinically as an artificial oxygen carrier. Emulsion droplets can be recognized by the body’s immune system and may be incorporated by macrophages. In the long term, PFC accumulates in the spleen and liver, where it can interact with enzymes.2 If the size of the droplet is less than 150 nm, it may also be taken up by pinocytosis by cells outside the immune system, e.g., endothelial cells.2,64−67 The DSD of the emulsion emulsified via pressure and temperature gradients had a maximum droplet size of approx. 200 nm, and the droplets may, therefore, be incorporated into cells via phago- or/and pinocytotic uptake.68,69 For this experiment, the emulsion was generated with human serum albumin using a clinically approved solution (STEEN solution) to avoid antigen recognition of BSA or unwanted activation of the macrophages by impurities of the BSA solution. Still, the utilized macrophages were able to incorporate the emulsion droplets to fuse the resulting endosomes with lysosomes, resulting in heterolysosomes (Figure 3C), which contain proteases and peptidases. These enzymes are used by the cell to degrade proteins into peptides and amino acids for both elimination of pathogens and energy generation. Interestingly, as shown in Figure 3D, the droplets did not lose their primary shape and size, indicating that the enzymes are not able to degrade the BSA shell of the droplets sufficiently, generating an unforeseen biological resilience. This property may even allow for the process of exocytosis, leading to desirable longer blood retention of the emulsion droplets.70 An overview of the observed cellular uptake is given in Figures S5–S40.

Summary

The size determination analysis of BSA-based PFC–water emulsions is challenging because of (i) their refractive index, which is extremely close to the refractive index of water, (ii) the large PDI that disqualifies them for analysis via DLS, and (iii) small droplet size that disqualifies them for analysis via optical microscopy. We identified SLS/PIDS technology as the only reliable method to determine the DSD of these emulsions. A variety of methods such as cTEM, TEM, and optical microscopy verified and confirmed our data obtained with this technology. To further obtain a comprehensive picture of the emulsion, methods such as ESA, cSEM, and 1H NMR were used to characterize the shell properties of a single emulsion droplet and its stability over time.

During HPH of PFC in water, using BSA as a surfactant, the increase of emulsification pressure and the increases of the passage count at 4 °C led to long-term destabilization of the emulsion. We assume that this effect was caused by the inertia of BSA, which hindered the diffusion and, therefore, the occupation of empty binding sites at the surface of nascent emulsion droplets. Using moderate emulsification conditions led to a surprisingly stable emulsion that showed only minor signs of decay for up to 119 days. However, this emulsion contained a broad droplet size distribution with droplet sizes larger than 5 μm, which is incompatible for clinical intravenous use as an oxygen carrier. To reduce this droplet size and decrease the DSD, it was necessary to overcome BSA’s inertia by introducing heat to the emulsification process. This was realized by application of a temperature and pressure gradient, leading to a stepwise increase of the temperature that was tolerated by the emulsion to up to 50 °C. These adjusted emulsification conditions limited droplet ripening that was observed at a time of about 6 months. This temperature-controlled protocol was applied to create an emulsion based on human serum albumin without further adjustment. This emulsion showed remarkable biological resistivity and promise to work in a clinical context.

Conclusions

Considering the extensive characterization, we concluded that the stability of a PFC–water emulsion with albumin as the surfactant is crucially dependent on albumin′s mobility during the emulsification process and on sufficient occupation of binding sites by the surfactant on the surface at the PFC–water interface. We discovered that applying a temperature and pressure gradient is a convenient way to increase albumin′s mobility during the emulsification process, which led to the generation of a nanoemulsion with a tremendously prolonged shelf life of at least 6 months. This temperature-controlled protocol also worked for HSA.

Acknowledgments

The authors thank Susanne Eitner and Sylvia Voortmann for their excellent technical assistance. Scanning electron microscopy (SEM) experiments were performed at the Imaging Center Essen, University Hospital Essen. cSEM was performed by Labor Dr. Schäffner GmbH, Solingen, Germany. They also acknowledge support by the Open Access Publication Fund of the University of Duisburg-Essen.

Glossary

Abbreviations

BSA

bovine serum albumin

cSEM

cryo-scanning electron microscopy

cTEM

cryo-transmission Electron Microscopy

DOT

droplet-tissue-oxygen-transmission

EP

emulsification pressure

HPH

high-pressure homogenization

PC

passage count

PFC

perfluorocarbon

PFD

perfluorodecalin

pO2

oxygen partial pressure

sO2

oxygen saturation

TEM

transmission electron microscopy

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.langmuir.1c03388.

  • (Scheme S1) Visualization of the emulsification process inside the microchannels of the microfluidizer; (Figure S1) refractive indices of the emulsion in dependence on the wavelength; (Figure S2) photograph of the nanoemulsion revealing the transparency; (Figure S3) ESA measurements of an emulsion produced using one passage and an emulsification pressure of 20,000 PSI and an emulsion using a temperature and pressure gradient; (Figures S5–S40) electron microscopy images of the uptake of emulsion droplets into THP-1 cells; and (Figures S41–S43) impact of the refractive index on DLS and SLS measurements: droplet size distribution and SLS intrinsic residual value (PDF)

Author Contributions

All authors have given approval to the final version of the manuscript. Study design: J.J. and K.B.F. Writing: J.J., S.H., O.A., D.M., A.H., O.E.P., H.J., K.B.F., C.S., and M.K. Experimentation: J.J., O.A., D.M., S.H., A.W., F.N., A.H., R.F., M.C., H.J., and S.P. Analyzed data: J.J., S.H., O.A., D.M., A.H., and R.F. Interpreted data and discussion: J.J., S.H., O.A., D.M., A.W., H.J., R.S., K.B.F., M.K., and C.S.

This project was funded by the German Federal Ministry of Economics, grant numbers 16KN039249 and 16KN039250. This work (cryo-TEM) was supported by grants from the Deutsche Forschungsgemeinschaft SFB 944 (Z-Project).

The authors declare the following competing financial interest(s): Dr. Johannes Jaegers, Dr. Sven Haferkamp, Dr. Oliver Arnolds, Fabian Nocke, Dr. Miriam Cantore, Stefanie Pütz, Dr. Carsten Schauerte, Prof. Raphael Stoll, Prof. Michael Kirsch and Prof. Katja Ferenz are holders of the patent application Organ life fluid - Bioactive perfusion medium for the normothermic perfusion and regeneration of isolated human kidneys German docket number: 102021211272.2.

Notes

The Ethics Council of the University Hospital Essen authorized the collection of blood for the purpose of experimental studies. (file number 19-8600-BO).

Supplementary Material

la1c03388_si_001.pdf (12.1MB, pdf)

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