OPSALC: On-Particle
Solvent-Assisted Lipid Coating
to Create Erythrocyte Membrane-like Coatings with Improved Hemocompatibility
Introduction Particles are an essential
foundation of nanomedicine, having contributed
to the establishment of targeted drug delivery, enhanced imaging,
and precision diagnostics. − Particles have recently started being explored as cell-like materials,
with emerging reports on engineered platelets, macrophages , and red blood cells (erythrocytes) − for targeted therapeutics. Particle administration is often done
via intravenous or intratumoral delivery, , which
includes blood contact, and therefore demands the use of hemocompatible
materials. Most particles (e.g., micro and nanomaterials) are
either poorly
or not fully hemocompatible, and demand surface modifications or coatings
to facilitate their use in blood-contact applications. Common strategies
to improve particle hemocompatibility include the application of surface
modifications or coatings, such as heparin, − albumin − or lipids, − but the use of heparin poses some risk of thrombotic events in patients , and the use of albumin coatings shows a short-lasting effect, predominantly
due to the displacement of albumin by other serum proteins in blood. Recently, the use of blood cell lipid membrane
“ghosts”, i.e. membranes isolated from leukocytes, platelets,
or erythrocytes, has shown great promise in the improvement of particle
hemocompatibility, demonstrating a notable reduction in macrophage
response and prolongation of circulation time. − Despite their advantages, blood cell membrane “ghosts”
are scarcely available, contain inherently instable membrane proteins, and may give rise to patient-specific complications
due to the immunogenicity of membrane antigens. To overcome these
obstacles, artificial lipid membrane coatings can be engineered using
synthetic lipids, ultimately providing a stable, tunable, and disease-free
coating. , However, an efficient and scalable method
to apply cell membrane-like lipid nanocoatings on particles remains
unexplored. Lipid coatings can be applied to material surfaces
through a number
of techniques, such as vesicle fusion, − Langmuir–Blodgett
deposition , or dip pen lithography. One of the most commonly employed techniques
in the lipid-coating of particles is vesicle fusion, wherein small
uniform lipid vesicles adhere to a particle surface, rupture and spread
out to form a continuous bilayer. This process has been studied in
great detail for forming supported lipid bilayers on planar surfaces. − Similarly, to efficiently coat particles with lipids through vesicle
fusion, the pH, ionic strength and the choice of charged lipid headgroups
need optimization to enhance vesicle absorption to the surface due
to attractive forces, while reducing intervesicle interactions and
particle aggregation. Consequently, for
most particle material-lipid combinations, lipid vesicles do not adsorb
to particles or, when they do adsorb, they often only partially rupture
at the surface, yielding incompletely covered particles. , To overcome these obstacles, exciting advances have been reported
on solvent-exchange protocols that allow for the lipid-coating of
a near-universal variety of planar solid surfaces. − In these solvent-exchange protocols,
the organic solvent, in which the lipids are dissolved, is gradually
diluted by the addition of water. This solvent–water gradient
will gradually induce a lipid assembly from monomeric lipids and inverted
micelles to micelles, and then vesicles. Shortly before the onset of the micelle-to-vesicle transition, a
lipid bilayer nucleates at the solid planar surface, forming a cell
membrane-like lipid layer coating. In
contrast to vesicle fusion, solvent-exchange protocols require less
handling and can be easily implemented in different scalable formats
such as continuous flow set-ups. In addition, particles would not
require a laborious tailoring of their surface prior to applying solvent-exchange. , While these strategies have been vastly successful in applying lipid
coatings to planar substrates, the successful lipid-coating of nonplanar
substrates, such as particles, through solvent-exchange methods has
not yet been documented. In this work, we introduce an on-particle
solvent-assisted lipid
coating (OPSALC) method, which combines the versatility of solvent
exchange lipid coating methods with a simple experimental setup, compatible
with a wide range of particle substrates. In short, an aqueous buffer
is gradually added, while stirring, to a concentrated mixture of lipids
and particles in a water-miscible alcohol. The induced gradient then
originates a lipid-layer coating on the particles through a gradual
lipid assembly ( a, Figure S1 ). We first validated the
method using a single-lipid solution of 1,2-dioleoyl- sn -glycero-3-phosphocholine (DOPC) on silica nanoparticles (SiO 2 NPs) and microparticles (SiO 2 MPs). Next, we applied
the OPSALC method to coat SiO 2 and polycaprolactone (PCL)
particles using four erythrocyte cell membrane-mimicking lipid coatings
( b) to study
their influence on the overall hemocompatibility of the particles. 

Experimental Section 2.1 Materials Colloidal silica nanoparticles
(diameter 0.10 ± 0.03 μm) were purchased from Polysciences
Inc. DiagNano green fluorescent silica microparticles (diameter 5
μm), monodisperse, with hydroxyl surface functionalization,
were purchased from CD Bioparticles. Polycaprolactone (PCL) M w ∼ 14,000, Mowiol 40–88, HEPES
(≥99.5%), calcium chloride dihydrate (ACS reagent, ≥
99.0%), sodium chloride (ACS reagent, ≥ 99.0%) and ethylenediamine-tetraacetic
acid disodium salt dihydrate (EDTA, ≥ 99.0%) were purchased
from Sigma-Aldrich. 1,2-dioleoyl- sn -glycero-3-phosphocholine
(DOPC), cholesterol, egg sphingomyelin (SM), and soy phosphatidylinositol
(PI) were purchased from Avanti Polar Lipids (Sigma). Texas Red-DHPE
(TR-DHPE) was purchased from ThermoFisher Scientific. All lipids were
purchased in powder form, dissolved in chloroform, and stored at −20
°C except TR-DHPE, which was solubilized in methanol and then
stored at −20 °C. Dulbecco’s Phosphate Buffered
Saline (DPBS, sterile), DPBS with CaCl 2 and MgCl 2 (sterile), chloroform (ACS spectrophotometric grade, ≥ 99.8%),
dichloromethane (DCM, ACS reagent, ≥ 99.5%), methanol (HPLC
grade, ≥ 99.9%), and isopropanol (IPA, ACS reagent, ≥
99.5%) were purchased from Sigma-Aldrich. Ethanol (absolute, ≥
99.5%) was purchased from VWR Chemicals. Ultrapure water (resistivity
18.2 MΩ•cm) was obtained from a Milli-Q setup (Millipore).
DPBS was sterile-filtered with 0.2 μm syringe filters prior
to each experiment. 2.2 OPSALC on SiO 2 Nanoparticles Suspensions of SiO 2 NPs were prepared in ethanol and
isopropanol at a concentration of 10 mg mL –1 . A
lipid solution of DOPC was prepared in the same solvents at concentrations
of 0.5, 1.0, 2.0, and 4.0 mg mL –1 . To coat the particles
through the OPSALC method, a concentrated lipid-particle suspension
was first prepared by adding 50 μL of SiO 2 NPs to
450 μL of lipid solution in a small glass vial. The gradient
was then induced by the injection of DPBS into the vial at a specific
addition rate (600 μL min –1 or 60 μL
min –1 ) using a syringe pump (KD Scientific). This
buffer addition was performed under magnetic stirring at 500 rpm,
up to a volume of 5 mL, reaching a final solvent:buffer ratio of 10:90
(v/v). The resulting suspension was then heated to 50 °C for
45 min and left to cool down to room temperature. Particles were washed
three times in ultrapure water by centrifuging at 10000 g for 10 min at 4 °C. During washing, the supernatant was removed
and the pellet was resuspended in fresh ultrapure water using a micropipette
(Eppendorf). After washing, particle pellets were kept at 4 °C
until further use. For the study on the effect of lipid concentration,
the lipid-particle mixture was prepared in ethanol using a buffer
addition rate of 600 μL min –1 and a lipid
concentration of 0.5, 1.0, 2.0, or 4.0 mg mL –1 ,
corresponding to an approximate lipid bilayer:particle ratio ( n ) of 4:1, 8:1, 16:1, and 32:1 ( Methods S1 ). For the study on buffer addition rate, the lipid-particle
mixture was prepared in ethanol using a lipid concentration of 1.0
mg mL –1 and a buffer addition rate of 600 or 60
μL min –1 . Lipid vesicles were produced as
the respective controls, under the same conditions, using pure ethanol
instead of the SiO 2 NP suspension. For the study on solvent,
the lipid-particle mixture was prepared in ethanol or isopropanol
using a lipid concentration of 1.0 mg mL –1 and a
buffer addition rate of 600 μL min –1 . Lipid
vesicles were produced as the respective controls, under the same
conditions, using pure ethanol or isopropanol instead of the SiO 2 NP suspension. Lastly, for the study on solvent:buffer ratio,
the lipid-particle mixture was prepared in ethanol using a lipid concentration
of 1.0 mg mL –1 and an addition rate of 600 μL
min –1 , at a final solvent:buffer ratio of 100:0,
50:50, or 10:90 (v/v). The final volume was maintained at 5 mL, and
the ratios were obtained by adding 4.5 mL of ethanol (100:0), 2 mL
of ethanol with 2.5 mL of DPBS (50:50), or 4.5 mL of DPBS (10:90). 2.3 OPSALC on SiO 2 Microparticles Suspensions of SiO 2 MPs were prepared in ethanol and
isopropanol at a concentration of 10 mg mL –1 . A
fluorescent lipid solution of DOPC (99.9 mol % DOPC, 0.1 mol % TR-DHPE)
was prepared in the same solvents at concentrations of 0.25, 0.5,
1.0, and 2.0 mg mL –1 , corresponding to
0.1, 0.2, 0.4, and 0.8 mg per mg of SiO 2 MPs, or n ≈ 44, 88, 176, and 353 lipid bilayers per particle,
respectively ( Methods S2 ). To coat the
particles through the OPSALC method, a concentrated lipid-particle
suspension was prepared by mixing 40 μL of SiO 2 MPs
with 160 μL of lipid solution in a small glass vial. The gradient
was then induced by the injection of DPBS into the vial at a manual
pipetting rate (∼3 mL min –1 ) or at controlled
addition rates (600 μL min –1 or 60 μL
min –1 ) using a syringe pump (KD Scientific). The
buffer was added under magnetic stirring at 180 rpm up to a volume
of 2 mL, reaching a final solvent:buffer ratio of 10:90 (v/v). The
resulting suspension was then heated to 50 °C for 45 min and
left to cool down to room temperature. Particles were washed three
times in ultrapure water by centrifuging at 50 g for
4 min, and were kept at 4 °C until further use. The conditions
used during the studies on solvent, buffer addition rate, solvent:buffer
ratio, and lipid concentration are specified in detail in Tables S1–4 . 2.4 Preparation of Erythrocyte Membrane-like Lipid
Formulations Based on lipidomics studies of the outer leaflet
of the erythrocyte membrane, we prepared
four lipid formulations of distinct biomimicry complexity. Fluorescent
and nonfluorescent versions of the same lipid formulation were prepared.
Fluorescent formulations included DT (DOPC, TR-DHPE), CT (DOPC, cholesterol,
TR-DHPE), ST (DOPC, cholesterol, sphingomyelin, TR-DHPE), and IT (DOPC,
cholesterol, sphingomyelin, phosphatidylinositol, TR-DHPE). Nonfluorescent
formulations (D, C, S, and I) included the same lipids at the same
lipid ratios, only without TR-DHPE. Detailed information on lipid
ratios can be found in Table S5 . 2.5 OPSALC of SiO 2 MPs with Membrane-like
Formulations SiO 2 MPs were prepared at 10 mg mL –1 in isopropanol. Fluorescent lipid formulations DT,
CT, ST, and IT were prepared at 0.5 mg mL –1 in the
same solvent. A concentrated lipid-particle suspension was first made
by adding 80 μL of SiO 2 NPs to 320 μL of lipid
solutions in small glass vials. The gradient was then induced with
the addition of 3.6 mL of DPBS, after which the suspension was heated
to 50 °C for 45 min and left to cool down to room temperature.
Particles were washed three times in ultrapure water by centrifuging
at 50 g for 4 min, and finally redispersed in DPBS
for analysis. 2.6 OPSALC of SiO 2 NPs with Membrane-like
Formulations SiO 2 NPs were prepared at 10 mg mL –1 in isopropanol. Nonfluorescent lipid formulations
D, C, S, and I were prepared at 1.0 mg mL –1 in the
same solvent. A concentrated lipid-particle suspension was first made
by adding 50 μL of SiO 2 NPs to 450 μL of lipid
solutions in small glass vials. The gradient was then induced with
the addition of 4.5 mL of DPBS, after which the suspension was heated
to 50 °C for 45 min and left to cool down to room temperature.
Particles were washed three times in ultrapure water by centrifuging
at 10000 g for 10 min at 4 °C and finally redispersed
in DPBS for analysis. 2.7 OPSALC of PCL MPs with Membrane-like Formulations PCL MPs were produced through a single emulsion solvent evaporation
technique. A solution of 0.3% w/v Mowiol 40–88 (25 mL) was
prepared in water, to which a 3% w/v solution of PCL in dichloromethane
(1 mL) was added. The two-phase mixture was homogenized using tip
ultrasonication for 10 s on ice, with a pulse of 1 on 1 s off and
an amplitude of 25% (Fisherbrand Q500 Sonicator). The emulsion was
stirred at 600 rpm overnight to allow the dichloromethane to evaporate.
The resulting suspension was washed three times in ultrapure water
by centrifuging at 1500 g for 15 min at 4 °C,
removing the supernatant, and redispersing in water using a micropipette
(Eppendorf). For OPSALC, PCL particles were resuspended in the appropriate
amount of isopropanol to yield a concentration of ∼10 mg mL –1 . Similar to SiO 2 MPs, a concentrated lipid-particle
suspension was made by adding 80 μL of PCL MPs to 320 μL
of fluorescent lipid solution (DT, CT, ST, or IT, 1.0 mg mL –1 ) for coating analysis. As an alternative, nonfluorescent lipid solutions
(D, C, S, or I, 1.0 mg mL –1 ) were used for the preparation
of hemocompatibility assays. The gradient was then induced with the
addition of 3.6 mL DPBS while magnetically stirring at 500 rpm. The
suspension was further heated to 50 °C for 45 min and left to
cool down to room temperature. Particles were washed three times in
ultrapure water by centrifuging at 2500 g for 15 min at 4 °C
and finally redispersed in DPBS. 2.8 Confocal Laser Scanning Microscopy Uncoated and OPSALC lipid-coated SiO 2 MP suspensions (0.2–0.4
mg mL –1 in DPBS) were imaged using a Zeiss LSM 880
confocal fluorescence microscope. A 20X air objective with a numeric
aperture of 0.8 and a 40X water immersion objective with a numeric
aperture of 1.2 were used. Image processing and analysis was performed
using Fiji software. Image acquisition
parameters (objective, acquisition speed, laser power, gain, and bit
depth) and processing methods were kept constant between all the conditions
in comparative studies to allow for a direct comparison. For solvent:buffer
ratio studies, ethanol and isopropanol conditions were acquired at
different laser powers and were therefore only compared qualitatively.
For lipid fluorescence intensity quantification, images were acquired
with the 20X objective at zoom 1.0 (n = 3 images per condition). To
determine the mean fluorescence intensity of TR-DHPE per particle,
an automatic threshold was applied to the red-channel image in order
to identify each particle as a region of interest (ROI). The mean
fluorescence intensity of each ROI was then measured, merged data
from a total of n = 3 images per condition, and plotted as the lipid
content per particle, per condition, using Origin software (OriginLab).
Z-stack images were acquired with a z -interval of
0.46 μm with the 40X objective. Orthogonal views of Z-stacks
were produced with Fiji software. Coating efficiency was determined
as the percentage of red-fluorescent particles (i.e., lipid-coated)
per green-fluorescent particles (i.e., all particles) within a specific
area of the sample. Calculations were performed for two sample areas
across two replicate batches, in a total of n = 557 particles. Where
indicated, images were equally adjusted for brightness (i.e., grayscale
range adjustment propagated for all images) for improved visualization. 2.9 Epifluorescence Microscopy PCL MPs
were analyzed in an ECHO Revolve epifluorescence microscope (ECHO).
Images were acquired at 100% light intensity, with an exposure of
10 ms for transmission and 450 ms for Texas Red channels. Images were
equally adjusted for Texas Red intensity and merged with transmission
images using Fiji software. 2.10 Dynamic Light Scattering (DLS) and Zeta Potential
(ZP) The size distribution and zeta potential of lipid-coated
SiO 2 NPs was determined using a ZetaSizer Lab with 90°
scatter angle measurements (Malvern). Three particle batches (n =
3) were produced, and each condition of each batch was measured in
5-fold in diluted DPBS (0.05X), at a particle concentration range
of 0.05–0.1 mg mL –1 . 2.11 Cryo-transmission Electron Microscopy (cryo-TEM) Uncoated SiO 2 NPs, OPSALC lipid-coated SiO 2 NPs, and lipid vesicle suspensions (approximately 5 mg mL –1 in DPBS) were imaged using a FEI Tecnai T20 transmission electron
microscope operating at 200 keV. Suspensions were vitrified with liquid
ethane (Vitrobot, FEI) and examined on a Gatan cryo-stage. Images
were taken under low-dose conditions using a slow scan CCD camera.
For better visualization of the coating nanostructure, images were
adjusted for brightness and contrast. Particle and vesicle size were
quantified using FIJI software. Lipid
coating states were visually identified (“uncoated”,
“partially coated”, “coated”, “coated–lamellar”,
or “coated–vesicular”) and quantified for the
analysis of coating efficiency. 2.12 Fluorescence Recovery after Photobleaching
(FRAP) SiO 2 MPs coated with fluorescent formulations
(0.2–0.4 mg mL –1 ) containing TR-DHPE were
subjected to FRAP measurements on a Nikon A1 confocal microscope (Nikon)
with a 20X objective (numerical aperture 0.75) in the Texas Red channel.
Circular ROIs on isolated particles with a diameter of 1 μm
(SiO 2 ) or 2 μm (PCL) were bleached for ∼2
s (SiO 2 ) or ∼100 ms (PCL) using the 488 and 561
nm LASERs simultaneously (SiO 2 ) or the 561 nm LASER only
(PCL), at maximum power. Reference ROIs, used for fading correction,
were always of equal dimensions as their respective bleach ROIs and
situated on a different particle. After bleaching, the bleach and
reference ROIs were imaged for up to 5 min to record fluorescence
recovery. Recovery curves are plotted as ratios of background corrected
intensities of bleached ROI/reference ROI as fading correction in
time. 2.13 Blood Collection Human whole blood
was collected from healthy volunteers at the Blood Donor Service of
the Technical Medical Centre of University Twente and used within
4 h of withdrawal. The research did not fall within the scope of the
Dutch Medical Research Involving Human Subjects Act. Informed consent
was obtained from all volunteers and the used blood collection procedure
was approved by the Medical Research Ethics Committee (METC Twente,
reference K11–23). For hemolysis and plasma coagulation assays,
blood was collected in Vacutainer tubes (BD) containing anticoagulant
trisodium citrate with citric acid and dextrose (ACD-A) and treated
in the first hour after withdrawal. For multiplex assays, blood was
collected in Greiner tubes without additive and immediately anticoagulated
with sodium heparin (LEO) to a final concentration of 1.5 IU ml –1 . 2.14 Hemolysis Assays Erythrocytes (here
abbreviated to RBCs, from Red Blood Cells) were isolated based on
a protocol described elsewhere. Briefly,
anticoagulated whole blood was first centrifuged at 300 g for 2 min, after which the position of the upper layers of plasma
and buffy coat were marked on the tube. Both upper layers were removed,
substituted with an equal volume of DPBS, and used to redisperse RBCs.
Cells were washed three times with DPBS using the same centrifugation
conditions and redispersed in the same volume of DPBS. This suspension
was then diluted 50 times in the same buffer. For hemolysis
assays, 950 μL of diluted RBC suspensions were incubated with
50 μL of a 4 mg mL –1 suspension of uncoated
or coated SiO 2 NPs, at a total particle concentration of
approximately 0.2 mg mL –1 . For the negative controls,
RBCs were incubated with 50 μL of DPBS, and for positive controls,
cells were incubated with 50 μL of a Triton-X solution. Particle
suspensions at the same concentration in pure DPBS were included as
controls for absorbance correction. All samples were incubated at
37 °C for 4 h, with gentle mechanical agitation once every hour,
and then centrifuged at 300 g for 5 min. The supernatant
was transferred to a well-plate for quantification of released hemoglobin
through absorbance measurements at a wavelength of 540 nm (150 μL,
n = 5 measurements per condition). Hemolysis rates (HR) were calculated
according to , 1 where A S,RBCs is the absorbance
of the supernatant of each condition of SiO 2 NPs incubated
with RBCs; < A S,DPBS > is the mean absorbance of the
supernatant of the same SiO 2 NP condition in DPBS; <
A UC, RBCs > is the mean absorbance of the supernatant
of uncoated SiO 2 NPs incubated with RBCs; and < A UC,DPBS > in the mean absorbance of the supernatant of uncoated
SiO 2 NPs in DPBS. The experiment was performed for three
different healthy donors. 2.15 Plasma Coagulation Time Assay Platelet-poor
plasma (PPP) was isolated from three different healthy donors. After
blood withdrawal, blood was centrifuged at 2500 g for 15 min. The upper layer of plasma was carefully transferred
into new tubes and centrifuged again at 2500 g for
15 min. PPP was carefully transferred to Protein LoBind tubes (Eppendorf),
aliquoted, and stored at −80 °C. PPP aliquots from three
different donors was thawed and pooled on the day of the experiment.
For the assay, 450 μL of pooled PPP was incubated with 50 μL
of uncoated or coated particle suspensions in DPBS at a total of 0.5
mg per condition. For the negative control, PPP was incubated with
50 μL of DPBS. Samples were incubated for 30 min at 37 °C,
after which the plasma-material mixture was carefully mixed 1:1 v/v
with recalcification buffer (10 mM HEPES, 20 mM CaCl 2 ,
115 mM NaCl, pH 7.2) in a well-plate. Absorbance was measured at 405
nm every minute for 45 min. All tubes and plates were previously blocked
with a 0.1 mg mL –1 albumin solution prior to contact
with plasma. Plasma coagulation time was determined by fitting
the kinetic curves to a Boltzmann model ( ) using Origin software, 2 where A 1 corresponds to the absorbance
baseline prior to coagulation; A 2 the absorbance plateau
after coagulation; and x 0 the center value, corresponding
to the estimated coagulation time, also reported as the “ripening
time” in previous reports. 2.16 Multiplexed Protein Assay Whole
blood treated with heparin (final concentration 1.5 IU ml-1) was incubated
with uncoated and coated PCL MPs, after which platelet-free plasma
was isolated and analyzed using a Luminex apparatus. Particle suspensions
in DPBS (0.8 mg in 100 μL) were added to 100 μL of DPBS
with calcium and magnesium, and 1 mL of whole blood. Samples were
incubated for at 37 °C under orbital shaking at 50 rpm in Biopur
tubes (Eppendorf) for 2 h. To stop the reaction, a solution of EDTA
was added up to a final concentration of 10 mM. Samples were centrifuged
at 1500 g for 10 min at room temperature, after which the supernatant
was transferred to new tubes and recentrifuged at 13000 g for 10 min.
The resulting platelet-free plasma was carefully transferred to cryotubes
and stored at −80 °C until analysis. 2.17 Statistical Analysis Normality tests
for the analysis of the mean fluorescence intensity of lipids were
performed through a Shapiro-Wilk test. Kruskal–Wallis ANOVA
with post hoc Dunn test, Mann–Whitney test
or one-way ANOVA with post hoc Tukey tests were performed
as indicated in the text. All analyses were performed using Origin
software. Statistical significance represented as n.s. (nonsignificant),
* ( p < 0.05), ** ( p < 0.01),
or *** ( p < 0.001). 

Materials Colloidal silica nanoparticles
(diameter 0.10 ± 0.03 μm) were purchased from Polysciences
Inc. DiagNano green fluorescent silica microparticles (diameter 5
μm), monodisperse, with hydroxyl surface functionalization,
were purchased from CD Bioparticles. Polycaprolactone (PCL) M w ∼ 14,000, Mowiol 40–88, HEPES
(≥99.5%), calcium chloride dihydrate (ACS reagent, ≥
99.0%), sodium chloride (ACS reagent, ≥ 99.0%) and ethylenediamine-tetraacetic
acid disodium salt dihydrate (EDTA, ≥ 99.0%) were purchased
from Sigma-Aldrich. 1,2-dioleoyl- sn -glycero-3-phosphocholine
(DOPC), cholesterol, egg sphingomyelin (SM), and soy phosphatidylinositol
(PI) were purchased from Avanti Polar Lipids (Sigma). Texas Red-DHPE
(TR-DHPE) was purchased from ThermoFisher Scientific. All lipids were
purchased in powder form, dissolved in chloroform, and stored at −20
°C except TR-DHPE, which was solubilized in methanol and then
stored at −20 °C. Dulbecco’s Phosphate Buffered
Saline (DPBS, sterile), DPBS with CaCl 2 and MgCl 2 (sterile), chloroform (ACS spectrophotometric grade, ≥ 99.8%),
dichloromethane (DCM, ACS reagent, ≥ 99.5%), methanol (HPLC
grade, ≥ 99.9%), and isopropanol (IPA, ACS reagent, ≥
99.5%) were purchased from Sigma-Aldrich. Ethanol (absolute, ≥
99.5%) was purchased from VWR Chemicals. Ultrapure water (resistivity
18.2 MΩ•cm) was obtained from a Milli-Q setup (Millipore).
DPBS was sterile-filtered with 0.2 μm syringe filters prior
to each experiment. 

OPSALC on SiO 2 Nanoparticles Suspensions of SiO 2 NPs were prepared in ethanol and
isopropanol at a concentration of 10 mg mL –1 . A
lipid solution of DOPC was prepared in the same solvents at concentrations
of 0.5, 1.0, 2.0, and 4.0 mg mL –1 . To coat the particles
through the OPSALC method, a concentrated lipid-particle suspension
was first prepared by adding 50 μL of SiO 2 NPs to
450 μL of lipid solution in a small glass vial. The gradient
was then induced by the injection of DPBS into the vial at a specific
addition rate (600 μL min –1 or 60 μL
min –1 ) using a syringe pump (KD Scientific). This
buffer addition was performed under magnetic stirring at 500 rpm,
up to a volume of 5 mL, reaching a final solvent:buffer ratio of 10:90
(v/v). The resulting suspension was then heated to 50 °C for
45 min and left to cool down to room temperature. Particles were washed
three times in ultrapure water by centrifuging at 10000 g for 10 min at 4 °C. During washing, the supernatant was removed
and the pellet was resuspended in fresh ultrapure water using a micropipette
(Eppendorf). After washing, particle pellets were kept at 4 °C
until further use. For the study on the effect of lipid concentration,
the lipid-particle mixture was prepared in ethanol using a buffer
addition rate of 600 μL min –1 and a lipid
concentration of 0.5, 1.0, 2.0, or 4.0 mg mL –1 ,
corresponding to an approximate lipid bilayer:particle ratio ( n ) of 4:1, 8:1, 16:1, and 32:1 ( Methods S1 ). For the study on buffer addition rate, the lipid-particle
mixture was prepared in ethanol using a lipid concentration of 1.0
mg mL –1 and a buffer addition rate of 600 or 60
μL min –1 . Lipid vesicles were produced as
the respective controls, under the same conditions, using pure ethanol
instead of the SiO 2 NP suspension. For the study on solvent,
the lipid-particle mixture was prepared in ethanol or isopropanol
using a lipid concentration of 1.0 mg mL –1 and a
buffer addition rate of 600 μL min –1 . Lipid
vesicles were produced as the respective controls, under the same
conditions, using pure ethanol or isopropanol instead of the SiO 2 NP suspension. Lastly, for the study on solvent:buffer ratio,
the lipid-particle mixture was prepared in ethanol using a lipid concentration
of 1.0 mg mL –1 and an addition rate of 600 μL
min –1 , at a final solvent:buffer ratio of 100:0,
50:50, or 10:90 (v/v). The final volume was maintained at 5 mL, and
the ratios were obtained by adding 4.5 mL of ethanol (100:0), 2 mL
of ethanol with 2.5 mL of DPBS (50:50), or 4.5 mL of DPBS (10:90). 

OPSALC on SiO 2 Microparticles Suspensions of SiO 2 MPs were prepared in ethanol and
isopropanol at a concentration of 10 mg mL –1 . A
fluorescent lipid solution of DOPC (99.9 mol % DOPC, 0.1 mol % TR-DHPE)
was prepared in the same solvents at concentrations of 0.25, 0.5,
1.0, and 2.0 mg mL –1 , corresponding to
0.1, 0.2, 0.4, and 0.8 mg per mg of SiO 2 MPs, or n ≈ 44, 88, 176, and 353 lipid bilayers per particle,
respectively ( Methods S2 ). To coat the
particles through the OPSALC method, a concentrated lipid-particle
suspension was prepared by mixing 40 μL of SiO 2 MPs
with 160 μL of lipid solution in a small glass vial. The gradient
was then induced by the injection of DPBS into the vial at a manual
pipetting rate (∼3 mL min –1 ) or at controlled
addition rates (600 μL min –1 or 60 μL
min –1 ) using a syringe pump (KD Scientific). The
buffer was added under magnetic stirring at 180 rpm up to a volume
of 2 mL, reaching a final solvent:buffer ratio of 10:90 (v/v). The
resulting suspension was then heated to 50 °C for 45 min and
left to cool down to room temperature. Particles were washed three
times in ultrapure water by centrifuging at 50 g for
4 min, and were kept at 4 °C until further use. The conditions
used during the studies on solvent, buffer addition rate, solvent:buffer
ratio, and lipid concentration are specified in detail in Tables S1–4 . 

Preparation of Erythrocyte Membrane-like Lipid
Formulations Based on lipidomics studies of the outer leaflet
of the erythrocyte membrane, we prepared
four lipid formulations of distinct biomimicry complexity. Fluorescent
and nonfluorescent versions of the same lipid formulation were prepared.
Fluorescent formulations included DT (DOPC, TR-DHPE), CT (DOPC, cholesterol,
TR-DHPE), ST (DOPC, cholesterol, sphingomyelin, TR-DHPE), and IT (DOPC,
cholesterol, sphingomyelin, phosphatidylinositol, TR-DHPE). Nonfluorescent
formulations (D, C, S, and I) included the same lipids at the same
lipid ratios, only without TR-DHPE. Detailed information on lipid
ratios can be found in Table S5 . 

OPSALC of SiO 2 MPs with Membrane-like
Formulations SiO 2 MPs were prepared at 10 mg mL –1 in isopropanol. Fluorescent lipid formulations DT,
CT, ST, and IT were prepared at 0.5 mg mL –1 in the
same solvent. A concentrated lipid-particle suspension was first made
by adding 80 μL of SiO 2 NPs to 320 μL of lipid
solutions in small glass vials. The gradient was then induced with
the addition of 3.6 mL of DPBS, after which the suspension was heated
to 50 °C for 45 min and left to cool down to room temperature.
Particles were washed three times in ultrapure water by centrifuging
at 50 g for 4 min, and finally redispersed in DPBS
for analysis. 

OPSALC of SiO 2 NPs with Membrane-like
Formulations SiO 2 NPs were prepared at 10 mg mL –1 in isopropanol. Nonfluorescent lipid formulations
D, C, S, and I were prepared at 1.0 mg mL –1 in the
same solvent. A concentrated lipid-particle suspension was first made
by adding 50 μL of SiO 2 NPs to 450 μL of lipid
solutions in small glass vials. The gradient was then induced with
the addition of 4.5 mL of DPBS, after which the suspension was heated
to 50 °C for 45 min and left to cool down to room temperature.
Particles were washed three times in ultrapure water by centrifuging
at 10000 g for 10 min at 4 °C and finally redispersed
in DPBS for analysis. 

OPSALC of PCL MPs with Membrane-like Formulations PCL MPs were produced through a single emulsion solvent evaporation
technique. A solution of 0.3% w/v Mowiol 40–88 (25 mL) was
prepared in water, to which a 3% w/v solution of PCL in dichloromethane
(1 mL) was added. The two-phase mixture was homogenized using tip
ultrasonication for 10 s on ice, with a pulse of 1 on 1 s off and
an amplitude of 25% (Fisherbrand Q500 Sonicator). The emulsion was
stirred at 600 rpm overnight to allow the dichloromethane to evaporate.
The resulting suspension was washed three times in ultrapure water
by centrifuging at 1500 g for 15 min at 4 °C,
removing the supernatant, and redispersing in water using a micropipette
(Eppendorf). For OPSALC, PCL particles were resuspended in the appropriate
amount of isopropanol to yield a concentration of ∼10 mg mL –1 . Similar to SiO 2 MPs, a concentrated lipid-particle
suspension was made by adding 80 μL of PCL MPs to 320 μL
of fluorescent lipid solution (DT, CT, ST, or IT, 1.0 mg mL –1 ) for coating analysis. As an alternative, nonfluorescent lipid solutions
(D, C, S, or I, 1.0 mg mL –1 ) were used for the preparation
of hemocompatibility assays. The gradient was then induced with the
addition of 3.6 mL DPBS while magnetically stirring at 500 rpm. The
suspension was further heated to 50 °C for 45 min and left to
cool down to room temperature. Particles were washed three times in
ultrapure water by centrifuging at 2500 g for 15 min at 4 °C
and finally redispersed in DPBS. 

Confocal Laser Scanning Microscopy Uncoated and OPSALC lipid-coated SiO 2 MP suspensions (0.2–0.4
mg mL –1 in DPBS) were imaged using a Zeiss LSM 880
confocal fluorescence microscope. A 20X air objective with a numeric
aperture of 0.8 and a 40X water immersion objective with a numeric
aperture of 1.2 were used. Image processing and analysis was performed
using Fiji software. Image acquisition
parameters (objective, acquisition speed, laser power, gain, and bit
depth) and processing methods were kept constant between all the conditions
in comparative studies to allow for a direct comparison. For solvent:buffer
ratio studies, ethanol and isopropanol conditions were acquired at
different laser powers and were therefore only compared qualitatively.
For lipid fluorescence intensity quantification, images were acquired
with the 20X objective at zoom 1.0 (n = 3 images per condition). To
determine the mean fluorescence intensity of TR-DHPE per particle,
an automatic threshold was applied to the red-channel image in order
to identify each particle as a region of interest (ROI). The mean
fluorescence intensity of each ROI was then measured, merged data
from a total of n = 3 images per condition, and plotted as the lipid
content per particle, per condition, using Origin software (OriginLab).
Z-stack images were acquired with a z -interval of
0.46 μm with the 40X objective. Orthogonal views of Z-stacks
were produced with Fiji software. Coating efficiency was determined
as the percentage of red-fluorescent particles (i.e., lipid-coated)
per green-fluorescent particles (i.e., all particles) within a specific
area of the sample. Calculations were performed for two sample areas
across two replicate batches, in a total of n = 557 particles. Where
indicated, images were equally adjusted for brightness (i.e., grayscale
range adjustment propagated for all images) for improved visualization. 

Epifluorescence Microscopy PCL MPs
were analyzed in an ECHO Revolve epifluorescence microscope (ECHO).
Images were acquired at 100% light intensity, with an exposure of
10 ms for transmission and 450 ms for Texas Red channels. Images were
equally adjusted for Texas Red intensity and merged with transmission
images using Fiji software. 

Dynamic Light Scattering (DLS) and Zeta Potential
(ZP) The size distribution and zeta potential of lipid-coated
SiO 2 NPs was determined using a ZetaSizer Lab with 90°
scatter angle measurements (Malvern). Three particle batches (n =
3) were produced, and each condition of each batch was measured in
5-fold in diluted DPBS (0.05X), at a particle concentration range
of 0.05–0.1 mg mL –1 . 

Cryo-transmission Electron Microscopy (cryo-TEM) Uncoated SiO 2 NPs, OPSALC lipid-coated SiO 2 NPs, and lipid vesicle suspensions (approximately 5 mg mL –1 in DPBS) were imaged using a FEI Tecnai T20 transmission electron
microscope operating at 200 keV. Suspensions were vitrified with liquid
ethane (Vitrobot, FEI) and examined on a Gatan cryo-stage. Images
were taken under low-dose conditions using a slow scan CCD camera.
For better visualization of the coating nanostructure, images were
adjusted for brightness and contrast. Particle and vesicle size were
quantified using FIJI software. Lipid
coating states were visually identified (“uncoated”,
“partially coated”, “coated”, “coated–lamellar”,
or “coated–vesicular”) and quantified for the
analysis of coating efficiency. 

Fluorescence Recovery after Photobleaching
(FRAP) SiO 2 MPs coated with fluorescent formulations
(0.2–0.4 mg mL –1 ) containing TR-DHPE were
subjected to FRAP measurements on a Nikon A1 confocal microscope (Nikon)
with a 20X objective (numerical aperture 0.75) in the Texas Red channel.
Circular ROIs on isolated particles with a diameter of 1 μm
(SiO 2 ) or 2 μm (PCL) were bleached for ∼2
s (SiO 2 ) or ∼100 ms (PCL) using the 488 and 561
nm LASERs simultaneously (SiO 2 ) or the 561 nm LASER only
(PCL), at maximum power. Reference ROIs, used for fading correction,
were always of equal dimensions as their respective bleach ROIs and
situated on a different particle. After bleaching, the bleach and
reference ROIs were imaged for up to 5 min to record fluorescence
recovery. Recovery curves are plotted as ratios of background corrected
intensities of bleached ROI/reference ROI as fading correction in
time. 

Blood Collection Human whole blood
was collected from healthy volunteers at the Blood Donor Service of
the Technical Medical Centre of University Twente and used within
4 h of withdrawal. The research did not fall within the scope of the
Dutch Medical Research Involving Human Subjects Act. Informed consent
was obtained from all volunteers and the used blood collection procedure
was approved by the Medical Research Ethics Committee (METC Twente,
reference K11–23). For hemolysis and plasma coagulation assays,
blood was collected in Vacutainer tubes (BD) containing anticoagulant
trisodium citrate with citric acid and dextrose (ACD-A) and treated
in the first hour after withdrawal. For multiplex assays, blood was
collected in Greiner tubes without additive and immediately anticoagulated
with sodium heparin (LEO) to a final concentration of 1.5 IU ml –1 . 

Hemolysis Assays Erythrocytes (here
abbreviated to RBCs, from Red Blood Cells) were isolated based on
a protocol described elsewhere. Briefly,
anticoagulated whole blood was first centrifuged at 300 g for 2 min, after which the position of the upper layers of plasma
and buffy coat were marked on the tube. Both upper layers were removed,
substituted with an equal volume of DPBS, and used to redisperse RBCs.
Cells were washed three times with DPBS using the same centrifugation
conditions and redispersed in the same volume of DPBS. This suspension
was then diluted 50 times in the same buffer. For hemolysis
assays, 950 μL of diluted RBC suspensions were incubated with
50 μL of a 4 mg mL –1 suspension of uncoated
or coated SiO 2 NPs, at a total particle concentration of
approximately 0.2 mg mL –1 . For the negative controls,
RBCs were incubated with 50 μL of DPBS, and for positive controls,
cells were incubated with 50 μL of a Triton-X solution. Particle
suspensions at the same concentration in pure DPBS were included as
controls for absorbance correction. All samples were incubated at
37 °C for 4 h, with gentle mechanical agitation once every hour,
and then centrifuged at 300 g for 5 min. The supernatant
was transferred to a well-plate for quantification of released hemoglobin
through absorbance measurements at a wavelength of 540 nm (150 μL,
n = 5 measurements per condition). Hemolysis rates (HR) were calculated
according to , 1 where A S,RBCs is the absorbance
of the supernatant of each condition of SiO 2 NPs incubated
with RBCs; < A S,DPBS > is the mean absorbance of the
supernatant of the same SiO 2 NP condition in DPBS; <
A UC, RBCs > is the mean absorbance of the supernatant
of uncoated SiO 2 NPs incubated with RBCs; and < A UC,DPBS > in the mean absorbance of the supernatant of uncoated
SiO 2 NPs in DPBS. The experiment was performed for three
different healthy donors. 

Plasma Coagulation Time Assay Platelet-poor
plasma (PPP) was isolated from three different healthy donors. After
blood withdrawal, blood was centrifuged at 2500 g for 15 min. The upper layer of plasma was carefully transferred
into new tubes and centrifuged again at 2500 g for
15 min. PPP was carefully transferred to Protein LoBind tubes (Eppendorf),
aliquoted, and stored at −80 °C. PPP aliquots from three
different donors was thawed and pooled on the day of the experiment.
For the assay, 450 μL of pooled PPP was incubated with 50 μL
of uncoated or coated particle suspensions in DPBS at a total of 0.5
mg per condition. For the negative control, PPP was incubated with
50 μL of DPBS. Samples were incubated for 30 min at 37 °C,
after which the plasma-material mixture was carefully mixed 1:1 v/v
with recalcification buffer (10 mM HEPES, 20 mM CaCl 2 ,
115 mM NaCl, pH 7.2) in a well-plate. Absorbance was measured at 405
nm every minute for 45 min. All tubes and plates were previously blocked
with a 0.1 mg mL –1 albumin solution prior to contact
with plasma. Plasma coagulation time was determined by fitting
the kinetic curves to a Boltzmann model ( ) using Origin software, 2 where A 1 corresponds to the absorbance
baseline prior to coagulation; A 2 the absorbance plateau
after coagulation; and x 0 the center value, corresponding
to the estimated coagulation time, also reported as the “ripening
time” in previous reports. 

Multiplexed Protein Assay Whole
blood treated with heparin (final concentration 1.5 IU ml-1) was incubated
with uncoated and coated PCL MPs, after which platelet-free plasma
was isolated and analyzed using a Luminex apparatus. Particle suspensions
in DPBS (0.8 mg in 100 μL) were added to 100 μL of DPBS
with calcium and magnesium, and 1 mL of whole blood. Samples were
incubated for at 37 °C under orbital shaking at 50 rpm in Biopur
tubes (Eppendorf) for 2 h. To stop the reaction, a solution of EDTA
was added up to a final concentration of 10 mM. Samples were centrifuged
at 1500 g for 10 min at room temperature, after which the supernatant
was transferred to new tubes and recentrifuged at 13000 g for 10 min.
The resulting platelet-free plasma was carefully transferred to cryotubes
and stored at −80 °C until analysis. 

Statistical Analysis Normality tests
for the analysis of the mean fluorescence intensity of lipids were
performed through a Shapiro-Wilk test. Kruskal–Wallis ANOVA
with post hoc Dunn test, Mann–Whitney test
or one-way ANOVA with post hoc Tukey tests were performed
as indicated in the text. All analyses were performed using Origin
software. Statistical significance represented as n.s. (nonsignificant),
* ( p < 0.05), ** ( p < 0.01),
or *** ( p < 0.001). 

Results and Discussion 3.1 Effect of Lipid Concentration on SiO 2 NP Lipid Coating To demonstrate the feasibility of the
OPSALC method, as well as to allow for its optimization, SiO 2 NPs were first used. These nanoparticles were selected as the standard
substrate due to their surface chemistry, low polydispersity, and
small size, which make them an excellent platform for particle colloidal
stability studies and characterization of the lipid coating. We studied the effect of lipid concentration on coating formation
using lipid solutions of 0.5, 1.0, 2.0, and 4.0 mg mL –1 , performing dynamic light scattering (DLS) measurements to assess
colloidal stability. These lipid concentrations correspond to a mathematical
estimation of the number of concentrically stacked lipid bilayers
around a single particle, also referred to as lipid bilayer:particle
ratios, of 4:1, 8:1, 16:1, and 32:1 ( a, Methods S1 ). Uncoated particles showed a main size intensity
peak around 158
nm in DLS studies, whereas SiO 2 NPs after coating showed
main peaks in the range of 161–216 nm ( Figure S2 , Table S6 ). These size
shifts could indicate the formation of a lipid coating, but also the
formation of aggregates. Among coated samples, the increase in lipid
concentration from lipid bilayer:particle ratios of 4:1 to 8:1 significantly
improved colloidal stability, as observed from the lower polydispersity
index (PDI) of the particle suspensions in different batches ( Figure S3 , Tables S6,7 ). A further increase in lipid concentration (16:1 and 32:1) did
not produce a further improvement in all batches. To determine coating
efficiency, cryogenic transmission electron microscopy (cryo-TEM)
images were recorded and analyzed. Conditions 8:1 and 32:1 produced
the highest percentage of fully coated particles (32% and 31%, respectively, Figure S4 ). For all fully coated particles, cryo-TEM
data revealed a single uniform lipid layer on the particles ( b–f), confirming
that the coating process did not produce multilayers. The observation
of single layers also corroborates previous conclusions on solvent
exchange methods for planar surfaces. , Overall, the
most consistent condition in terms of low polydispersity and high
coating efficiency was 8:1, with the lowest PDI (mean of 0.146), lowest
Z-average (mean of 180.1 nm), and highest coating efficiency ( Tables S6,7 , Figures S2–4 ). This condition (8:1), corresponding to 1.0 mg mL –1 of lipid solution, was used in all further SiO 2 NP experiments. 3.2 Effect of Buffer Addition Rate on SiO 2 NP Lipid Coating Next, we assessed the effect of
buffer addition rate on coating formation. We compared the particle
size distributions obtained from an addition rate of 600 μL
min –1 or a rate of 60 μL min –1 , and observed that the lower rate produced a marginally broader
SiO 2 NP size distribution with some large aggregates ( g–i). To better
understand how lipids assemble with the different buffer addition
rates, we also analyzed the lipid vesicles formed in both conditions,
in the absence of particles. Similar to the coated particle size distributions,
lipid vesicles produced at 60 μL min –1 revealed
a broader size distribution, whereas 600 μL min –1 produced more monodisperse populations ( Figure S5a,b , Table S8 ). To gain
a better understanding of these results, we imaged the coated particles
and lipid vesicles using cryo-TEM. Both buffer addition rates produced
single-layer lipid coatings ( j,k); however, the coating efficiency was higher at
600 μL min –1 as compared to 60 μL min –1 (38% vs. 20% fully coated particles, l). In line with the DLS observations,
we also confirmed that lipid vesicles produced at 600 μL min –1 were smaller and less broad in size distribution
than those produced at 60 μL min –1 , with main
size peaks at ∼40 nm and ∼60 nm, respectively ( Figure S5c–e ). Here, it is worth to note
that DLS sizes tend to be larger than cryo-TEM sizes due to the larger
hydrodynamic radius observed in DLS and the presence of clusters in
suspension; therefore, cryo-TEM data was preferred for vesicle size
analysis. The faster addition of water is thought to have induced
a faster reduction of the critical micelle concentration of lipids,
possibly preventing the system from fully reaching a state of thermodynamic
equilibrium. This effect could have likely led to a larger fraction
of lipid monomers and smaller lipid assemblies, such as micelles,
as compared to larger lipid vesicles, thereby favoring the pathway
of lipid-layer deposition during the process ( a(i)). This effect would also explain the
smaller size of lipid vesicles observed at the faster addition rate.
This is in agreement with earlier reports that indicate that lipid
layers are predominantly formed from adsorbed monomers and micelles. , , Overall, this hypothesis could
explain the higher coating efficiency observed at 600 μL min –1 , and therefore we used this buffer addition rate
in all further experiments. 3.3 Effect of Solvent on SiO 2 NP Lipid
Coating Next, we explored the effect of solvent on the formation
of a lipid coating by comparing ethanol and IPA buffer gradients.
Ethanol was used as solvent in the previous sections, as it represented
a good solvent for dispersing SiO 2 NPs with no aggregation.
However, different solvents are known to change the fractions of monomers
and lipid assemblies upon buffer addition, influencing the efficiency
of the coating process. Based on the differences in dielectric constants
for IPA and ethanol, DOPC is expected to result in a larger fraction
of lipid monomers and small lipid assemblies in IPA, boosting the formation of lipid bilayers, as shown in previous
studies on planar surfaces. DLS
measurements showed that SiO 2 NPs were more stable in ethanol
than IPA buffer gradients ( Figure S6a,b ). However, lipid-coated SiO 2 NPs tended to have a smaller
average diameter in IPA than in ethanol (201 nm vs 223 nm, Table S9 ) as well as a lower occurrence of aggregates
( m,n). Lipid-coated
particles were also analyzed using cryo-TEM, revealing the formation
of single-layer coatings on the fully coated particles ( Figure S6e , d). A more in-depth analysis revealed that
the fraction of fully coated SiO 2 NPs was lower for IPA
(18%, vide infra n) than for ethanol (38%, l). In contrast, partially
coated SiO 2 NPs were more frequent in IPA (26%) than in
ethanol (20%). The complete formation of a lipid layer in IPA gradients
appeared to be slowed down, despite the higher available fraction
of lipid monomers and small assemblies. This effect contradicts earlier
observations for IPA exchange coatings on planar surfaces, suggesting that other effects (e.g., lipid–surface,
lipid–solvent interactions) may have played a role in coating
formation. On the other hand, DLS analysis of lipid vesicles revealed
slightly smaller vesicles for IPA (136 nm) than for ethanol (149 nm, Figure S6c,d,f,g , Table S9 ). This difference in vesicle size is much less pronounced for the
different solvents than observed for different buffer addition rates,
where vesicle diameters were 97 nm for 600 μL min –1 and 212 nm for 60 μL min –1 ( Table S8 ). This correlates well with the smaller
differences in the quality of coated SiO 2 NPs comparing
IPA and ethanol, while a change in buffer addition rate from 60 μL
min –1 to 600 μL min –1 has
a much larger effect on the quality of the coating process. 3.4 Effect of Solvent:Buffer Ratio on SiO 2 NP Lipid Coating Subsequently, the effect of solvent:buffer
ratio on the lipid-coating of SiO 2 NPs was studied. We
imaged SiO 2 NPs using cryo-TEM after halting the buffer
addition process at an ethanol:buffer ratio of 50:50 and 10:90 ( o,p). For 50:50,
14% of SiO 2 NPs presented a vesicular coating and 25% presented
a full lamellar (i.e., single-layer) coating ( q), whereas for 10:90 no vesicular coating
was observed and 38% of SiO 2 NPs presented a full lamellar
coating ( l).
These cryo-TEM results show that the vesicles in the bulk are attracted
to the surface of SiO 2 NPs to some extent, suggesting a
small contribution of vesicle rupture to the formation of lamellar
coatings ( a(ii)).
However, at a final solvent to buffer ratio of 10:90, no absorbed
vesicles could be observed, which implies that vesicles play a minor
role in the coating process. Furthermore, halting the buffer addition
at 50:50 will allow the system time to reach equilibrium, therefore
allowing a larger fraction of lipids to assemble into micelles and
eventually vesicles, as compared to the uninterrupted buffer addition
up to 10:90. Taking into account the earlier solvent-exchange studies
on planar surfaces, , reporting minor to no contribution
of vesicle rupture to the coating formation during buffer exchange,
vesicles present in the 50:50 case could have also adsorbed to the
uncoated parts of a particle surface without necessarily participating
in coating formation. These hypotheses, however, warrant a more in-depth
biophysical study to unveil the exact contribution of vesicles to
coating formation. Lastly, it is important to address the significant
fraction of uncoated (∼40%) and partially coated (∼20%)
SiO 2 NPs observed in cryo-TEM. This phenomenon represents
a common issue in the lipid-coating of nanoparticles, and has been
similarly reported elsewhere using vesicle fusion techniques. , It is our understanding that the natural process of lipid–surface
interactions occur at the micelle/vesicle state, and that the natural
curvature of nanoparticles reduces the extent of micelle/vesicle adsorption,
thus limiting the occurrence of vesicle fusion and lipid bilayer formation.
In contrast with other methods, however, the coating efficiency of
OPSALC has potential for significant improvement. For example, the
use of different lipids could potentially improve lipid–surface
attraction and may facilitate lipid packing within the layer. In addition,
the use of considerably higher lipid excess to particle surface area
should improve the fraction of fully coated particles, as we used
5 × 10 –17 mg mL –1 nm –2 while earlier studies on solvent assisted lipid coating
of planar surfaces have used 1 × 10 –15 mg mL –1 nm –2 . 3.5 Applying OPSALC to Coat SiO 2 Microparticles Subsequently, we attempted to lipid-coat SiO 2 MPs using
the OPSALC procedure. Similar to coating the nanoparticles, we found
that the type of solvent, the rate of addition of buffer, solvent:buffer
ratio and lipid concentration were determining our success of achieving
a homogeneous lipid coverage on SiO 2 MPs ( ). Confocal microscopy analysis
revealed that the coating efficiency for SiO 2 MPs was 99.7%
( Figure S7 ). A higher lipid fluorescence
per particle was observed for IPA than for ethanol gradients ( a–d), indicating
coatings with a higher lipid content in the case of IPA. This higher
lipid fluorescence could indicate a denser or thicker coating, or
a better surface coverage, but could also be rationalized with more
lipids penetrating the pores of the particles. Homogeneous coatings
were found for all tested buffer addition rates using ethanol:buffer
gradients (>3 mL min –1 , 600 μL min –1 , 60 μL min –1 , e–g). The analysis of
fluorescence
intensity plot profiles of coated particles revealed different degrees
of lipid penetration into the pores of the particles depending on
the addition rate, with the lowest condition (60 μL min –1 ) yielding the highest degree of penetration ( Figure S8 ). Similarly, upon increasing lipid
concentration, an increase of lipid penetration into the particles
was observed ( Figure S11 ). Concentrations
of 0.25, 0.5, 1.0, and 2.0 mg mL –1 were used, which
corresponded to 0.1, 0.2, 0.4, and 0.8 mg per mg of SiO 2 MPs and n ≈ 44, 88, 176, and 353 lipid bilayers per particle,
respectively ( Method S2 ). Furthermore, the amount and
homogeneity of the coating were substantially different when halting
the buffer addition at solvent:buffer ratios of 100:0, 75:25, 50:50,
25:75 and 10:90 for both ethanol and IPA. In the case of ethanol,
high intensity (i.e., high lipid content) and homogeneous coatings
were observed at buffer contents of ≥75%, whereas at lower
buffer contents (<75%) coatings were less homogeneous, showing
few high intensity (i.e., high lipid content) features distributed
throughout a homogeneous low intensity (low lipid content) coating.
( i–m, Figures S9,10 ). It should be noted that lipid penetration increased,
not only with
a decrease in buffer addition rate and higher lipid concentration,
but also with the use of IPA instead of ethanol, an effect clearly
visible for 25:75 solvent:buffer ratios ( Figures S10,11 , h, red circles). Lower buffer addition rates are believed to provide
more time for lipid monomers and small lipid assemblies to diffuse
into the pores of the particles, whereas higher lipid concentrations
and the use of IPA supply higher amounts of lipid monomers and smaller
assemblies, which can penetrate the 8–10 nm sized pores. Therefore,
we tentatively assign the increased lipid penetration to (i) the availability
of higher fractions of lipid monomers and small lipid assemblies,
and (ii) the longer time allowed for lipids to diffuse into the pores
of the particles. Altogether, we concluded that the optimal
conditions for OPSALC
to yield a homogeneous coating on SiO 2 MPs (with limited
penetration) are a moderate excess of lipids (n ≈ 44 lipid
bilayers per particle), a minimum buffer addition rate of 600 μL
min –1 , and a final solvent:buffer ratio of 10:90
using either ethanol or IPA as the solvent. 3.6 Erythrocyte Membrane-like Lipid Coatings on
SiO 2 MPs Next, we explored the use of OPSALC in
the development of erythrocyte membrane-like coatings on SiO 2 MPs. Lipidomics studies of the erythrocyte membrane revealed that
its outer leaflet, the one in direct contact with blood, has a main
composition of phosphatidylcholines (42.3 mol %), sphingomyelins (55.3
mol %), phosphatidylserine (1.5 mol %), and phosphatidylinositol (0.9
mol %), with a varying concentration
of cholesterol (30–40 mol %). Based
on these reports, we selected a phosphatidylcholine lipid (DOPC),
a sphingomyelin lipid (egg SM, mainly 16:0 fatty acid tails), a phosphatidylinositol
lipid (soy PI, mainly 18:1 fatty acid tails), and cholesterol (Chol)
as the components of the cell membrane-like formulations. Phosphatidylserine,
although present in the outer leaflet of the membrane, is a typical
biomarker of cell aging and certain pathologies , and was therefore excluded from the cell membrane design. The different
lipids were combined into formulations, according to the lipidomics-derived
molar ratios: DOPC only (D), DOPC and Chol (C), DOPC, Chol and egg
SM (S); and DOPC, Chol, egg SM and soy PI (I) ( a). For microscopy, fluorescent versions of these formulations
(DT, CT, ST, IT, respectively) were prepared by adding 0.1 mol % of
TR-DHPE. As expected, all fluorescent erythrocyte membrane-like
formulations
yielded homogeneous lipid coatings on SiO 2 MPs using the
OPSALC method ( b,c), retaining at least 70% of initial fluorescence intensity for
1 week ( f, Figure S12 ). Subsequently, measurements of fluorescence
recovery after photobleaching (FRAP) clearly distinguished lipid mobility
in the different coatings on SiO 2 MPs ( d,e). For DT and CT coatings,
nearly identical recovery kinetics were found, with higher lipid mobilities
than for ST and IT coatings, which showed near identical recovery
kinetics among themselves as well. Obviously, FRAP experiments on
microparticles, wherein the bleached area constitutes a significant
fraction of the total particle surface area, will show incomplete
recovery. The differences in mobility observed in FRAP could be explained
by differences in lipid phases formed by the different lipid formulations.
Miscibility diagrams of DOPC and cholesterol show that D and C coatings
are expected to be in a liquid-disordered phase at room temperature. In contrast, egg SM has a much higher chain
melting temperature (38 °C) than DOPC (−17 °C) and
causes mixtures of DOPC, Chol and egg SM, like formulation S and I,
to be in the two-liquid phase, containing liquid-disordered and liquid-ordered
domains at room temperature. It has been
shown that liquid-ordered phases have significantly lower lipid mobility
than liquid-disordered ones, corroborating our findings that D and
C coatings are more mobile than S and I coatings. Furthermore, it
stands to reason that lipid mixtures (i.e, formulations S and I),
which result in partially liquid-ordered layers due to their tighter
packing and stronger lipid–lipid attraction, are more prone
to form larger, more stable lipid assemblies during OPSALC. This would
also mean that lower amounts of lipid monomers and small lipid assemblies
are present, limiting the possibility of penetration of lipids into
the pores of the particles. This observation, in turn, is in agreement
with the smaller penetration depth for S and I, as effectively observed
in confocal microscopy ( b,c). The observed differences show the potential of using
OPSALC to tailor the penetration depth by varying not only buffer
addition rate, lipid concentration and choice of solvent, but also
by varying the phase of the lipid layers. Overall, the OPSALC
method was used to successfully coat SiO 2 MPs with various
erythrocyte membrane-mimicking lipid coatings,
with distinct characteristics depending on the exact formulation applied. 3.7 Erythrocyte Membrane-like Coatings on SiO 2 NPs To confirm that erythrocyte membrane-like formulations
could also be applied to nanomaterials, we studied the efficiency
of OPSALC on SiO 2 NPs. Particles were coated through IPA:buffer
gradients using formulations D, C, S, and I, and subsequently analyzed
using DLS and zeta potential (ZP) measurements. Formulations D and
C resulted in a larger particle size distribution than S and I ( g) and the polydispersity
decreased linearly going from D, C, S to I ( h). All coatings produced a reduction in
the ZP, from −48 mV in uncoated particles to approximately
−30 mV in coated ones ( i). Among the tested conditions, particles coated with
formulation I produced the lowest ZP value (−34.4 mV) and the
lowest polydispersity index (0.26), thus displaying the highest colloidal
stability for lipid-coated SiO 2 NPs. To further characterize
the coatings, SiO 2 NPs were analyzed with cryo-TEM. All
formulations successfully produced single-layer coatings on the particles
( j–n),
corroborating our previous cryo-TEM results. Coating efficiency was
further quantified as percentage of uncoated, partially coated, and
coated particles, revealing that formulations S and I generated a
considerably higher percentage of fully coated particles than formulations
D and C (D: 18%, C: 18%, S: 50%, I: 41% o). Previously, we had proposed that lipid–surface
and lipid–medium interactions were the main factor determining
the efficiency of OPSALC. Interestingly, we have now observed that
lipid–lipid interactions also play a major role in coating
efficiency, as formulations with tighter lipid packing (S, I) produce
a considerably higher percentage of coated particles. Presumably,
the differences in packing and hydration of the partially liquid-ordered
layers improved lipid–surface interactions, compensating for
the expected occurrence of lower amounts of monomers and small lipid
assemblies. Overall, we confirmed that SiO 2 NPs can
be coated with erythrocyte membrane-like formulations using the OPSALC
method, and observed that coating SiO 2 NPs with formulations
producing liquid-ordered and -disordered phases (S, I) was more effective
than coating them with formulations producing a liquid-disordered
phase only (D, C). 3.8 Erythrocyte Membrane-like Coatings on PCL
MPs Next, we attempted to coat hydrophobic PCL MPs using
the OPSALC method. PCL was chosen as a hydrophobic particle substrate
due to its bioresorbable composition and its common use in drug delivery
and tissue engineering. It is often preferred over other hydrophobic
polymers (e.g., PLA or PLGA) due to its low cost, high availability,
and lower rate of biodegradation. , Here, the
fluorescent erythrocyte membrane-like formulations used for SiO 2 MPs were also employed for coating visualization studies. During the coating process, we observed that coated microparticles
were easier to redisperse in water than uncoated microparticles, and
that S- and I-coated PCL MPs were significantly easier to redisperse
than D- and C-coated PCL MPs ( p). Fluorescence imaging revealed the highest fluorescence
intensity for I-coated PCL MPs, with C- and S-coated particles of
only slightly lower intensity, and a significantly lower intensity
for D-coated PCL MPs ( q). FRAP measurements revealed slow recovery rates, comparable
for all types of coating ( r,s), indicating that all lipid layers had limited
mobility. We propose that this reduced mobility is due to hydrophobic
interactions between the fatty acid tails of the lipids and the surface
of PCL particles, which may have resulted in a single immobile monolayer,
as suggested elsewhere for another planar hydrophobic substrate. Altogether, these results indicate that OPSALC
can be employed to successfully coat hydrophobic PCL MPs with different
mixtures of lipids, resulting in layers with low mobility irrespective
of lipid composition. 3.9 Hemocompatibility Testing of Lipid-Coated
Particles To demonstrate the functionality of distinct lipid
coatings, the effect of lipid formulation on particle hemocompatibility
was investigated. We first conducted hemolysis and plasma coagulation
assays on SiO 2 NPs, as they are known for their highly
hemolytic and procoagulant nature. , Hemolysis
assays showed that uncoated SiO 2 NPs are extremely hemolytic
at particle concentrations as low as 0.2 mg mL –1 , and specifically more hemolytic than the positive control (1% v/v
Triton-X final concentration), which led to its use as the reference
for hemolysis rate calculation. In contrast, SiO 2 NPs coated
with any of the lipid formulations yielded a drastic reduction of
erythrocyte damage, coupled to a reduction in the amount of hemoglobin
release and the resulting hemolysis rates (from 100% to below 5%, a,b). Moreover, no
significant difference was observed between lipid formulations and
the negative control, confirming its inert behavior. It must be noted,
however, that the particle aggregation observed for D- and C-coated
SiO 2 NPs ( g) could be partially responsible for the reduction in hemolysis
rate. As a result, the low hemolysis rates of coatings S and I are
more reliable than the hemolysis rates of coatings D and C, as they
more accurately indicate the hemoprotective effect of the lipid layer
on single particles. To determine whether the lipid composition of the
particle coatings
have an effect on plasma coagulation time, coated and uncoated SiO 2 NPs were incubated with platelet-poor plasma, recalcified,
and analyzed via the increase in scattering during fibrin clot formation.
SiO 2 surfaces are known for their procoagulant properties
through the surface-induced activation of factor XII, or Hageman factor,
an important mediator of the intrinsic coagulation pathway. , Plasma coagulation curves demonstrated that S and I coatings could
induce a more pronounced delay in coagulation onset than D and C coatings
( c), demonstrating
a more significant attenuation of the pro-coagulating properties of
SiO 2 NPs, although this advantage was not consistently
observed. The coagulation curves for two other tested batches can
be found in Figure S13 . Plasma coagulation
time was then calculated as average of all batches tested. Overall,
the average coagulation time induced by different coatings was not
significantly different; however, a significant difference was observed
between the uncoated and all coated conditions ( d). Collectively, these results indicate
that all tested coatings were effective in attenuating the coagulating
effect of SiO 2 NPs. To further study the hemocompatibility
effects of the lipid coatings,
we explored the complement activation, platelet activation, leukocyte
recruitment, and inflammation response induced by uncoated vs lipid-coated
PCL MPs. Despite the low toxicity of this polymer, hydrophobic surfaces
are known to adsorb fibrinogen, IgG, and C3, proteins involved in
platelet/leukocyte adhesion and complement activation. , Since hydrophobic micromaterials have been researched as drug carriers, , porous functional scaffolds, and, more
recently, oxygen-generating components of tissue engineered implants, , hemocompatibility has become a key aspect of their biocompatibility. Indeed, PCL MPs stimulated the production of all complement activation,
platelet activation, and leukocyte recruitment markers when compared
to nonstimulated blood ( e). The relatively high levels of C5a, a common biomarker
for all complement activation pathways, indicated that PCL MPs tend to induce complement activation to some
extent. Lipid coatings, however, attenuated this complement-activating
effect with an interesting trend: increasingly complex lipid formulations
produced a gradually safer response. Neutrophil chemotactic marker
IL-8 followed a similar trend, with more
complex lipid coatings resulting in lower IL-8 production. Platelet
activation markers (β-TG and PDGF-BB), , platelet-secreted monocyte and lymphocyte attractant RANTES, and monocyte attractant MCP-1 were found at generally low levels for all coating formulations.
Finally, activated T-cell cytokine MIF was found at similarly high levels in uncoated, D-coated and C-coated
PCL MPs, but found at lower levels in S-coated and I-coated PCL MPs.
These results suggested that PCL MPs were not entirely bioinert in
blood, and that lipid coatings with more complex erythrocyte membrane
mimicry were able to notably improve PCL MP hemocompatibility. To
expand on these results, the levels of inflammatory biomolecules (IL-1RA,
IL-1β, IL-6, and TNF-α) were also analyzed. While there
was no statistically significant difference between the negative control
and any PCL MPs for any of these biomarkers, there was a consistent
tendency for D-coated PCL MPs to have an almost statistical difference
from the negative controls ( f–i), indicating a slightly more pro-inflammatory
profile than uncoated and other coated particles. Ultimately,
all results corroborated that S and I lipid formulations
generated a better improvement on blood-contacting particle hemocompatibility
as compared to D and C lipid formulations, indicating a potential
benefit in using biomimetically complex lipid coatings in biomedical
applications. 

Effect of Lipid Concentration on SiO 2 NP Lipid Coating To demonstrate the feasibility of the
OPSALC method, as well as to allow for its optimization, SiO 2 NPs were first used. These nanoparticles were selected as the standard
substrate due to their surface chemistry, low polydispersity, and
small size, which make them an excellent platform for particle colloidal
stability studies and characterization of the lipid coating. We studied the effect of lipid concentration on coating formation
using lipid solutions of 0.5, 1.0, 2.0, and 4.0 mg mL –1 , performing dynamic light scattering (DLS) measurements to assess
colloidal stability. These lipid concentrations correspond to a mathematical
estimation of the number of concentrically stacked lipid bilayers
around a single particle, also referred to as lipid bilayer:particle
ratios, of 4:1, 8:1, 16:1, and 32:1 ( a, Methods S1 ). Uncoated particles showed a main size intensity
peak around 158
nm in DLS studies, whereas SiO 2 NPs after coating showed
main peaks in the range of 161–216 nm ( Figure S2 , Table S6 ). These size
shifts could indicate the formation of a lipid coating, but also the
formation of aggregates. Among coated samples, the increase in lipid
concentration from lipid bilayer:particle ratios of 4:1 to 8:1 significantly
improved colloidal stability, as observed from the lower polydispersity
index (PDI) of the particle suspensions in different batches ( Figure S3 , Tables S6,7 ). A further increase in lipid concentration (16:1 and 32:1) did
not produce a further improvement in all batches. To determine coating
efficiency, cryogenic transmission electron microscopy (cryo-TEM)
images were recorded and analyzed. Conditions 8:1 and 32:1 produced
the highest percentage of fully coated particles (32% and 31%, respectively, Figure S4 ). For all fully coated particles, cryo-TEM
data revealed a single uniform lipid layer on the particles ( b–f), confirming
that the coating process did not produce multilayers. The observation
of single layers also corroborates previous conclusions on solvent
exchange methods for planar surfaces. , Overall, the
most consistent condition in terms of low polydispersity and high
coating efficiency was 8:1, with the lowest PDI (mean of 0.146), lowest
Z-average (mean of 180.1 nm), and highest coating efficiency ( Tables S6,7 , Figures S2–4 ). This condition (8:1), corresponding to 1.0 mg mL –1 of lipid solution, was used in all further SiO 2 NP experiments. 

Effect of Buffer Addition Rate on SiO 2 NP Lipid Coating Next, we assessed the effect of
buffer addition rate on coating formation. We compared the particle
size distributions obtained from an addition rate of 600 μL
min –1 or a rate of 60 μL min –1 , and observed that the lower rate produced a marginally broader
SiO 2 NP size distribution with some large aggregates ( g–i). To better
understand how lipids assemble with the different buffer addition
rates, we also analyzed the lipid vesicles formed in both conditions,
in the absence of particles. Similar to the coated particle size distributions,
lipid vesicles produced at 60 μL min –1 revealed
a broader size distribution, whereas 600 μL min –1 produced more monodisperse populations ( Figure S5a,b , Table S8 ). To gain
a better understanding of these results, we imaged the coated particles
and lipid vesicles using cryo-TEM. Both buffer addition rates produced
single-layer lipid coatings ( j,k); however, the coating efficiency was higher at
600 μL min –1 as compared to 60 μL min –1 (38% vs. 20% fully coated particles, l). In line with the DLS observations,
we also confirmed that lipid vesicles produced at 600 μL min –1 were smaller and less broad in size distribution
than those produced at 60 μL min –1 , with main
size peaks at ∼40 nm and ∼60 nm, respectively ( Figure S5c–e ). Here, it is worth to note
that DLS sizes tend to be larger than cryo-TEM sizes due to the larger
hydrodynamic radius observed in DLS and the presence of clusters in
suspension; therefore, cryo-TEM data was preferred for vesicle size
analysis. The faster addition of water is thought to have induced
a faster reduction of the critical micelle concentration of lipids,
possibly preventing the system from fully reaching a state of thermodynamic
equilibrium. This effect could have likely led to a larger fraction
of lipid monomers and smaller lipid assemblies, such as micelles,
as compared to larger lipid vesicles, thereby favoring the pathway
of lipid-layer deposition during the process ( a(i)). This effect would also explain the
smaller size of lipid vesicles observed at the faster addition rate.
This is in agreement with earlier reports that indicate that lipid
layers are predominantly formed from adsorbed monomers and micelles. , , Overall, this hypothesis could
explain the higher coating efficiency observed at 600 μL min –1 , and therefore we used this buffer addition rate
in all further experiments. 

Effect of Solvent on SiO 2 NP Lipid
Coating Next, we explored the effect of solvent on the formation
of a lipid coating by comparing ethanol and IPA buffer gradients.
Ethanol was used as solvent in the previous sections, as it represented
a good solvent for dispersing SiO 2 NPs with no aggregation.
However, different solvents are known to change the fractions of monomers
and lipid assemblies upon buffer addition, influencing the efficiency
of the coating process. Based on the differences in dielectric constants
for IPA and ethanol, DOPC is expected to result in a larger fraction
of lipid monomers and small lipid assemblies in IPA, boosting the formation of lipid bilayers, as shown in previous
studies on planar surfaces. DLS
measurements showed that SiO 2 NPs were more stable in ethanol
than IPA buffer gradients ( Figure S6a,b ). However, lipid-coated SiO 2 NPs tended to have a smaller
average diameter in IPA than in ethanol (201 nm vs 223 nm, Table S9 ) as well as a lower occurrence of aggregates
( m,n). Lipid-coated
particles were also analyzed using cryo-TEM, revealing the formation
of single-layer coatings on the fully coated particles ( Figure S6e , d). A more in-depth analysis revealed that
the fraction of fully coated SiO 2 NPs was lower for IPA
(18%, vide infra n) than for ethanol (38%, l). In contrast, partially
coated SiO 2 NPs were more frequent in IPA (26%) than in
ethanol (20%). The complete formation of a lipid layer in IPA gradients
appeared to be slowed down, despite the higher available fraction
of lipid monomers and small assemblies. This effect contradicts earlier
observations for IPA exchange coatings on planar surfaces, suggesting that other effects (e.g., lipid–surface,
lipid–solvent interactions) may have played a role in coating
formation. On the other hand, DLS analysis of lipid vesicles revealed
slightly smaller vesicles for IPA (136 nm) than for ethanol (149 nm, Figure S6c,d,f,g , Table S9 ). This difference in vesicle size is much less pronounced for the
different solvents than observed for different buffer addition rates,
where vesicle diameters were 97 nm for 600 μL min –1 and 212 nm for 60 μL min –1 ( Table S8 ). This correlates well with the smaller
differences in the quality of coated SiO 2 NPs comparing
IPA and ethanol, while a change in buffer addition rate from 60 μL
min –1 to 600 μL min –1 has
a much larger effect on the quality of the coating process. 

Effect of Solvent:Buffer Ratio on SiO 2 NP Lipid Coating Subsequently, the effect of solvent:buffer
ratio on the lipid-coating of SiO 2 NPs was studied. We
imaged SiO 2 NPs using cryo-TEM after halting the buffer
addition process at an ethanol:buffer ratio of 50:50 and 10:90 ( o,p). For 50:50,
14% of SiO 2 NPs presented a vesicular coating and 25% presented
a full lamellar (i.e., single-layer) coating ( q), whereas for 10:90 no vesicular coating
was observed and 38% of SiO 2 NPs presented a full lamellar
coating ( l).
These cryo-TEM results show that the vesicles in the bulk are attracted
to the surface of SiO 2 NPs to some extent, suggesting a
small contribution of vesicle rupture to the formation of lamellar
coatings ( a(ii)).
However, at a final solvent to buffer ratio of 10:90, no absorbed
vesicles could be observed, which implies that vesicles play a minor
role in the coating process. Furthermore, halting the buffer addition
at 50:50 will allow the system time to reach equilibrium, therefore
allowing a larger fraction of lipids to assemble into micelles and
eventually vesicles, as compared to the uninterrupted buffer addition
up to 10:90. Taking into account the earlier solvent-exchange studies
on planar surfaces, , reporting minor to no contribution
of vesicle rupture to the coating formation during buffer exchange,
vesicles present in the 50:50 case could have also adsorbed to the
uncoated parts of a particle surface without necessarily participating
in coating formation. These hypotheses, however, warrant a more in-depth
biophysical study to unveil the exact contribution of vesicles to
coating formation. Lastly, it is important to address the significant
fraction of uncoated (∼40%) and partially coated (∼20%)
SiO 2 NPs observed in cryo-TEM. This phenomenon represents
a common issue in the lipid-coating of nanoparticles, and has been
similarly reported elsewhere using vesicle fusion techniques. , It is our understanding that the natural process of lipid–surface
interactions occur at the micelle/vesicle state, and that the natural
curvature of nanoparticles reduces the extent of micelle/vesicle adsorption,
thus limiting the occurrence of vesicle fusion and lipid bilayer formation.
In contrast with other methods, however, the coating efficiency of
OPSALC has potential for significant improvement. For example, the
use of different lipids could potentially improve lipid–surface
attraction and may facilitate lipid packing within the layer. In addition,
the use of considerably higher lipid excess to particle surface area
should improve the fraction of fully coated particles, as we used
5 × 10 –17 mg mL –1 nm –2 while earlier studies on solvent assisted lipid coating
of planar surfaces have used 1 × 10 –15 mg mL –1 nm –2 . 

Applying OPSALC to Coat SiO 2 Microparticles Subsequently, we attempted to lipid-coat SiO 2 MPs using
the OPSALC procedure. Similar to coating the nanoparticles, we found
that the type of solvent, the rate of addition of buffer, solvent:buffer
ratio and lipid concentration were determining our success of achieving
a homogeneous lipid coverage on SiO 2 MPs ( ). Confocal microscopy analysis
revealed that the coating efficiency for SiO 2 MPs was 99.7%
( Figure S7 ). A higher lipid fluorescence
per particle was observed for IPA than for ethanol gradients ( a–d), indicating
coatings with a higher lipid content in the case of IPA. This higher
lipid fluorescence could indicate a denser or thicker coating, or
a better surface coverage, but could also be rationalized with more
lipids penetrating the pores of the particles. Homogeneous coatings
were found for all tested buffer addition rates using ethanol:buffer
gradients (>3 mL min –1 , 600 μL min –1 , 60 μL min –1 , e–g). The analysis of
fluorescence
intensity plot profiles of coated particles revealed different degrees
of lipid penetration into the pores of the particles depending on
the addition rate, with the lowest condition (60 μL min –1 ) yielding the highest degree of penetration ( Figure S8 ). Similarly, upon increasing lipid
concentration, an increase of lipid penetration into the particles
was observed ( Figure S11 ). Concentrations
of 0.25, 0.5, 1.0, and 2.0 mg mL –1 were used, which
corresponded to 0.1, 0.2, 0.4, and 0.8 mg per mg of SiO 2 MPs and n ≈ 44, 88, 176, and 353 lipid bilayers per particle,
respectively ( Method S2 ). Furthermore, the amount and
homogeneity of the coating were substantially different when halting
the buffer addition at solvent:buffer ratios of 100:0, 75:25, 50:50,
25:75 and 10:90 for both ethanol and IPA. In the case of ethanol,
high intensity (i.e., high lipid content) and homogeneous coatings
were observed at buffer contents of ≥75%, whereas at lower
buffer contents (<75%) coatings were less homogeneous, showing
few high intensity (i.e., high lipid content) features distributed
throughout a homogeneous low intensity (low lipid content) coating.
( i–m, Figures S9,10 ). It should be noted that lipid penetration increased,
not only with
a decrease in buffer addition rate and higher lipid concentration,
but also with the use of IPA instead of ethanol, an effect clearly
visible for 25:75 solvent:buffer ratios ( Figures S10,11 , h, red circles). Lower buffer addition rates are believed to provide
more time for lipid monomers and small lipid assemblies to diffuse
into the pores of the particles, whereas higher lipid concentrations
and the use of IPA supply higher amounts of lipid monomers and smaller
assemblies, which can penetrate the 8–10 nm sized pores. Therefore,
we tentatively assign the increased lipid penetration to (i) the availability
of higher fractions of lipid monomers and small lipid assemblies,
and (ii) the longer time allowed for lipids to diffuse into the pores
of the particles. Altogether, we concluded that the optimal
conditions for OPSALC
to yield a homogeneous coating on SiO 2 MPs (with limited
penetration) are a moderate excess of lipids (n ≈ 44 lipid
bilayers per particle), a minimum buffer addition rate of 600 μL
min –1 , and a final solvent:buffer ratio of 10:90
using either ethanol or IPA as the solvent. 

Erythrocyte Membrane-like Lipid Coatings on
SiO 2 MPs Next, we explored the use of OPSALC in
the development of erythrocyte membrane-like coatings on SiO 2 MPs. Lipidomics studies of the erythrocyte membrane revealed that
its outer leaflet, the one in direct contact with blood, has a main
composition of phosphatidylcholines (42.3 mol %), sphingomyelins (55.3
mol %), phosphatidylserine (1.5 mol %), and phosphatidylinositol (0.9
mol %), with a varying concentration
of cholesterol (30–40 mol %). Based
on these reports, we selected a phosphatidylcholine lipid (DOPC),
a sphingomyelin lipid (egg SM, mainly 16:0 fatty acid tails), a phosphatidylinositol
lipid (soy PI, mainly 18:1 fatty acid tails), and cholesterol (Chol)
as the components of the cell membrane-like formulations. Phosphatidylserine,
although present in the outer leaflet of the membrane, is a typical
biomarker of cell aging and certain pathologies , and was therefore excluded from the cell membrane design. The different
lipids were combined into formulations, according to the lipidomics-derived
molar ratios: DOPC only (D), DOPC and Chol (C), DOPC, Chol and egg
SM (S); and DOPC, Chol, egg SM and soy PI (I) ( a). For microscopy, fluorescent versions of these formulations
(DT, CT, ST, IT, respectively) were prepared by adding 0.1 mol % of
TR-DHPE. As expected, all fluorescent erythrocyte membrane-like
formulations
yielded homogeneous lipid coatings on SiO 2 MPs using the
OPSALC method ( b,c), retaining at least 70% of initial fluorescence intensity for
1 week ( f, Figure S12 ). Subsequently, measurements of fluorescence
recovery after photobleaching (FRAP) clearly distinguished lipid mobility
in the different coatings on SiO 2 MPs ( d,e). For DT and CT coatings,
nearly identical recovery kinetics were found, with higher lipid mobilities
than for ST and IT coatings, which showed near identical recovery
kinetics among themselves as well. Obviously, FRAP experiments on
microparticles, wherein the bleached area constitutes a significant
fraction of the total particle surface area, will show incomplete
recovery. The differences in mobility observed in FRAP could be explained
by differences in lipid phases formed by the different lipid formulations.
Miscibility diagrams of DOPC and cholesterol show that D and C coatings
are expected to be in a liquid-disordered phase at room temperature. In contrast, egg SM has a much higher chain
melting temperature (38 °C) than DOPC (−17 °C) and
causes mixtures of DOPC, Chol and egg SM, like formulation S and I,
to be in the two-liquid phase, containing liquid-disordered and liquid-ordered
domains at room temperature. It has been
shown that liquid-ordered phases have significantly lower lipid mobility
than liquid-disordered ones, corroborating our findings that D and
C coatings are more mobile than S and I coatings. Furthermore, it
stands to reason that lipid mixtures (i.e, formulations S and I),
which result in partially liquid-ordered layers due to their tighter
packing and stronger lipid–lipid attraction, are more prone
to form larger, more stable lipid assemblies during OPSALC. This would
also mean that lower amounts of lipid monomers and small lipid assemblies
are present, limiting the possibility of penetration of lipids into
the pores of the particles. This observation, in turn, is in agreement
with the smaller penetration depth for S and I, as effectively observed
in confocal microscopy ( b,c). The observed differences show the potential of using
OPSALC to tailor the penetration depth by varying not only buffer
addition rate, lipid concentration and choice of solvent, but also
by varying the phase of the lipid layers. Overall, the OPSALC
method was used to successfully coat SiO 2 MPs with various
erythrocyte membrane-mimicking lipid coatings,
with distinct characteristics depending on the exact formulation applied. 

Erythrocyte Membrane-like Coatings on SiO 2 NPs To confirm that erythrocyte membrane-like formulations
could also be applied to nanomaterials, we studied the efficiency
of OPSALC on SiO 2 NPs. Particles were coated through IPA:buffer
gradients using formulations D, C, S, and I, and subsequently analyzed
using DLS and zeta potential (ZP) measurements. Formulations D and
C resulted in a larger particle size distribution than S and I ( g) and the polydispersity
decreased linearly going from D, C, S to I ( h). All coatings produced a reduction in
the ZP, from −48 mV in uncoated particles to approximately
−30 mV in coated ones ( i). Among the tested conditions, particles coated with
formulation I produced the lowest ZP value (−34.4 mV) and the
lowest polydispersity index (0.26), thus displaying the highest colloidal
stability for lipid-coated SiO 2 NPs. To further characterize
the coatings, SiO 2 NPs were analyzed with cryo-TEM. All
formulations successfully produced single-layer coatings on the particles
( j–n),
corroborating our previous cryo-TEM results. Coating efficiency was
further quantified as percentage of uncoated, partially coated, and
coated particles, revealing that formulations S and I generated a
considerably higher percentage of fully coated particles than formulations
D and C (D: 18%, C: 18%, S: 50%, I: 41% o). Previously, we had proposed that lipid–surface
and lipid–medium interactions were the main factor determining
the efficiency of OPSALC. Interestingly, we have now observed that
lipid–lipid interactions also play a major role in coating
efficiency, as formulations with tighter lipid packing (S, I) produce
a considerably higher percentage of coated particles. Presumably,
the differences in packing and hydration of the partially liquid-ordered
layers improved lipid–surface interactions, compensating for
the expected occurrence of lower amounts of monomers and small lipid
assemblies. Overall, we confirmed that SiO 2 NPs can
be coated with erythrocyte membrane-like formulations using the OPSALC
method, and observed that coating SiO 2 NPs with formulations
producing liquid-ordered and -disordered phases (S, I) was more effective
than coating them with formulations producing a liquid-disordered
phase only (D, C). 

Erythrocyte Membrane-like Coatings on PCL
MPs Next, we attempted to coat hydrophobic PCL MPs using
the OPSALC method. PCL was chosen as a hydrophobic particle substrate
due to its bioresorbable composition and its common use in drug delivery
and tissue engineering. It is often preferred over other hydrophobic
polymers (e.g., PLA or PLGA) due to its low cost, high availability,
and lower rate of biodegradation. , Here, the
fluorescent erythrocyte membrane-like formulations used for SiO 2 MPs were also employed for coating visualization studies. During the coating process, we observed that coated microparticles
were easier to redisperse in water than uncoated microparticles, and
that S- and I-coated PCL MPs were significantly easier to redisperse
than D- and C-coated PCL MPs ( p). Fluorescence imaging revealed the highest fluorescence
intensity for I-coated PCL MPs, with C- and S-coated particles of
only slightly lower intensity, and a significantly lower intensity
for D-coated PCL MPs ( q). FRAP measurements revealed slow recovery rates, comparable
for all types of coating ( r,s), indicating that all lipid layers had limited
mobility. We propose that this reduced mobility is due to hydrophobic
interactions between the fatty acid tails of the lipids and the surface
of PCL particles, which may have resulted in a single immobile monolayer,
as suggested elsewhere for another planar hydrophobic substrate. Altogether, these results indicate that OPSALC
can be employed to successfully coat hydrophobic PCL MPs with different
mixtures of lipids, resulting in layers with low mobility irrespective
of lipid composition. 

Hemocompatibility Testing of Lipid-Coated
Particles To demonstrate the functionality of distinct lipid
coatings, the effect of lipid formulation on particle hemocompatibility
was investigated. We first conducted hemolysis and plasma coagulation
assays on SiO 2 NPs, as they are known for their highly
hemolytic and procoagulant nature. , Hemolysis
assays showed that uncoated SiO 2 NPs are extremely hemolytic
at particle concentrations as low as 0.2 mg mL –1 , and specifically more hemolytic than the positive control (1% v/v
Triton-X final concentration), which led to its use as the reference
for hemolysis rate calculation. In contrast, SiO 2 NPs coated
with any of the lipid formulations yielded a drastic reduction of
erythrocyte damage, coupled to a reduction in the amount of hemoglobin
release and the resulting hemolysis rates (from 100% to below 5%, a,b). Moreover, no
significant difference was observed between lipid formulations and
the negative control, confirming its inert behavior. It must be noted,
however, that the particle aggregation observed for D- and C-coated
SiO 2 NPs ( g) could be partially responsible for the reduction in hemolysis
rate. As a result, the low hemolysis rates of coatings S and I are
more reliable than the hemolysis rates of coatings D and C, as they
more accurately indicate the hemoprotective effect of the lipid layer
on single particles. To determine whether the lipid composition of the
particle coatings
have an effect on plasma coagulation time, coated and uncoated SiO 2 NPs were incubated with platelet-poor plasma, recalcified,
and analyzed via the increase in scattering during fibrin clot formation.
SiO 2 surfaces are known for their procoagulant properties
through the surface-induced activation of factor XII, or Hageman factor,
an important mediator of the intrinsic coagulation pathway. , Plasma coagulation curves demonstrated that S and I coatings could
induce a more pronounced delay in coagulation onset than D and C coatings
( c), demonstrating
a more significant attenuation of the pro-coagulating properties of
SiO 2 NPs, although this advantage was not consistently
observed. The coagulation curves for two other tested batches can
be found in Figure S13 . Plasma coagulation
time was then calculated as average of all batches tested. Overall,
the average coagulation time induced by different coatings was not
significantly different; however, a significant difference was observed
between the uncoated and all coated conditions ( d). Collectively, these results indicate
that all tested coatings were effective in attenuating the coagulating
effect of SiO 2 NPs. To further study the hemocompatibility
effects of the lipid coatings,
we explored the complement activation, platelet activation, leukocyte
recruitment, and inflammation response induced by uncoated vs lipid-coated
PCL MPs. Despite the low toxicity of this polymer, hydrophobic surfaces
are known to adsorb fibrinogen, IgG, and C3, proteins involved in
platelet/leukocyte adhesion and complement activation. , Since hydrophobic micromaterials have been researched as drug carriers, , porous functional scaffolds, and, more
recently, oxygen-generating components of tissue engineered implants, , hemocompatibility has become a key aspect of their biocompatibility. Indeed, PCL MPs stimulated the production of all complement activation,
platelet activation, and leukocyte recruitment markers when compared
to nonstimulated blood ( e). The relatively high levels of C5a, a common biomarker
for all complement activation pathways, indicated that PCL MPs tend to induce complement activation to some
extent. Lipid coatings, however, attenuated this complement-activating
effect with an interesting trend: increasingly complex lipid formulations
produced a gradually safer response. Neutrophil chemotactic marker
IL-8 followed a similar trend, with more
complex lipid coatings resulting in lower IL-8 production. Platelet
activation markers (β-TG and PDGF-BB), , platelet-secreted monocyte and lymphocyte attractant RANTES, and monocyte attractant MCP-1 were found at generally low levels for all coating formulations.
Finally, activated T-cell cytokine MIF was found at similarly high levels in uncoated, D-coated and C-coated
PCL MPs, but found at lower levels in S-coated and I-coated PCL MPs.
These results suggested that PCL MPs were not entirely bioinert in
blood, and that lipid coatings with more complex erythrocyte membrane
mimicry were able to notably improve PCL MP hemocompatibility. To
expand on these results, the levels of inflammatory biomolecules (IL-1RA,
IL-1β, IL-6, and TNF-α) were also analyzed. While there
was no statistically significant difference between the negative control
and any PCL MPs for any of these biomarkers, there was a consistent
tendency for D-coated PCL MPs to have an almost statistical difference
from the negative controls ( f–i), indicating a slightly more pro-inflammatory
profile than uncoated and other coated particles. Ultimately,
all results corroborated that S and I lipid formulations
generated a better improvement on blood-contacting particle hemocompatibility
as compared to D and C lipid formulations, indicating a potential
benefit in using biomimetically complex lipid coatings in biomedical
applications. 

Conclusions We demonstrated that on-particle
solvent-assisted lipid coating
(OPSALC) can be successfully used to lipid-coat particles of different
sizes and materials. The efficiency of the OPSALC method depends,
among others, on the extent of the solvent:buffer gradient, on the
fractions and types of lipid species formed throughout the process,
and on the balance of lipid–particle and lipid–medium
interactions. Moreover, we demonstrate that OPSALC enables a facile
and scalable lipid-coating of particles using biologically relevant
lipid formulations in a safer, more sustainable, and ethically more
acceptable manner than natural cell membrane coatings. Different erythrocyte
membrane-like lipid formulations have been tested, offering new entries
into cell membrane engineering when designing blood-contacting lipid
coatings. Moreover, we demonstrated that erythrocyte membrane-like
lipid formulations can significantly improve particle hemocompatibility,
as well as enhance the colloidal stability of hydrophobic microparticles
in water. Particles coated with synthetic cell membranes can be incorporated
in the design of novel drug delivery systems, engineering cell mimics,
building blocks for tissue engineering, and medical devices, providing
a hemocompatibility enhancement toolbox for future biomedical and
pharmaceutical innovation. 

