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Mikrochim Acta
Mikrochim Acta
Mikrochimica Acta
0026-3672
1436-5073
Springer Vienna Vienna

39168870
6631
10.1007/s00604-024-06631-7
Original Paper
Exploring the influence of silicon oxide microchips shape on cellular uptake using imaging flow cytometry
Bruce Gordon 1
Bagherpour Saman 23
Duch Marta 4
Plaza José Antonio 4
Stolnik Snow 5
http://orcid.org/0000-0003-2031-4405
Pérez-García Lluïsa mlperez@ub.edu

123
1 https://ror.org/01ee9ar58 grid.4563.4 0000 0004 1936 8868 Division of Advanced Materials and Healthcare Technologies, School of Pharmacy, University of Nottingham, Nottingham, NG7 2 UK
2 https://ror.org/021018s57 grid.5841.8 0000 0004 1937 0247 Departament de Farmacologia, Facultat de Farmàcia I Ciències de L’Alimentació, Toxicologia I Química Terapèutica, Universitat de Barcelona, Av. Joan XXIII 27-31, 08028 Barcelona, Spain
3 grid.5841.8 0000 0004 1937 0247 Institut de Nanociència i Nanotecnologia (IN2UB), Universitat de Barcelona, 08028 Barcelona, Spain
4 grid.424142.5 0000 0004 1803 4225 Instituto de Microelectrónica de Barcelona IMB-CNM (CSIC), Campus UAB, Cerdanyola del Vallès, 08193 Barcelona, Spain
5 https://ror.org/01ee9ar58 grid.4563.4 0000 0004 1936 8868 Division of Regenerative Medicine and Cellular Therapies, School of Pharmacy, University of Nottingham, Nottingham, NG7 2 UK
21 8 2024
21 8 2024
2024
191 9 5542 5 2024
8 8 2024
© The Author(s) 2024
2024
https://creativecommons.org/licenses/by/4.0/ Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.
Nano- and micro-carriers of therapeutic molecules offer numerous advantages for drug delivery, and the shape of these particles plays a vital role in their biodistribution and their interaction with cells. However, analysing how microparticles are taken up by cells presents methodological challenges. Qualitative methods like microscopy provide detailed imaging but are time-consuming, whereas quantitative methods such as flow cytometry enable high-throughput analysis but struggle to differentiate between internalised and surface-bound particles. Instead, imaging flow cytometry combines the best of both worlds, offering high-resolution imaging with the efficiency of flow cytometry, allowing for quantitative analysis at the single-cell level. This study focuses on fluorescently labelled silicon oxide microchips of various morphologies but related surface areas and volumes: rectangular cuboids and apex-truncated square pyramid microchips fabricated using photolithography techniques, offering a reliable basis for comparison with the more commonly studied spherical particles. Imaging flow cytometry was utilised to evaluate the effect of particle shape on cellular uptake using RAW 264.7 cells and revealed phagocytosis of particles with all shapes. Increasing the particle dose enhanced the uptake, while macrophage stimulation had minimal effect. Using a ratio particle:cell of 10:1 cuboids and spheres showed an uptake rate of approximately 50%, in terms of the percentage of cells with internalised particles, and the average number of particles taken up per cell ranging from about 1–1.5 particle/cell for all the different shapes. This study indicates how differently shaped micro-carriers offer insights into particle uptake variations, demonstrating the potential of non-spherical micro-carriers for precise drug delivery applications.

Graphical Abstract

Supplementary Information

The online version contains supplementary material available at 10.1007/s00604-024-06631-7.

Keywords

Cellular uptake
Functionalization
Imaging flow cytometry
Fluorescence label
Microfabrication
Silicon oxide microchips
http://dx.doi.org/10.13039/501100004837 Ministerio de Ciencia e Innovación PID2020-115663GB-C3 1 PID2020-115663GB-C3-2 Plaza José Antonio Pérez-García Lluïsa Agaur2021 SGR 01085 FISDUR Bagherpour Saman Pérez-García Lluïsa http://dx.doi.org/10.13039/501100000266 Engineering and Physical Sciences Research Council EP/L01646X Stolnik Snow http://dx.doi.org/10.13039/100004330 GlaxoSmithKline Universitat de BarcelonaOpen Access funding provided thanks to the CRUE-CSIC agreement with Springer Nature.

issue-copyright-statement© Springer-Verlag GmbH Austria, part of Springer Nature 2024
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pmcIntroduction

Nano- and microscale particle carriers, designed to transport therapeutic molecules, provide numerous benefits for biomedical applications, including drug delivery, biosensing, targeted cancer therapies, gene editing, and regenerative medicine [1]. Recently, the amount of research in this area has risen drastically; although there still remain many hurdles to increasing the number of approved pharmaceutical products [2], some associated with their cellular uptake, influenced by some features of the particles such as particle size, surface chemistry, charge, and mechanical properties. Development of the research in this area provides opportunities to rationally design drug delivery systems based on the desired therapeutic outcome.

Particle shape is also an important determinant in the biodistribution of particles after administration to the body, and particular attention has been paid to the movement of micron-sized particles of different shapes in flow [3]. The hydrodynamics of non-spherical microparticles in blood often result in their margination towards the blood vessel walls, leading researchers to explore their use in vascular targeting [4]. Decuzzi et al. [5] have demonstrated differences in biodistribution between spheres, disks, cylinders, and hemispherical particles, opening another avenue for consideration in the targeting of different areas of the body using different particle shapes.

Studies of the uptake of silicon oxide particles of different shapes have primarily been focussed on respirable silica dusts in order to understand how the particle morphology affects toxicity in macrophages [6–8]. Recently, an increased number of studies have begun to examine silicon oxide particle morphology as a tool for enhancing control over drug delivery, for example by modulating uptake by cells [9–12].

It is difficult to make clear comparisons between these studies because the effects of particle shape are difficult to study in isolation and because experimental conditions tend to not coincide. Differences in particle size, surface chemistry, and cell type are ever present and have been shown to play important roles in particle internalisation [13–15]. Additionally, cellular association is often used as a proxy for internalisation. Moreover, the majority of studies examining the effect of silicon oxide particle shape utilise particles that are in the nano size range [16–20], whereas there are fewer studies that look at silicon oxide particles in the micron size range (> 1 μm).

The methodology of analysing particle internalisation is a key aspect to be considered. There are numerous methods available to assess the internalisation of particles into cells in vitro, which can be divided into qualitative or semiquantitative methods such as confocal/fluorescence/light microscopy and scanning electron microscopy and quantitative (flow cytometry, plate reader methods, inductively coupled mass spectrometry (ICP-MS)) [21]. Qualitative methods rely on high-resolution imaging to allow visualisation of particle internalisation events and are useful in providing descriptive information about the sample. However, often these methods are time-consuming and expensive which therefore leads to a reduction in the number of samples that can be assessed, thus decreasing the reliability of the quantitative information that can be obtained. Furthermore, due to the selection of which cells are imaged by the researcher, these methods can be subject to unconscious sampling bias that can reduce the validity of the obtained results [22].

On the other hand, quantitative methods are able to assess high numbers of cells in order to give highly reliable information about the sample but lack the sensitivity in discriminating between internalised and surface-bound particles [23]. For example, assessment of cells by flow cytometry or plate reader methods will give information about the proportion of cells that are associated with fluorescent particles, but not the location of particles with respect to the cells. Further methodological steps can be used, for example by quenching extracellular fluorescence or by staining surface-bound particles with a secondary fluorophore to discriminate between cells with internal and cells with surface-bound particles. However, these methods are very system specific, requiring efficient quenching/labelling, and therefore are not suitable as a general method.

Imaging flow cytometry (IFC) is a recently developed method that aims to overcome some of these limitations by combining the spatial resolution of microscopy with the high-throughput nature of flow cytometry. IFC generates quantitative image data for every event present, such as cells, cell clusters, and debris, which enables the detailed analysis of individual cells within diverse populations, contributing to our knowledge of cellular heterogeneity [24]. IFC has been used by a number of groups to assess the internalisation of nanoparticles, and it requires particle fluorescent labelling [25, 26].

Our group has expertise in the functionalization of hexahedral-shaped silicon-based microchips, fabricated by silicon-based technologies, for intracellular pH sensing [27] which showed that the chemical integrity and functionality of immobilised compounds (proteins, antibodies, or fluorophores) remain after conjugation with these types of microchips. Moreover, we have demonstrated the versatility of particle size and shapes that can be fabricated using photolithography [28, 29] followed by exhibiting the low polydispersity micro-fabricated particles in the aqueous media [30, 31]. However, the effect of silicon oxide microparticle shape is yet to be investigated.

In this work, silicon oxide microchips (or microparticles) of rectangular cuboid and apex-truncated square pyramid morphologies, which are called cuboids and pyramids, fabricated using photolithography, are compared with silicon oxide microspheres acquired from a commercial supplier to assess their interactions with RAW 264.7 macrophages. All microchips offer different morphologies but similar surface areas and volumes initially. Two siloxane linkers of different length were used for the functionalization of the microchips with fluorescent labels, and the interaction with RAW 264.7 macrophages of the functionalised particles was assessed using IFC.

Experimental

The Materials and General Methods sections are included in the supporting information.

Silicon oxide microchips fabricated by silicon-based technologies for the production of chips

For silicon oxide microchips preparation, the wet oxidation method was applied to grow the silicon oxide layer with a thickness of 1 μm on the surface of silicon and the plasma-enhanced chemical vapour deposition (PECVD) to deposited a silicon oxide layer with a thickness of 3 µm. The main process parameters of the 1-µm thermal oxide layer are as follows: 1100 °C and growing time 2 h 25 min. Meanwhile, the parameters for the 3-µm PECVD silicon oxide layer are as follows: 400 °C, plasma power 400 W, pressure 9 torr, gases (TEOS 500 SCCM and O2 SCCM), and deposition time 6 min 15 s. Then, on the silicon oxide layer, spinning a positive photoresist with a thickness of 1.2 μm was performed. Afterward, UV light through the photomask was applied for defining the shape of particles, as well as the lateral nominal dimensions (3 × 3 µm2). After that, the C2F6 and CHF3 mixture were used for dry etching the silicon oxide layer, followed by the photoresist stripping. Then, anchors were defined underneath particles by employing quasi-isotropic reactive ion etching (RIE). Finally, silicon oxidation (1100 °C and 5 h 15 min) was applied to convert the silicon pillars in silicon oxide pillars [27], and this annealing also makes more comparable the PECVD silicon oxide to the thermal grown one.

Covalent labelling and removal of silicon oxide microchips anchored on silicon wafers

Silicon wafers possessing silicon oxide microchips attached to the surface by supporting anchors were cut using a diamond scribe into ~ 5 × 5 mm pieces. Pieces of wafer were transferred to a 10-mL glass vial (four wafer pieces per vial) with the silicon oxide particle pattern facing up. Acidic piranha solution (1 mL) was prepared by adding H2O2 (35%, 300 μL) dropwise to H2SO4 (98%, 700 μL), and this solution transferred into the vial containing the wafer pieces. This solution was freshly prepared each time to avoid loss of activity. After 1 h, the acidic piranha solution was removed, and the surfaces rinsed with Milli-Q water (3 × 1 mL), dried under a nitrogen stream, and added to a fresh 10-mL glass vial.

Basic piranha solution was prepared by adding NH4OH (20%, 150 μL) to Milli-Q water (700 μL), and to this mixture, H2O2 (35%, 150 μL) was added dropwise. Basic piranha solution was transferred to the vial containing the wafer pieces for 30 min. This solution was freshly prepared each time to avoid loss of activity. The basic piranha solution was removed, and the wafer pieces washed with Milli-Q water (3 × 1 mL) and dried under a stream of nitrogen before being transferred to a fresh 10-mL glass vial.

Freshly activated wafer pieces were placed in a 10-mL glass vial with the silicon oxide particle facing up and submerged in siloxane solution: either 2% v/v APTMS solution in acetone (1 mL) for 30 min or 0.1% v/v AUTES solution in EtOH (1 mL) for 2 h. Siloxane solutions were removed, and the wafer pieces washed with 3 × 1 mL of the corresponding solvent (acetone for APTMS-treated wafer pieces, EtOH for AUTES treated wafer pieces) before being dried with a nitrogen air stream and placed in a fresh 10-mL glass vial. RBITC solution in EtOH (1 mL) was then added to the wafer pieces which were protected from light and left overnight. For experiments comparing the covalent labelling methods, a 200-μM solution of RBITC in EtOH (1 mL) was used, and for all subsequent experiments, 30-μM RBITC solution in EtOH (1 mL) was used. Alternatively, BDP 630 NHS ester (30 μM) in Na2HPO4/NaH2PO4 buffer pH 8 (700 μL) was added to the wafer pieces, protected from light and left overnight. Wafer pieces were washed with 3 × 1 mL EtOH (RBITC-labelled wafer pieces) or Milli-Q water (BDP 630-labelled wafer pieces) and dried with a stream of nitrogen before removal of particles by peeling.

For the removal of silicon oxide particles from surface, a drop of Fluoromount™ aqueous mounting medium was placed onto each of the wafer pieces and spread using a micropipette tip to ensure complete coverage. This was then left to air-dry (~ 20 min). After drying, the mounting medium was peeled away from the wafer piece surface using tweezers and placed in a 1-mL microcentrifuge tube [27]. Milli-Q water (1 mL) was added to the microcentrifuge tube and sonicated to aid dissolution of the mounting medium. After complete dissolution had occurred, the microcentrifuge tube was centrifuged for 10 min at 5700 RPM (6175 G), resulting in a pellet forming. The supernatant was removed, and the cycle of centrifugation and washing with Milli-Q water (1 mL) was repeated two times. Particles were stored in EtOH (700 μL) at 4 °C.

Covalent labelling of spherical silicon oxide microchips in suspension

A total of 1 mg (20 μL) silicon oxide spheres as received (stock concentration 5% w/v) were transferred to a 1.5-mL microcentrifuge tube and centrifuged at 5700 RPM (6175 G) for 10 min and the supernatant removed. Particles were suspended in acidic piranha solution (700 μL) for 1 h with shaking to prevent particles from falling out of suspension. Particles were centrifuged at 5700 rpm (6175 G) for 10 min and the acidic piranha solution removed. Particles were washed by resuspension in 700-μL Milli-Q water and then centrifuged 5700 rpm (6175 G) for 10 min. This washing process was repeated three times and then the supernatant removed. Particles were suspended in basic piranha solution (700 μL) and left shaking for 30 min. Particles were centrifuged at 5700 rpm (6175 G) for 10 min and the basic piranha solution removed. Particles were washed using the same resuspension in 700-μL Milli-Q water and centrifugation steps used to wash the particles after removal of the acidic piranha solution.

Freshly activated silicon oxide particles were suspended in 0.1% v/v AUTES in EtOH (700 μL) and left to shake for 2 h. Particles were centrifuged at 5700 RPM (6175 G) for 10 min and the silane solution removed. Particles were washed by resuspension in 700-μL EtOH and centrifugation at 5700 RPM (6175 G) for 10 min before removal of the supernatant. This washing process was repeated three times and then the supernatant removed. Particles were suspended in RBITC (30 μM) in EtOH (700 μL) or BDP 630 NHS ester (30 μM) in Na2HPO4/NaH2PO4 buffer pH 8 (700 μL) and left on a shaker overnight while protected from light. Particles were washed with EtOH 3 × 700 μL (RBITC-labelled particles) or Milli-Q water 3 × 700 μL (BDP 630-labelled particles) by repeated suspension and centrifugation as above. Particles were stored in 700-μL EtOH at 4 °C.

Determination of trypan blue quenching efficiency

Measurement of median fluorescence intensity (MFI) of particles was measured by imaging flow cytometry using a 50-μL suspension containing RBITC-labelled particles in 1% HEPES in HBSS. MFI was measured with and without the addition of trypan blue solution (0.4%, 10 μL) immediately before sample analysis. Flow cytometer settings were kept the same for each sample, and details can be found in the flow cytometry section.

Qualitative particle uptake of silicon oxide microchips by RAW 264.7 macrophages assessed by confocal microscopy

BDP 630-labelled particles were centrifuged at 5700 RPM (6175 G) for 10 min. From this point onwards, all work took place inside a sterile cell culture hood to maintain the sterility of the particles. Stock particle suspensions were prepared by removal of the EtOH and the addition of 1-mL buffer comprising 1% HEPES in HBSS. A volume of each particle stock solution containing 1,500,000 particles was transferred into fresh microcentrifuge tubes and the volume made up to 1 mL with fresh 1% HEPES in HBSS buffer.

RAW 264.7 cells grown to 60–80% confluency were harvested by scraping and counted by the protocol described in the supporting information. A total of 300,000 cells per well in 1-mL culture media were seeded onto collagen-coated glass coverslips inside clear polystyrene 6-well plates and incubated at 37 °C, 5% CO2, and 95% humidity overnight to allow cell attachment. Culture media was then removed, and the cells washed with 1-mL pre-warmed (37 °C) PBS. The PBS was removed by aspiration, and 1 mL per well pre-warmed particle suspensions each containing 1,500,000 particles added to each well. Cells were kept in an incubator for 4 h after which time the particle suspensions were removed by aspiration and the cells washed with 3 × 1 mL pre-warmed PBS. Cells were fixed by the addition of 1 mL per well of 4% formaldehyde in PBS for 15 min and then washed with 3 × 1 mL PBS. Permeabilization of cells was achieved by adding 1 mL per well 0.1% Triton X-100 in PBS for 3 min. Cells were washed with PBS (3 × 1 mL) and the PBS removed. The immunofluorescence staining and preparation of confocal slides are explained in the supporting information.

Quantitative uptake of silicon oxide microchips by RAW 264.7 cells assessed by flow cytometry

RBITC-modified particles were prepared according to the same procedure explained in the “Preparation of fixed cells for flow cytometry: imaging flow cytometry (IFC) and trypan blue quenching (TBQ) method” section. RAW 264.7 cells were also harvested once they reach 60–80% confluency by scraping according to the described protocol in the supporting information. A total of 100,000 cells in culture media were seeded per well into a 12-well plate and incubated overnight at 37 °C, 5% CO2, and 95% humidity. In experiments where RAW 264.7 cells were stimulated with LPS, culture media was used without 1% penicillin/streptomycin and was supplemented with 100-ng per mL LPS. Culture media was removed by aspiration and the cells washed with 1-mL pre-warmed (37 °C) PBS. PBS was removed and 1 mL pre-warmed particle suspensions applied at ratios of 5 or 10 particles per cell. Cells were incubated for 4 h after which time the particle suspensions were removed and the cells washed with warm HBSS (3 × 1 mL). Cells were prepared for flow cytometry as either fixed or live cells depending on the analysis method.

Preparation of fixed cells for flow cytometry: imaging flow cytometry (IFC) and trypan blue quenching (TBQ) method

For both IFC method and TBQ, HBSS was removed from each well by aspiration and 250-μL Accutase added to each well, and the cells were placed in an incubator for 5 min to detach cells. A total of 250-μL HBSS was then added to each well and the total volume of each well transferred to fresh microcentrifuge tubes. Samples were centrifuged at 250 G for 5 min and the supernatants removed.

In the IFC experiment, cells were suspended in 250-μL 4% formaldehyde in PBS for 20 min before being centrifuged for 5 min (250 G) and the supernatant removed. Fixed cells were suspended in 50-μL HBSS and stored at 4 °C until analysis by flow cytometry. Samples were always analysed within 1 week of sample preparation. Regarding the TBQ method, cells were suspended in 50-μL HBSS and kept on ice until analysis by flow cytometry. All samples were analysed within 2 h of sample preparation. Immediately before analysis of each sample 10 μL 0.4%, trypan blue solution was added to quench surface-bound particle fluorescence.

Results and discussion

Characterisation of fabricated microchips

Silicon oxide microchips anchored on to silicon wafers were microfabricated using photolithography (Scheme 1). The method involves growing a thermal layer of silicon oxide on top of a silicon wafer (nominal 1-µm thick for the cuboids) and a PECVD deposition (nominal 3-µm thick for the pyramids), and a photomask is then used to selectively etch the particles that have a morphology which is defined by the photomask and the thickness of the original silicon oxide layer. After fabrication, particles remain anchored to the silicon wafer by a thin silicon support. This silicon support is thermally oxidised to allow single material microchips. This oxidation also anneals the PECVD oxide (3-µm pyramids) to make it more similar to the grown silicon oxide (1-µm cuboids). The anchors are posteriorly broken to release the particles into suspension.Scheme 1 Fabrication of A cuboids and B pyramids silicon oxide microchips. Ai, Bi Silicon substrate, Aii, Bii growing or depositing silicon oxide layer on the surface of silicon, Aiii, Biii spinning a positive photoresist on the silicon oxide layer, Aiv, Biv defining the dimensions of microchips by the utilisation of UV light through the photomask and a dry etching (RIE) of the silicon oxide layer, Av, Bv defining anchors by RIE process of the silicon substrate, Avi, Bvi photoresist removal, and Avii, Bvii silicon oxidation to convert the silicon pillars in silicon oxide pillars

SEM images of microfabricated silicon oxide microchips show highly uniform particle dimensions for both cuboids and pyramids (Fig. 1A and C). To ensure that all microchips are of the same size and feature precision, several key factors were controlled rigorously during the photolithography process, including alignment precision, uniform exposure of UV lights, and equipment calibration. In addition, environmental factors like temperature, humidity, and vibration were tightly controlled in the fabrication facility (cleanroom) to prevent any variations in the process. Figure 1B and D shows cuboid and pyramid microchips, respectively, removed from their silicon support, and a ca. 200-nm piece of the support pillar remains attached to the particle. Uniformity of particle size and shape is essential to ensure that any effects of particle shape on cellular uptake in later experiments are not caused by polydispersity. Particle dimensions were measured using ImageJ®, and the results are displayed in Table S1 in the supporting information.Fig. 1 SEM images of fabricated silicon oxide microchips. A Cuboids, B zoom detail of cuboid microchips, C pyramids, and D zoom detail of pyramid microchips

Both cuboids and pyramids have a wider base than at the top of the particles, and some rounding can be observed on the particle edges due to the limit of our photolithographic and etching processes, which is particularly evident in the pyramids. Surface area and volume of the microchips are related, although lateral dimensions are dissimilar, so are their morphology. For the calculation of particle volume and surface area, this curvature was not taken into account, and flat edges assumed. Silicon oxide spheres, acquired from a commercial supplier, had a diameter of 3 μm according to the manufacturer’s analysis.

Labelled silicon oxide microchips anchored on silicon wafers characterisation

Covalent labelling methods have been used by a vast number of groups in order to label silica nanoparticles with fluorescent molecules or other molecules of interest [32], and from the wide variety of silanes available, 3-aminopropyltrimethoxysilane (APTMS) and 11-aminoundecyltriethoxysilane (AUTES) were chosen (Fig. S1), in order to explore the influence of the spacer link in the functionalization process. APTMS and AUTES are both amine-terminated silanes, which offer stable terminal groups that can be coupled with amine reactive fluorophores such as rhodamine B isothiocyanate (RBITC) (Fig. 2 (1)).Fig. 2 1 Functionalization sequence for labelling of activated silicon oxide microchip surfaces with silane linkers APTMS/AUTES and RBITC. 2 Fluorescence of covalently labelled particles. A APTMS and RBITC and B AUTES and RBITC. i Brightfield, ii fluorescence, iii fluorescence surface plots, and iv fluorescence intensity profiles. Yellow lines on Aii/Bii represent the location from which fluorescence line plots Aiv/Biv were taken

Successful activation of the surface, using acidic and basic piranha treatment, was confirmed by measuring water contact angle; activated surfaces had a water contact angle < 10° indicating hydrophilic nature due to hydroxyl groups present on the surface. After activation, siloxanes were deposited on particles attached to wafers and their water contact angles measured. The results are shown in Table S2. The increase in contact angle from hydrophilic (< 10°) to hydrophobic (> 40°) is consistent with the presence of hydrocarbon chains of the silanes and thus indicates successful deposition of each of the silanes. APTMS and AUTES displayed comparable water contact angles to those reported previously in the literature [33]. APTMS exhibits lower contact angles (ca. 38°) than AUTES (ca. 61°) due to the shorter length of its hydrocarbon chains [34].

RBITC was added to APTMS- and AUTES-modified surfaces as shown in Fig. 2 (1). Fluorescence microscopy images of functionalized chips on surfaces showed that each of the silanes had been successfully immobilised onto the particle surface, which had subsequently been labelled (Fig. 2 (2)). In each case, plots of the normalised fluorescence intensity of the surfaces indicate the uniformity of the labelling on the surface of the particles, which was very consistent and exhibits surface homogeneity with the average normalised fluorescence intensity of 3.98 ± 0.07 (a.u.) measured from analysing 500 microchips. This is important for cell studies as an uneven distribution of fluorophore on the particle surface will cause a difference in interaction between the particle and cell and thus interfere with the results. AUTES was chosen as the linker for subsequent experiments because it has been demonstrated that linkers with longer carbon chains form stable monolayers that are more resistant to hydrolysis than shorter chains [33], which can be relevant to insure the stability of the labelling.

Prior to the uptake experiments, it was necessary to determine the stability of the fluorophores on the labelled particles, to ensure no fluorophore would release in buffer conditions at 37 °C. Wafer pieces possessing particles labelled covalently using AUTES-RBITC were incubated in PBS at 37 °C for 2 h. The fluorescence intensity of the particles was assessed before and after incubation using fluorescence microscopy (Fig. S2 in the supporting information). Median fluorescence intensity values were calculated by imaging 5 areas of each wafer piece and measuring the fluorescence intensity of 500 particles per area. The results show that no significant decrease in fluorescence intensity was observed with covalently labelled AUTES-RBITC particles.

Characterisation of fluorescently labelled microchips in suspension

Measurement of median fluorescence intensity (MFI) and surface coverage of fluorophore

Silicon oxide spheres, cuboids, and pyramids were fluorescently labelled using AUTES and RBITC. For the spheres, this process was performed in suspension, whereas cuboids and pyramids were fluorescently labelled while still attached to wafers by a supporting anchor followed by peeling away from the wafer by using a mounting medium as shown in Scheme S1 in the supporting information. All resulting labelled particles in suspension were imaged using fluorescence microscopy (Fig. S3 in the supporting information) revealing their successful labelling. Distinct particle shapes can be identified in the fluorescence images which correspond to the brightfield images indicating that RBITC is evenly distributed across the surface of the particles, and that the labelling was not affected by the release of the particles from the wafer. The normalised fluorescence intensity measured from fluorescent microscopy images for spheres, released cuboids, and released pyramids in the suspension was 3.67 ± 0.06 (a.u.), 3.57 ± 0.07 (a.u.), and 3.84 ± 0.08 (a.u.) for ~ 200 particles for each type, indicating the homogenous labelling according to standard deviations related to each type of particles.

In order to compare the degree of fluorescent labelling of each particle type, imaging flow cytometry was also used to measure the median fluorescence intensity (MFI) (n > 1000 particles of each shape). The obtained values and fluorescence intensity histograms are included in Fig. S4 (supporting information). The values show that each particle type has similar MFI values indicating a similar degree of labelling. To confirm this, the percentage of RBITC-labelled surface was estimated. This was done by comparing the MFI values of each particle type with a calibration curve of particles (3-μm spheres) that had known percentages of labelled RBITC (Fig. S5 in supporting information), and the results show that all particles had lower than 2% of surface labelled with RBITC.

It is important to have a similar and reduced amount of RBITC on the surface of the particles because rhodamine B is a relatively hydrophobic molecule (logP 1.95) and could promote uptake into cells [35]. Juliano et al. found that the uptake of dendrimers into HeLa cells was enhanced by their conjugation with Oregon Green, although the exact mechanism was not deduced [36]. Additionally, it has been shown that rhodamine B interacts with type BI scavenger receptors and so could enhance uptake via specific receptor interactions [37]. Importantly, by having a low percentage of surface amines labelled with rhodamine, these effects on interaction with the cells will be reduced. In a study looking at the required amount of particle surface functionalisation to enhance cellular uptake, it was found that at least 4% of the surface functional groups (maleimide) needed to be covered with wheat germ agglutinin (WGA) in order to significantly increase uptake of PEG-PLA nanoparticles into Calu-3 cells in comparison with non-functionalised particles [38].

Measurement of trypan blue quenching efficiency

To distinguish internalised particles from surface-bound particles, we used a protocol based on fluorescence quenching of surface-bound particles. In order for this method to be effective, the degree of fluorescence quenching must be known to ensure that surface-bound particles are significantly less fluorescent than internalised particles. To calculate this, the MFI of each particle type was measured before and after quenching with trypan blue (n > 1000 particles). The results are also displayed in Fig. S4 (1), showing that in each case the MFI was decreased after trypan blue quenching by over 90%, allowing distinction of internalised and surface-bound particles.

Microchip interaction with RAW 264.7 cells

Qualitative determination of microchips uptake

To examine if each of the differently shaped microchips could be internalised by RAW 264.7 macrophages, laser scanning confocal microscopy (LSCM) and immunostaining were used. Macrophages were incubated with fluorescently labelled particles of each shape for 4 h before being fixed and stained. BDP 630 was used in place of RBITC to fluorescently label the particles for this qualitative examination of particle uptake. Representative Z-projections are displayed along with their orthogonal views and 3D representations in Fig. 3. Cell nuclei, stained with DAPI, are shown in blue, actin filaments stained with iFluor 488 phalloidin are shown in green, and particles are displayed in red. Actin staining with phalloidin shows part of the cell cytoskeleton, either with cells appearing rounded or in some cases spread out with pseudopodia seen protruding from cells. Particles appear as hollow shapes in the orthogonal views (Fig. 3Bi, Bii, Ci, Cii, Di), because they are labelled only on their surface and so no fluorescence is seen inside the particles. In comparison with control cells that had not received any particle treatment (Fig. 3A), macrophages, which had been treated with particles, displayed similar morphologies and nuclear shapes. The actin cytoskeleton staining (green) can be used to determine if the particles have been internalised. In the orthogonal views, particles found within the green boundary of the cytoskeleton can be considered to have been internalised. These images show that each of the particle shapes had been internalised by RAW 264.7 macrophages. Particles were typically found in the perinuclear region of the cell which is in line with the established literature which details that upon phagocytosis, particles are contained within the phagosome which is transported towards the nucleus fusing with lysosomes to become a phagolysosome [39].Fig. 3 LSCM images of RAW 264.7 cells with internalised microchip shapes. i Z projections and orthogonal views, ii 3D reconstructions of cells treated with A 1% HEPES in HBSS only, B spheres, C cuboids, and D pyramids. Nuclear staining (blue), actin staining (green), and particles (red)

Intracellular localisation of spheres, cuboids, and pyramids

Immunostaining was used to determine the intracellular localisation of the particles after internalisation had taken place. Cells were incubated with particles for 4 h and then fixed and permeabilised. Cell nuclei were stained blue with DAPI, and lysosomal-associated membrane protein 1 (LAMP-1) antibody stain (green) was applied to stain lysosomes/late endosomes. Images of stained cells with each particle shape are displayed in Fig. 4 with particles displayed in red. In untreated cells (Fig. 4A), lysosomes/late endosomes were stained and appeared as small green dots inside the cell surrounding the nucleus. Lysosomes/late endosomes were not stained in all cells, and this could be because not all cells contained lysosomes/late endosomes or more likely due to a lack of staining, perhaps resulting from a lack of permeabilization. In cells to which particles had been applied, lysosomes/late endosomes could also be seen as small green dots throughout the cell. However, in cells that had internalised particles, a green ring was often observed surrounding the particle. This indicates that LAMP-1 was present in the membrane surrounding the particle. Upon maturation of the phagosome, lysosomes and endosomes fuse with the phagosomal membrane to deposit their contents. A ‘kiss-and-run’ mechanism has been proposed as a major mechanism for this, where complete fusion of the lysosomal/endosomal and phagosomal membranes is prevented by fission after some transfer of luminal contents has taken place [40]. Proteins such as LAMP-1 are left behind in the late phagosome/phagolysosome, thereby indicating that fusion has taken place [41]. The white arrows in Fig. 4C and D indicate particles that are surrounded by LAMP-1-positive membranes and are therefore contained within the late phagosome/phagolysosome. Particles that were clearly not internalised by cells displayed no such ring of green fluorescence, thus demonstrating that the green fluorescence results from the LAMP-1-positive membrane rather than an artefact of overlapping fluorescent emission or direct binding of the LAMP-1 antibody to the particle.Fig. 4 Intracellular localisation of particles after phagocytosis. Cells treated with A 1% HEPES in HBSS only, B spheres, C cuboids, and D pyramids. DAPI (blue), LAMP-1 (green), particles (red). White arrows indicate LAMP-1 colocalisation with particles

Also present are particles that had been internalised by cells, but that did not display a ring of fluorescence. It is therefore likely that these particles were inside the phagosome, and that endosome/lysosome-phagosome fusion had not yet taken place. No difference was seen between the different shapes in terms of localisation. As in the case of the actin staining, the majority of particles were situated in the perinuclear region of the cell. Images of at least 40 cells from each sample were examined and the percentage of particles surrounded by LAMP-1-positive membranes calculated. The percentage of spheres and pyramids was similar (60.9 and 53.3%, respectively); however, the percentage of cuboids surrounded by LAMP-1-positive membrane was lower (31.0%).

Quantitative determination of microchip uptake using imaging flow cytometry (IFC) and trypan blue quenching (TBQ) methods

Determination of microchip’s uptake in the absence and presence of LPS

RAW 264.7 macrophages were incubated with rhodamine B-labelled particle suspensions at particle:cell ratios of 5:1 and 10:1, with or without prior stimulation with LPS as macrophages are highly dynamic cells that exhibit phenotypic changes in response to environmental cues. The results are shown in Fig. S6. Determination of particle uptake was calculated in two ways. Firstly, the percentage of cells, which had internalised particles, was measured by counting the number of cells, which had a fluorescent intensity above the intensity of untreated cells and dividing by the total number of cells (Fig. S6A). Secondly, by fluorescent spot counting of cells with internal particles, the mean number of particles internalised per cell was calculated (Fig. S6B). The data in Fig. S6 shows the percentage of cells with internal particles indicates that there was no statistical difference between the two methods (IFC and TBQ) in any case. Owing to the way in which the IFC method discriminates internal from external particles (Figs. S7, S8, S9, S10, and S11 in supporting information), it might be expected that an over estimate of the percentage of cells with internal particles is observed. This is because although surface-bound particles can be discriminated in the X- and Y-planes, the depth of field in the Z-plane is not sufficient to distinguish whether particles are internal or surface bound. The TBQ method (Figures S12 and S13 in supporting information) does not suffer this drawback and instead relies on adequate fluorescence quenching to gate surface-bound particles. A study by Fei et al. [25] reported that it is not possible to suitably distinguish between internal- and surface-bound particles without the addition of a secondary fluorescent label. In contrast, a study by Vranic et al. [24] showed similar results between trypan blue quenching of fluorescein isothiocyanate (FITC)-labelled silica nanoparticles and imaging flow cytometry image analysis. Owing to the conflicting reports, it was necessary to assess the validity of this method for the particles used in this work by performing a comparison with a well-established trypan blue quenching method. The results of the current work agree with the study by Vranic et al. that showed equivalent uptake using TBQ and IFC [25]. A possible explanation for the lack of difference between the two methods is that the majority of particles are internalised rather than surface bound, and so the overestimation of surface-bound particles by the IFC method is minimised. Particle uptake at 4 °C was assessed using the IFC method (Fig. S14 in supporting information) and showed a reduction in particle uptake in comparison to 37 °C. Phagocytosis, an energy-dependent process, is reduced at this temperature, confirming that the IFC method was able to measure particle internalisation. The mean number of particles per cell was calculated and the data presented in Fig. S6B. In the majority of cases, IFC calculated a higher mean number of particles per cell than the TBQ method. This may result from the fact that surface-bound particles in cells that have both internal and external particles are included in the image analysis spot count but not in the trypan blue spot count (due to quenching).

RAW 264.7 cells were also stimulated with lipopolysaccharide (LPS) for 24 h prior to dosing in order to encourage differentiation to the M1 phenotype because macrophages are highly dynamic cells that exhibit phenotypic changes in response to environmental cues [42]. This was assessed visually by light microscopy as can be seen in Fig. S15 (supporting information). Cells that had undergone stimulation with LPS exhibited morphological changes as can been seen in Fig. S15B. In general, cells appeared larger and displayed vacuole-like and filopodia structures, indicative of M1-activated macrophages [43].

Particle uptake by cells was analysed using both IFC and TBQ, and the results are shown in Fig. S16 (supporting information). Figure S16A shows the effect of LPS stimulation on the percentage of cells that had internalised particles. LPS stimulation did not alter significantly the percentage of cells taking up particles. The only significant difference was observed in the case of spheres that were administered at a dose of 10 particles per cell and analysed using the TBQ method where LPS stimulation resulted in a significantly lower percentage of cells with internal particles. Similarly, there was very little difference in the average number of particles taken up per cell in LPS stimulated and unstimulated cells (Fig. S16B). The majority of cells internalised one or two particles with a much smaller proportion of cells internalising three or more particles. A study by Rieger et al. using IFC demonstrated that upon stimulation with LPS, RAW 264.7 macrophages association with microparticles increased overall; however, the cause for the increase in association was not an increase in internalisation but an increase in surface binding of zymosan particles [44].

If the same was true in the case of the silicon oxide particles used in this study, the increased binding of particles to the cell surface would not be detected by the TBQ method as these additional particles would be quenched. The IFC method can quantify this effect by analysing the percentage of cells with surface-bound particles as shown in Fig. 5. At a dose five particles per cell, the results indicate that approximately 30% of the cells have internalised spheres and cuboids. However, the percentage of cells with surface-bound spheres and cuboids is about 5% and 10%, respectively. In the case of pyramids, approximately 20% of cells take up the microchips, and about 5% of cells have surface-bound microchips. When the ratio of particles per cell was doubled, the percentage of cells with internalised spheres and cuboids increased to around 50% and that of pyramids to about 35%. On the other hand, the percentage of cells with surface-bound spheres and cuboids was ca. 5% and 10%, respectively, and that of pyramids to about 5%. Higher surface binding was not observed upon stimulation with LPS; instead, no statistically significant difference in overall particle association was observed in LPS-stimulated cells.Fig. 5 The effect of LPS stimulation on internalisation and surface binding for A 5 particles and B 10 particles administered per cells using the IFC method. *, **, ***, and **** indicate statistical significance (p < 0.05, 0.01, 0.001, 0.0001, respectively) as calculated by multiple unpaired T-tests. Values are representative of three biological repeats ± SEM (N = 3)

Effect of microchip shape on uptake by RAW 264.7 macrophages

Macrophages have previously been shown to display differential uptake of polymer nanoparticles depending on their morphology [45–47]. As such, it was expected that differences in the internalisation of spheres, cuboids, and pyramids would be observed. The results from analysing differences in internalisation between the different particle shapes using both IFC and TBQ methods are shown in Fig. 6. Using both analysis methods, an increase in the percentage of cells with internalised particles was observed as the particle dosage per cell doubled from 5 to 10 particles per cell (Fig. 6). This effect was seen in both LPS-stimulated and non-stimulated cells. However, the mean number of particles internalised per cell with the average of 1–1.5 particle/cell did not change with increasing the dose of particles per cell (Fig. S17). Using the IFC, the data is presented as the percentage of cells associated with particles and further distinguishes the percentage of cells with internal and surface-bound particles (Fig. 6Ai, ii). In this case, no significant differences between spheres and cuboids in terms of the percentage of cells associated with particles, the percentage of cells with internalised particles, or the percentage cells with surface-bound particles. This was the case for both LPS-stimulated and non-stimulated cells. In addition, analysis by TBQ shows no significant differences in internalisation of particles between spheres and cuboids (Fig. 6Bi, ii). However, significantly fewer cells internalised pyramids compared to spheres indicating that the ability of macrophages to internalise pyramids is lower. This may result from the anisotropic nature of these particles, which can present to cells in multiple orientations as opposed to spherical particles, which can only make contact with the cell in one orientation. Actin polymerisation must then occur in order to wrap the plasma membrane around the particle, and differences in the capacity of membrane wrapping, depending on particle orientation, could therefore result in reduced uptake.Fig. 6 The effect of particle shape on cellular uptake. The effect of particle shape on % of cells with internal particles Ai and Aii analysed using IFC and Bi and Bii analysed using TBQ method. *, **, ***, and **** indicate statistical significance (p < 0.05, 0.01, 0.001, 0.0001, respectively) as calculated by two-way ANOVA with multiple comparisons. Values are representative of three biological repeats ± SEM (n = 3)

Curiously, this effect was seen only in cells that had not been stimulated with LPS. There were no differences between any of the shapes in LPS-stimulated cells. Filopodia and membrane ruffles on the surface of macrophages are dynamic structures which have been shown to contribute to macropinocytosis and phagocytosis. Upon stimulation with LPS, actin polymerisation is increased resulting in more filopodia and membrane ruffles [48]. This change in the macrophage skeletal structure could therefore explain the reduced the effect of particle shape seen in LPS-stimulated cells. No differences were seen in the number of particles internalised per cell for any particle shape (Fig. S17). The majority of cells that internalised particles only internalised one particle regardless of particle shape, and this was consistent for both particles’ doses. In a similar study by Sharma et al. which looked at the internalisation of polystyrene spheres, oblate ellipsoids, and prolate ellipsoids by RAW 264.7 macrophages, there was no difference in internalisation seen between the different shapes with volumes above 24 μm3 (particles stretched from a 3.6-μm Φ sphere) [45]. There were, however, significant differences seen between the shapes that had volumes up to 0.69 μm3. The volumes of the particles used in this work were 14.1 μm3 (spheres), 11.6 μm3 (cuboids), and 17.2 μm3 (pyramids) and so are in between the volumes of particle groups used by Sharma et al. On the other hand, shape effects on internalisation of larger microparticles by macrophages have been reported by Doshi et al. who found that hyaluronic acid-coated disk-like microparticles (diameter = 6 μm) were able to avoid uptake by J774 macrophages compared with spherical particles (6 μm) of the same surface composition which exhibited high uptake [46]. Additionally, Champion and Mitragotri have shown that worm-like particles (aspect ratio > 20, length ~ 10 μm) are internalised significantly less that spheres of equivalent volume [47].

Conclusions

Silicon oxide microchips of different morphologies — either commercially available spheres or microfabricated rectangular cuboids and apex-truncated square pyramids — have been fluorescently labelled in order to study their interaction with living cells. Both surface- and in suspension-controlled covalent functionalization have resulted in a uniform, reduced — 2% of surface coverage per microchip — and stable labelling of the microchips. Imaging flow cytometry (IFC) has been used as a method for quantifying microparticle uptake, compared with the trypan blue quenching method and proved to give similar results demonstrating its usefulness as a technique which does not rely on specific quenching of fluorophores. The effect of particle shape on cellular uptake was evaluated. RAW 264.7 cells could phagocytose all particle shapes, which were shown to follow the established phagosomal maturation pathway. The uptake of particles increased with increasing dose and was not significantly altered upon stimulation of macrophages with LPS. There was a negligible difference in the uptake of particles between spheres and cuboids, regarding both the percentage of cells containing internalised particles and the number of particles internalised per cell. The findings reveal that at a rate microchip:cell of 5:1, ca. 30% of the cells internalise spheres and cuboids, with the average of 1–1.5 particle/cell, whereas the proportion of cells with surface-bound spheres and cuboids is approximately 5% and 10%, respectively. However, pyramids demonstrated a decreased uptake about 20% of cells engulf the microchips and the same average of 1–1.5 particle/cell, while around 5% have surface-bound microchips. When the particle dose was doubled to 10:1, an increase to ca. 50% of the percentage of cells with internalised spheres and cuboids was observed, whereas for pyramids it was ca. 35%. Moreover, the percentage of cells with surface-bound spheres and cuboids was approximately 5% and 10%, respectively, and for pyramids it was about 5%. However, the number of microchips internalised per cell did not change after increasing the dosage of particles, and it was ca. 1.5 microchip/cell, a remarkable value considering the overall size of the microchips.

Overall, the microfabricated cuboids show a comparable rate of internalisation to the widely studied and commonly used spheres. This suggests that cuboids could be viewed as superior choices for micro-carrier applications because they enable controlled multi-functionalization in terms of space, for example through micro-contact printing techniques. In addition, cuboid microchips offer several advantages over spherical particles, making them superior in biomedical applications. The microfabrication of cuboid microchips allows for precise size and shape control, avoiding the issue of polydispersity commonly seen with spherical particles. This precise control also enhances their versatility and morphology, enabling tailored designs for the biological applications. The high surface area of cuboid microchips supports high loading capacities and facilitates chemical functionalization, improving their functionality. Furthermore, cuboid microchips are biocompatible, easier to observe under microscopic analysis, and exhibit prolonged retention within cells, often accumulating in targeted tissues. These characteristics collectively make cuboid microchips a more effective and reliable option for biomedical applications.

Supplementary Information

Below is the link to the electronic supplementary material.Supplementary file1 (DOCX 5156 KB)

Acknowledgements

This work was supported by project PID2020-115663GB-C3 1 and 2 funded by Ministerio de Ciencia e Innovación/Agencia Española de Investigación MCIN/AEI/10.13039/501100011033. We also thank Agència de Gestió d'Ajuts Universitaris i de Recerca (AGAUR, Generalitat de Catalunya) for project 2021 SGR 01085. G. B. thanks EPSRC Grant EP/L01646X and GSK for a predoctoral grant. S. B. thanks Generalitat de Catalunya for a predoctoral FISDUR scholarship.

Author contribution

All authors contributed to the study conception and design. Material preparation, data collection, and analysis were performed by Gordon Bruce, as part of his doctoral thesis, Saman Bagherpour contributed to analysis, and Marta Duch to microfabrication. The first draft of the manuscript was written by Gordon Bruce, and all authors commented on previous versions of the manuscript. All authors read and approved the final manuscript.

Funding

Open Access funding provided thanks to the CRUE-CSIC agreement with Springer Nature. Ministerio de Ciencia e Innovación, PID2020-115663 GB-C3-1, José Antonio Plaza, PID2020-115663 GB-C3-2, Lluïsa Pérez-García, Agaur, 2021SGR01085, Lluïsa Pérez-García, FISDUR, Saman Bagherpour, Engineering and Physical Sciences Research Council, EP/L01646X,Snow Stolnik, GlaxoSmithKline

Data availability

The datasets generated during and/or analysed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethical approval.

This declaration is not applicable

Conflict of interest

The authors declare no competing interests.

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
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References

1. Fang L Micro- and nano-carrier systems: the non-invasive and painless local administration strategies for disease therapy in mucosal tissues Nanomedicine NBM 2017 13 153 171 10.1016/j.nano.2016.08.025
Fang L et al (2017) Micro- and nano-carrier systems: the non-invasive and painless local administration strategies for disease therapy in mucosal tissues. Nanomedicine NBM 13:153–171. 10.1016/j.nano.2016.08.02510.1016/j.nano.2016.08.025
2. Timin AS Litvak MM Gorin DA Atochina-Vasserman EN Atochin DN Sukhorukov GB Cell-based drug delivery and use of nano-and microcarriers for cell functionalization Adv Healthc Mater 2018 7 1700818 10.1002/adhm.201700818
Timin AS, Litvak MM, Gorin DA, Atochina-Vasserman EN, Atochin DN, Sukhorukov GB (2018) Cell-based drug delivery and use of nano-and microcarriers for cell functionalization. Adv Healthc Mater 7:1700818. 10.1002/adhm.20170081810.1002/adhm.201700818
3. Cooley M Sarode A Hoore M Fedosov DA Mitragotri S Sen Gupta A Influence of particle size and shape on their margination and wall-adhesion: implications in drug delivery vehicle design across nano-to-micro scale Nanoscale 2018 10 15350 15364 10.1039/C8NR04042G 30080212
Cooley M, Sarode A, Hoore M, Fedosov DA, Mitragotri S, Sen Gupta A (2018) Influence of particle size and shape on their margination and wall-adhesion: implications in drug delivery vehicle design across nano-to-micro scale. Nanoscale 10:15350–15364. 10.1039/C8NR04042G30080212 10.1039/C8NR04042G
4. Decuzzi P Pasqualini R Arap W Ferrari M Intravascular delivery of particulate systems: does geometry really matter? Pharm Res 2009 26 235 243 10.1007/s11095-008-9697-x 18712584
Decuzzi P, Pasqualini R, Arap W, Ferrari M (2009) Intravascular delivery of particulate systems: does geometry really matter? Pharm Res 26:235–243. 10.1007/s11095-008-9697-x18712584 10.1007/s11095-008-9697-x
5. Decuzzi P Godin B Tanaka T Lee S-Y Chiappini C Liu X Ferrari M Size and shape effects in the biodistribution of intravascularly injected particles J Control Release 2010 141 320 327 10.1016/j.jconrel.2009.10.014 19874859
Decuzzi P, Godin B, Tanaka T, Lee S-Y, Chiappini C, Liu X, Ferrari M (2010) Size and shape effects in the biodistribution of intravascularly injected particles. J Control Release 141:320–327. 10.1016/j.jconrel.2009.10.01419874859 10.1016/j.jconrel.2009.10.014
6. Schimmelpfeng J Drosselmeyer E Hofheinz V Seidel A Influence of surfactant components and exposure geometry on the effects of quartz and asbestos on alveolar macrophages Environ Health Perspect 1992 97 225 231 10.1289/ehp.9297225 1327736
Schimmelpfeng J, Drosselmeyer E, Hofheinz V, Seidel A (1992) Influence of surfactant components and exposure geometry on the effects of quartz and asbestos on alveolar macrophages. Environ Health Perspect 97:225–231. 10.1289/ehp.92972251327736 10.1289/ehp.9297225
7. Fenoglio I Croce A Di Renzo F Tiozzo R Fubini B Pure-silica zeolites (porosils) as model solids for the evaluation of the physicochemical features determining silica toxicity to macrophages Chem Res Toxicol 2000 13 489 500 10.1021/tx990169u 10858322
Fenoglio I, Croce A, Di Renzo F, Tiozzo R, Fubini B (2000) Pure-silica zeolites (porosils) as model solids for the evaluation of the physicochemical features determining silica toxicity to macrophages. Chem Res Toxicol 13:489–500. 10.1021/tx990169u10858322 10.1021/tx990169u
8. Hart GA Hesterberg TW In vitro toxicity of respirable-size particles of diatomaceous earth and crystalline silica compared with asbestos and titanium dioxide J Occup Environ Med 1998 40 29 42 10.1097/00043764-199801000-00008 9467118
Hart GA, Hesterberg TW (1998) In vitro toxicity of respirable-size particles of diatomaceous earth and crystalline silica compared with asbestos and titanium dioxide. J Occup Environ Med 40:29–429467118 10.1097/00043764-199801000-00008
9. Singh P Srivastava S Singh SK Nanosilica: recent progress in synthesis, functionalization, biocompatibility, and biomedical applications ACS Biomater Sci Eng 2019 5 4882 4898 10.1021/acsbiomaterials.9b00464 33455238
Singh P, Srivastava S, Singh SK (2019) Nanosilica: recent progress in synthesis, functionalization, biocompatibility, and biomedical applications. ACS Biomater Sci Eng 5:4882–4898. 10.1021/acsbiomaterials.9b0046433455238 10.1021/acsbiomaterials.9b00464
10. Encinas N Angulo M Astorga C Colilla M Izquierdo-Barba I Vallet-Regí M Mixed-charge pseudo-zwitterionic mesoporous silica nanoparticles with low-fouling and reduced cell uptake properties Acta Biomater 2019 84 317 327 10.1016/j.actbio.2018.12.012 30529082
Encinas N, Angulo M, Astorga C, Colilla M, Izquierdo-Barba I, Vallet-Regí M (2019) Mixed-charge pseudo-zwitterionic mesoporous silica nanoparticles with low-fouling and reduced cell uptake properties. Acta Biomater 84:317–327. 10.1016/j.actbio.2018.12.01230529082 10.1016/j.actbio.2018.12.012
11. Manzano M Vallet-Regí M Mesoporous silica nanoparticles for drug delivery Adv Funct Mater 2020 30 1902634 10.1002/adfm.201902634
Manzano M, Vallet-Regí M (2020) Mesoporous silica nanoparticles for drug delivery. Adv Funct Mater 30:1902634. 10.1002/adfm.20190263410.1002/adfm.201902634
12. Huang Y Silica nanoparticles: biomedical applications and toxicity Biomed Pharmacother 2022 151 113053 10.1016/j.biopha.2022.113053 35594717
Huang Y et al (2022) Silica nanoparticles: biomedical applications and toxicity. Biomed Pharmacother 151:113053. 10.1016/j.biopha.2022.11305335594717 10.1016/j.biopha.2022.113053
13. Kettler K Veltman K van de Meent D van Wezel A Hendriks AJ Cellular uptake of nanoparticles as determined by particle properties, experimental conditions, and cell type Environ Toxicol Chem 2014 33 481 492 10.1002/etc.2470 24273100
Kettler K, Veltman K, van de Meent D, van Wezel A, Hendriks AJ (2014) Cellular uptake of nanoparticles as determined by particle properties, experimental conditions, and cell type. Environ Toxicol Chem 33:481–492. 10.1002/etc.247024273100 10.1002/etc.2470
14. Lu F Wu S-H Hung Y Mou C-Y Size effect on cell uptake in well-suspended, uniform mesoporous silica nanoparticles Small 2009 5 1408 1413 10.1002/smll.200900005 19296554
Lu F, Wu S-H, Hung Y, Mou C-Y (2009) Size effect on cell uptake in well-suspended, uniform mesoporous silica nanoparticles. Small 5:1408–1413. 10.1002/smll.20090000519296554 10.1002/smll.200900005
15. Kurtz-Chalot A Villiers C Pourchez J Boudard D Martini M Marche PN Cottier M Forest V Impact of silica nanoparticle surface chemistry on protein corona formation and consequential interactions with biological cells Mater Sci Eng C 2017 75 16 24 10.1016/j.msec.2017.02.028
Kurtz-Chalot A, Villiers C, Pourchez J, Boudard D, Martini M, Marche PN, Cottier M, Forest V (2017) Impact of silica nanoparticle surface chemistry on protein corona formation and consequential interactions with biological cells. Mater Sci Eng C 75:16–24. 10.1016/j.msec.2017.02.02810.1016/j.msec.2017.02.028
16. Hao N Li L Zhang Q Huang X Meng X Zhang Y Chen D Tang F Li L The shape effect of PEGylated mesoporous silica nanoparticles on cellular uptake pathway in HeLa cells Microporous Mesoporous Mater 2012 162 14 23 10.1016/j.micromeso.2012.05.040
Hao N, Li L, Zhang Q, Huang X, Meng X, Zhang Y, Chen D, Tang F, Li L (2012) The shape effect of PEGylated mesoporous silica nanoparticles on cellular uptake pathway in HeLa cells. Microporous Mesoporous Mater 162:14–23. 10.1016/j.micromeso.2012.05.04010.1016/j.micromeso.2012.05.040
17. Karaman D Sen Desai D Senthilkumar R Johansson EM Råtts N Odén M Eriksson JE Sahlgren C Toivola DM Rosenholm JM Shape engineering vs organic modification of inorganic nanoparticles as a tool for enhancing cellular internalization Nanoscale Res. Lett 2012 7 358 10.1186/1556-276X-7-358 22747910
Karaman D, Sen Desai D, Senthilkumar R, Johansson EM, Råtts N, Odén M, Eriksson JE, Sahlgren C, Toivola DM, Rosenholm JM (2012) Shape engineering vs organic modification of inorganic nanoparticles as a tool for enhancing cellular internalization. Nanoscale Res. Lett 7:358. 10.1186/1556-276X-7-35822747910 10.1186/1556-276X-7-358
18. Breznan D Das DD O’Brien JS MacKinnon-Roy C Nimesh S Vuong NQ Bernatchez S DeSilva N Hill M Kumarathasan P Differential cytotoxic and inflammatory potency of amorphous silicon dioxide nanoparticles of similar size in multiple cell lines Nanotoxicology 2017 11 223 235 10.1080/17435390.2017.1287313 28142331
Breznan D, Das DD, O’Brien JS, MacKinnon-Roy C, Nimesh S, Vuong NQ, Bernatchez S, DeSilva N, Hill M, Kumarathasan P et al (2017) Differential cytotoxic and inflammatory potency of amorphous silicon dioxide nanoparticles of similar size in multiple cell lines. Nanotoxicology 11:223–235. 10.1080/17435390.2017.128731328142331 10.1080/17435390.2017.1287313
19. Yazdimamaghani M Barber ZB Hadipour Moghaddam SP Ghandehari H Influence of silica nanoparticle density and flow conditions on sedimentation, cell uptake, and cytotoxicity Mol Pharm 2018 15 2372 2383 10.1021/acs.molpharmaceut.8b00213 29719153
Yazdimamaghani M, Barber ZB, Hadipour Moghaddam SP, Ghandehari H (2018) Influence of silica nanoparticle density and flow conditions on sedimentation, cell uptake, and cytotoxicity. Mol Pharm 15:2372–2383. 10.1021/acs.molpharmaceut.8b0021329719153 10.1021/acs.molpharmaceut.8b00213
20. Ferreira MPA Ranjan S Correia AMR Mäkilä EM Kinnunen SM Zhang H Shahbazi M-A Almeida PV Salonen JJ Ruskoaho HJ In vitro and in vivo assessment of heart-homing porous silicon nanoparticles Biomaterials 2016 94 93 104 10.1016/j.biomaterials.2016.03.046 27107168
Ferreira MPA, Ranjan S, Correia AMR, Mäkilä EM, Kinnunen SM, Zhang H, Shahbazi M-A, Almeida PV, Salonen JJ, Ruskoaho HJ et al (2016) In vitro and in vivo assessment of heart-homing porous silicon nanoparticles. Biomaterials 94:93–104. 10.1016/j.biomaterials.2016.03.04627107168 10.1016/j.biomaterials.2016.03.046
21. Elsaesser A Taylor A de Yanés GS McKerr G Kim E-M O’Hare E Howard CV Quantification of nanoparticle uptake by cells using microscopical and analytical techniques Nanomedicine 2010 5 1447 1457 10.2217/nnm.10.118 21128725
Elsaesser A, Taylor A, de Yanés GS, McKerr G, Kim E-M, O’Hare E, Howard CV (2010) Quantification of nanoparticle uptake by cells using microscopical and analytical techniques. Nanomedicine 5:1447–1457. 10.2217/nnm.10.11821128725 10.2217/nnm.10.118
22. Lee J-Y Kitaoka M A beginner’s guide to rigor and reproducibility in fluorescence imaging experiments Mol Biol Cell 2018 29 1519 1525 10.1091/mbc.E17-05-0276 29953344
Lee J-Y, Kitaoka M (2018) A beginner’s guide to rigor and reproducibility in fluorescence imaging experiments. Mol Biol Cell 29:1519–1525. 10.1091/mbc.E17-05-027629953344 10.1091/mbc.E17-05-0276
23. Ashraf S Hassan Said A Hartmann R Assmann M-A Feliu N Lenz P Parak WJ Quantitative particle uptake by cells as analyzed by different methods Angew Chemie Int Ed 2020 59 5438 5453 10.1002/anie.201906303
Ashraf S, Hassan Said A, Hartmann R, Assmann M-A, Feliu N, Lenz P, Parak WJ (2020) Quantitative particle uptake by cells as analyzed by different methods. Angew Chemie Int Ed 59:5438–5453. 10.1002/anie.20190630310.1002/anie.201906303
24. Mikami H Kawaguchi M Huang C-J Matsumura H Sugimura T Huang K Lei C Ueno S Miura T Ito T Virtual-freezing fluorescence imaging flow cytometry Nat Commun 2020 11 1162 10.1038/s41467-020-14929-2 32139684
Mikami H, Kawaguchi M, Huang C-J, Matsumura H, Sugimura T, Huang K, Lei C, Ueno S, Miura T, Ito T et al (2020) Virtual-freezing fluorescence imaging flow cytometry. Nat Commun 11:1162. 10.1038/s41467-020-14929-232139684 10.1038/s41467-020-14929-2
25. Vranic S Boggetto N Contremoulins V Mornet S Reinhardt N Marano F Baeza-Squiban A Boland S Deciphering the mechanisms of cellular uptake of engineered nanoparticles by accurate evaluation of internalization using imaging flow cytometry Part Fibre Toxicol 2013 10 2 10.1186/1743-8977-10-2 23388071
Vranic S, Boggetto N, Contremoulins V, Mornet S, Reinhardt N, Marano F, Baeza-Squiban A, Boland S (2013) Deciphering the mechanisms of cellular uptake of engineered nanoparticles by accurate evaluation of internalization using imaging flow cytometry. Part Fibre Toxicol 10:2. 10.1186/1743-8977-10-223388071 10.1186/1743-8977-10-2
26. Fei C Lillico DME Hall B Rieger AM Stafford JL Connected component masking accurately identifies the ratio of phagocytosed and cell-bound particles in individual cells by imaging flow cytometry Cytom Part A 2017 91 372 381 10.1002/cyto.a.23050
Fei C, Lillico DME, Hall B, Rieger AM, Stafford JL (2017) Connected component masking accurately identifies the ratio of phagocytosed and cell-bound particles in individual cells by imaging flow cytometry. Cytom Part A 91:372–381. 10.1002/cyto.a.2305010.1002/cyto.a.23050
27. Torras N Agusil JP Vázquez P Duch M Hernández-Pinto AM Samitier J De La Rosa EJ Esteve J Suárez T Pérez-García L Suspended planar-array chips for molecular multiplexing at the microscale Adv Mater 2016 28 1449 1454 10.1002/adma.201504164 26649987
Torras N, Agusil JP, Vázquez P, Duch M, Hernández-Pinto AM, Samitier J, De La Rosa EJ, Esteve J, Suárez T, Pérez-García L et al (2016) Suspended planar-array chips for molecular multiplexing at the microscale. Adv Mater 28:1449–1454. 10.1002/adma.20150416426649987 10.1002/adma.201504164
28. Agusil JP Torras N Duch M Esteve J Pérez-García L Samitier J Plaza JA Highly anisotropic suspended planar-array chips with multidimensional sub-micrometric biomolecular patterns Adv Funct Mater 2017 27 1605912 10.1002/adfm.201605912
Agusil JP, Torras N, Duch M, Esteve J, Pérez-García L, Samitier J, Plaza JA (2017) Highly anisotropic suspended planar-array chips with multidimensional sub-micrometric biomolecular patterns. Adv Funct Mater 27:1605912. 10.1002/adfm.20160591210.1002/adfm.201605912
29. Arjona MI Duch M Hernández-Pinto A Vázquez P Agusil JP Gómez-Martínez R Redondo-Horcajo M Amirthalingam E Pérez-García L Suárez T Intracellular mechanical drugs induce cell-cycle altering and cell death Adv Mater 2022 34 2109581 10.1002/adma.202109581
Arjona MI, Duch M, Hernández-Pinto A, Vázquez P, Agusil JP, Gómez-Martínez R, Redondo-Horcajo M, Amirthalingam E, Pérez-García L, Suárez T et al (2022) Intracellular mechanical drugs induce cell-cycle altering and cell death. Adv Mater 34:2109581. 10.1002/adma.20210958110.1002/adma.202109581
30. Durán S Duch M Patiño T Torres A Penon O Gómez-Martínez R Barrios L Esteve J Nogués C Pérez-García L Technological development of intracellular polysilicon–chromium–gold chips for orthogonal chemical functionalization Sensors Actuators B Chem 2015 209 212 224 10.1016/j.snb.2014.11.077
Durán S, Duch M, Patiño T, Torres A, Penon O, Gómez-Martínez R, Barrios L, Esteve J, Nogués C, Pérez-García L et al (2015) Technological development of intracellular polysilicon–chromium–gold chips for orthogonal chemical functionalization. Sensors Actuators B Chem 209:212–224. 10.1016/j.snb.2014.11.07710.1016/j.snb.2014.11.077
31. Limón D Hornick JE Cai K Beldjoudi Y Duch M Plaza JA Pérez-García L Stoddart JF Polysilicon microchips functionalized with bipyridinium-based cyclophanes for a highly efficient cytotoxicity in cancerous cells ACS Nano 2022 16 5358 5375 10.1021/acsnano.1c08090 35357125
Limón D, Hornick JE, Cai K, Beldjoudi Y, Duch M, Plaza JA, Pérez-García L, Stoddart JF (2022) Polysilicon microchips functionalized with bipyridinium-based cyclophanes for a highly efficient cytotoxicity in cancerous cells. ACS Nano 16:5358–5375. 10.1021/acsnano.1c0809035357125 10.1021/acsnano.1c08090
32. Liberman A Mendez N Trogler WC Kummel AC Synthesis and surface functionalization of silica nanoparticles for nanomedicine Surf Sci Rep 2014 69 132 158 10.1016/j.surfrep.2014.07.001 25364083
Liberman A, Mendez N, Trogler WC, Kummel AC (2014) Synthesis and surface functionalization of silica nanoparticles for nanomedicine. Surf Sci Rep 69:132–158. 10.1016/j.surfrep.2014.07.00125364083 10.1016/j.surfrep.2014.07.001
33. Giraud L Nadarajah R Matar Y Bazin G Sun J Zhu XX Giasson S Amino-functionalized monolayers covalently grafted to silica-based substrates as a robust primer anchorage in aqueous media Appl Surf Sci 2016 370 476 485 10.1016/j.apsusc.2016.02.141
Giraud L, Nadarajah R, Matar Y, Bazin G, Sun J, Zhu XX, Giasson S (2016) Amino-functionalized monolayers covalently grafted to silica-based substrates as a robust primer anchorage in aqueous media. Appl Surf Sci 370:476–485. 10.1016/j.apsusc.2016.02.14110.1016/j.apsusc.2016.02.141
34. Perera HJ Mortazavian H Blum FD Surface properties of silane-treated diatomaceous earth coatings: effect of alkyl chain length Langmuir 2017 33 2799 2809 10.1021/acs.langmuir.7b00015 28244761
Perera HJ, Mortazavian H, Blum FD (2017) Surface properties of silane-treated diatomaceous earth coatings: effect of alkyl chain length. Langmuir 33:2799–2809. 10.1021/acs.langmuir.7b0001528244761 10.1021/acs.langmuir.7b00015
35. Tabata Y Ikada Y Effect of surface wettability of microspheres on phagocytosis J Colloid Interface Sci 1989 127 132 140 10.1016/0021-9797(89)90013-1
Tabata Y, Ikada Y (1989) Effect of surface wettability of microspheres on phagocytosis. J Colloid Interface Sci 127:132–140. 10.1016/0021-9797(89)90013-110.1016/0021-9797(89)90013-1
36. Yoo H Juliano RL Enhanced delivery of antisense oligonucleotides with fluorophore-conjugated PAMAM dendrimers Nucleic Acids Res 2000 28 4225 4231 10.1093/nar/28.21.4225 11058121
Yoo H, Juliano RL (2000) Enhanced delivery of antisense oligonucleotides with fluorophore-conjugated PAMAM dendrimers. Nucleic Acids Res 28:4225–4231. 10.1093/nar/28.21.422511058121 10.1093/nar/28.21.4225
37. Yan X Poelstra K Scherphof GL Kamps JAAM A role for scavenger receptor B-I in selective transfer of rhodamine-PE from liposomes to cells Biochem Biophys Res Commun 2004 325 908 914 10.1016/j.bbrc.2004.10.104 15541376
Yan X, Poelstra K, Scherphof GL, Kamps JAAM (2004) A role for scavenger receptor B-I in selective transfer of rhodamine-PE from liposomes to cells. Biochem Biophys Res Commun 325:908–914. 10.1016/j.bbrc.2004.10.10415541376 10.1016/j.bbrc.2004.10.104
38. Shen Y Chen J Liu Q Feng C Gao X Wang L Zhang Q Jiang X Effect of wheat germ agglutinin density on cellular uptake and toxicity of wheat germ agglutinin conjugated PEG–PLA nanoparticles in Calu-3 cells Int J Pharm 2011 413 184 193 10.1016/j.ijpharm.2011.04.026 21550388
Shen Y, Chen J, Liu Q, Feng C, Gao X, Wang L, Zhang Q, Jiang X (2011) Effect of wheat germ agglutinin density on cellular uptake and toxicity of wheat germ agglutinin conjugated PEG–PLA nanoparticles in Calu-3 cells. Int J Pharm 413:184–193. 10.1016/j.ijpharm.2011.04.02621550388 10.1016/j.ijpharm.2011.04.026
39. Keller S Berghoff K Kress H Phagosomal transport depends strongly on phagosome size Sci Rep 2017 7 17068 10.1038/s41598-017-17183-7 29213131
Keller S, Berghoff K, Kress H (2017) Phagosomal transport depends strongly on phagosome size. Sci Rep 7:17068. 10.1038/s41598-017-17183-729213131 10.1038/s41598-017-17183-7
40. Desjardins M Biogenesis of phagolysosomes: the ‘kiss and run’ hypothesis Trends Cell Biol 1995 5 183 186 10.1016/0962-8924(95)80001-W 14731444
Desjardins M (1995) Biogenesis of phagolysosomes: the ‘kiss and run’ hypothesis. Trends Cell Biol 5:183–186. 10.1016/0962-8924(95)80001-W14731444 10.1016/0962-8924(95)80001-W
41. Huynh KK Eskelinen E-L Scott CC Malevanets A Saftig P Grinstein S LAMP proteins are required for fusion of lysosomes with phagosomes EMBO J 2007 26 313 324 10.1038/sj.emboj.7601511 17245426
Huynh KK, Eskelinen E-L, Scott CC, Malevanets A, Saftig P, Grinstein S (2007) LAMP proteins are required for fusion of lysosomes with phagosomes. EMBO J 26:313–324. 10.1038/sj.emboj.760151117245426 10.1038/sj.emboj.7601511
42. Mantovani A Sica A Sozzani S Allavena P Vecchi A Locati M The chemokine system in diverse forms of macrophage activation and polarization Trends Immunol 2004 25 677 686 10.1016/j.it.2004.09.015 15530839
Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M (2004) The chemokine system in diverse forms of macrophage activation and polarization. Trends Immunol 25:677–686. 10.1016/j.it.2004.09.01515530839 10.1016/j.it.2004.09.015
43. Kim EJ Lee MY Jeon YJ Silymarin inhibits morphological changes in LPS-stimulated macrophages by blocking NF-ΚB pathway Kjpp 2015 19 211 218 10.4196/kjpp.2015.19.3.211
Kim EJ, Lee MY, Jeon YJ (2015) Silymarin inhibits morphological changes in LPS-stimulated macrophages by blocking NF-ΚB pathway. Kjpp 19:211–218. 10.4196/kjpp.2015.19.3.21110.4196/kjpp.2015.19.3.211
44. Rieger AM Hall BE Barreda DR Macrophage activation differentially modulates particle binding, phagocytosis and downstream antimicrobial mechanisms Dev Comp Immunol 2010 34 1144 1159 10.1016/j.dci.2010.06.006 20600280
Rieger AM, Hall BE, Barreda DR (2010) Macrophage activation differentially modulates particle binding, phagocytosis and downstream antimicrobial mechanisms. Dev Comp Immunol 34:1144–1159. 10.1016/j.dci.2010.06.00620600280 10.1016/j.dci.2010.06.006
45. Sharma G Valenta DT Altman Y Harvey S Xie H Mitragotri S Smith JW Polymer particle shape independently influences binding and internalization by macrophages J Control Release 2010 147 408 412 10.1016/j.jconrel.2010.07.116 20691741
Sharma G, Valenta DT, Altman Y, Harvey S, Xie H, Mitragotri S, Smith JW (2010) Polymer particle shape independently influences binding and internalization by macrophages. J Control Release 147:408–412. 10.1016/j.jconrel.2010.07.11620691741 10.1016/j.jconrel.2010.07.116
46. Doshi N Swiston AJ Gilbert JB Alcaraz ML Cohen RE Rubner MF Mitragotri S Cell-based drug delivery devices using phagocytosis-resistant backpacks Adv Mater 2011 23 H105 H109 10.1002/adma.201004074 21365691
Doshi N, Swiston AJ, Gilbert JB, Alcaraz ML, Cohen RE, Rubner MF, Mitragotri S (2011) Cell-based drug delivery devices using phagocytosis-resistant backpacks. Adv Mater 23:H105–H109. 10.1002/adma.20100407421365691 10.1002/adma.201004074
47. Champion JA Mitragotri S Shape induced inhibition of phagocytosis of polymer particles Pharm Res 2009 26 244 249 10.1007/s11095-008-9626-z 18548338
Champion JA, Mitragotri S (2009) Shape induced inhibition of phagocytosis of polymer particles. Pharm Res 26:244–249. 10.1007/s11095-008-9626-z18548338 10.1007/s11095-008-9626-z
48. Freeman SA Grinstein S Phagocytosis: receptors, signal integration, and the cytoskeleton Immunol Rev 2014 262 193 215 10.1111/imr.12212 25319336
Freeman SA, Grinstein S (2014) Phagocytosis: receptors, signal integration, and the cytoskeleton. Immunol Rev 262:193–215. 10.1111/imr.1221225319336 10.1111/imr.12212
