Macrophage responses to the physical burden of cell-sized particles

Hua Yue a, Lan Yuan b, Weiwei Zhang c, Shujia Zhang c, Wei Wei *a and Guanghui Ma *ade
aState Key Laboratory of Biochemical Engineering, Institute of Process Engineering, Chinese Academy of Sciences, Beijing 100190, China. E-mail:;
bPeking University Medical and Health Analysis Center, Beijing 100191, China
cCollege of Environment and Chemical Engineering, Dalian University, Liaoning Dalian 116622, China
dUniversity of Chinese Academy of Sciences, Beijing 100049, China
eJiangsu National Synergetic Innovation Center for Advanced Materials, Nanjing 211816, China

Received 19th June 2017 , Accepted 2nd December 2017

First published on 4th December 2017

The role of a biophysical signal on cell response has excited tremendous interest recently. Herein, we exploited an “intake method” together with uniform autofluorescent cell-sized particles to investigate macrophage responses against different particle burdens. Our work not only revealed an insatiable macrophage uptake of cell-sized microparticles (MPs) but also widened the theoretical size and volume range for particle entry. MPs outperform NPs in the utmost volume of intracellular particles, indicating a converse size-associated event than originally anticipated. Such a superior volume burden (2.64-fold of the cell volume) for MPs is highly correlated to less membrane loss (19-fold below that for NPs) and more evident deformation of the cell membrane/nucleus. In addition, the cells with a high burden of MPs exhibit moderate migration activity, which is in line with mild cytokine release. These results highlight the indispensable role of physical burden on the regulation of macrophage functions, providing important views for desired biological outcomes and minimal side effects.


Macrophages reside in every tissue and are the “first responders” to eat principal ‘targets’ such as microbes, all injected particles, and cells (senescing, dead or cancer cells).1 When macrophages encounter natural or man-made particles, their interactions are the basic topic in a variety of fields, such as pharmacology, cell biology, toxicology, and tissue engineering.2 Besides the assessment of cytotoxicity, in-depth studies on macrophage scavenging, activation and immunoregulation are highly appreciated.3,4 For example, particles with a smaller size (ranging from 20 nm to 1 μm),5,6 positive charge,7,8 hydrophobic surface,9,10 and specific chemical component11,12 are favored for macrophage activation or efficient immunity. However, prior to reaching the cell/tissue target, rapid clearance or a fierce cytokine response against the particles might result in low bioavailability or inflammation diseases.13,14 Since the desired biological outcomes vary case by case,15–18 there is still a rising demand for exploring the underlying bio-particle interactions.

To date, the majority of studies investigating the biological outcomes have focused on the particles in a nanosize scale, and systematic clarification upon particle burden lags far behind. It has been well documented that nanoparticles (NPs) could be more readily recognized and taken up into cells, as their size is similar to that of a virus or bacteria. Although NPs are associated with regulated cell viability, cytokine secretion and cell migration,19 the acquired conclusions may not be directly applicable to the microparticles (MPs). In contrast, only few and scattered documents have mentioned the cell uptake capacity of MPs up to 5 μm, let alone reported on the cell-sized MPs (at a size ranging from host immune cells to tumor cells).20,21 The maximum level of particle burden is largely unknown, and the observation of a resultant biological outcome is lacking. Therefore, it is necessary to uncover the cell uptake capacity and the resultant cell response against MPs at different particle burdens.

Particle burden inside the cells actually belongs to a physical cue. While the role of chemical signals has long been recognized, the importance of biophysical signals has only recently been recognized to regulate cell responses.22 The widely reported biophysical cue is surface topography, which reveals cellular behaviors in a well-defined geometry. Nevertheless, this is an exterior method, and it is difficult to isolate the influence of internal cell components on the whole-cell response.23 In addition, the well-defined geometry may not represent the natural random niche for macrophages. Particle burden refers to an interior signal and a physical environment along with the spontaneous arrangement of particles and cell organelles. Under a confined intracellular space, how the burden signals transduce through the whole cell to alter the cell geometry and resultant cell function is of great interest and importance.

Given the aforementioned knowledge gaps, we sought to determine the effect of physical burden on a series of macrophage responses. To overcome the handicap that resulted from the limited means of investigation, chitosan particles with uniform size (8 μm and 400 nm) as well as remarkable auto-fluorescence were fabricated using a microporous emulsification technique. Notably, the fluorescence intensity of the particles has been previously demonstrated to be directly proportional to the particle volume and quality.20,24 Such a combination of techniques highly facilitated the quantification analysis and ensured the reliability of the present study. Our results reveal that peritoneal macrophages (a model macrophage type) are capable of internalizing the 8 μm particles (up to a number of 10) with a total volume much larger than the volume of the cell and NPs. This event is highly dependent on the cell adjustment capability of the membrane and nucleus within the limited intracellular space. Besides, the subcellular organelle activity and cell migration behavior are also varied with different burden levels. This work highlights the indispensable role of a physical signal transduced from the particle burden to the whole cell in altering the macrophage response.

Results and discussion

Micro/nano particles prepared by conventional methods usually have a broad size distribution, which compromise subsequent biological evaluation and further applications in biomedical areas. To control the particle size ranging from nano to micro, we employed a microporous membrane emulsification technique that was developed in our research group. By choosing a membrane with a specific pore size, the MPs at 8 μm and the NPs (400 nm, termed as control particles) were successfully prepared with narrow size distributions (Fig. 1 and Fig. S1, ESI). Without any staining process, brilliant auto-fluorescence for the particles could be obtained, which was easily observed using laser confocal microscopy (CLSM). Meanwhile, the linear relationship between the fluorescence intensity (FI) and the volume of chitosan particles was again verified (Fig. S2, ESI). Employment of such bright and uniform particles not only guaranteed convincing results but also tremendously facilitated imaging and quantification.
image file: c7tb01673e-f1.tif
Fig. 1 Scanning electron microscopy (SEM) image (A), laser confocal microscopy (CLSM) image (B) and the size distribution (C) showing the uniform-sized microparticles (MPs, 8 μm) prepared by the membrane emulsification method. Scale bars: (A) 2 μm and (B) 25 μm. The inset scheme in (C) shows the principle of the membrane emulsification process.

Endocytosis by macrophages is considered a prerequisite for the process of exogenous substances and the subsequent regulation of an immunological response. To test the cell entry feasibility of gigantic man-made particles, auto-fluorescent MPs (8 μm) at different concentrations were added to the peritoneal macrophages (with an average diameter of 12 μm) (Fig. 2). By utilizing three imaging methods, an evident engulfment of particles at a size range around that of the host/tumor cells was systematically revealed. The scanning electron microscopy (SEM) images exhibited a series of internalization processes, including the initial filopodia exploration and capture, the subsequent membrane spreading and wrapping of particles, and the final fascinating morphology change of the cells. Notably, the particles were closely surrounded by the cell membrane, with a prominent outward curve that described the shape of internalized MPs. With the elevated particle concentration, macrophages (with a red membrane and green nucleus in the CLSM images) engulfed the MPs (with an auto-fluorescent yellow color) at a number of 1 to 10 (Fig. 2A–C and Fig. S3, ESI), reinforcing the huge appetite of macrophages under a confined cell volume. With an increase in the MP number, the membrane contour appears more visible, indicating an ingenious modulation for accommodation of MPs. The transmission electron microscopy (TEM) images also clearly showed the arrangement of the particles’ inner cells as well as the interaction between the cells and particles.

image file: c7tb01673e-f2.tif
Fig. 2 SEM, CLSM and TEM (transmission electron microscopy) images showing the response of the cell membrane and nucleus, upon the burden of the MPs at low (A), middle (B), and high (C) levels after 8 h of interaction. The morphologies of the cell membrane (red – rhodamine phalloidin) and nucleus (green – 4′,6-diamidino-2-phenylindole) in the CLSM images were strikingly changed with the increased burden of particles. Scale bars: 5 μm. (D) Schematic graph of the cells before and after the uptake of MPs.

In virtue of the fluorescence obtained from the internalized particles, the internalization profile was quantified by a flow cytometry assay (FCM), and a dose dependent uptake event for MPs was revealed (Fig. 3A). As the FI was in direct proportion to the internalized particle volume (Fig. S2, ESI), three burden levels (low: FI ≤ 200; middle: FI = 200–500; high: FI > 500) were defined, and the typical FI point for each burden (MP-L, MP-M, and MP-H) was highlighted by a colored arrow. After calculation (Fig. S4, ESI), the MP number (n) for the different burdens (low, middle and high) could be further set as 1–2, 3–5, and ≥6, respectively. After exposure to the corresponding concentration of the MPs for each burden, the cells exhibited time-dependent kinetics of FI (Fig. S5, ESI), and most of the internalization (>80%) occurred within the first 8 h. Besides, the side scattered light signals (SS) were enhanced, also reflecting an increased number of particles within the cells at an elevated burden level (Fig. S6, ESI). For the NP group, a dose dependent internalization was present at lower concentrations (≤80 μg mL−1), following a relatively feeble saturation plateau (Fig. 3B). Notably, the saturation FI of the internalized NPs only reached the middle burden level (NP-M), which was significantly below that for MPs.

image file: c7tb01673e-f3.tif
Fig. 3 The cellular internalization events (A–C) and cell viability (D and E) under different particle burdens. Cellular internalization of MPs (A) and NPs (B) at different particle concentrations after 24 h of incubation. The particle burden levels were defined according to the fluorescence intensity (FI) of the internalized particles. The corresponding MP numbers (n) are 1–2, 3–5, and ≥6, respectively, and a typical FI point for each burden (MP-L, MP-M, and MP-H) is highlighted by a colored arrow. Notably, the FI for the internalized NPs is not significantly elevated at a concentration above 80 μg, which only touches the middle burden level (NP-M). (C) Internalization phase diagram with calculated membrane loss (S*, the ratio of total particle surface to cell membrane surface) and the volume magnification (V*, the ratio of total particle volume to cell volume). The typical burden levels of low (L), middle (M), and high (L) were also marked by colored arrows. At the same volume burden (V*), the calculated membrane loss S* for NPs (400 nm) was 19-fold above that for MPs, which requires an enormous membrane supplement (e.g. the S* is 79.4 at supposed high NP burden as indicated by the hollow square). (D) Cell counting kit-8 (CCK-8) assay of cells after incubation with MPs at high burden and NPs at middle burden. (E) Lactate dehydrogenase (LDH) leakage of a positive control (P-ctrl), negative control (N-ctrl), and cell group at different burden levels after 48 h particle incubation. Each experiment was performed in triplicate, and data were represented as means ± standard deviation. P* < 0.05, P** < 0.01.

The above results confirmed a formidable internalization capacity of MPs and an utmost uptake volume burden that outperformed the NPs. Such an event indicated the different role of size in cell uptake than originally anticipated. On the one hand, the cell-sized particles were found to be insatiably phagocytized by macrophages, thus greatly widening the size range of particle entry and possible bio-applications. On the other hand, the particle burden capacity was a conversely size-associated event, while nanosize was unexpectedly inferior to microsize. In this sense, the micro-sized particles should not be overlooked either in biomedical strategies or in human health maintenance. For example, MPs present the potential to deliver active substances via an encapsulation method, as the MPs are superior in the total particle volume accumulated in cells. This is particularly applicable for implosive therapy or disease diagnosis when administrating a high dose of drug or radiation. Meanwhile, the insatiable MP uptake by macrophages in the lungs or mucosa may help to clear hazardous particles and maintain health when exposed to high levels of particulate matter (PM2.5 or 10). A similar outcome can also be predicted for macrophages in clearing senescing or even cancer cells.

To figure out the underlying mechanism for different burden capacities, we checked the intermediate status of the macrophages when they came across the particles. Intriguingly, every internalized microparticle was surrounded by a red cell membrane in the CLSM images (Fig. 2), and a similar phenomenon was found for the internalized NPs (Fig. S7, ESI). In fact, the membrane rings on particles were observed in some other research,24 especially when ample particles were given. However, the relationship between this intermediate phenomenon and the particle burden capacity has not been clarified. In this work, the red actin rings on the particles were shown to be derived from membrane wrapping during particle entry. The continuous engulfment of particles resulted in an increased membrane loss of the cell surface and thus an urgent need for a vast membrane supplement. Given there was a limited period for sufficient membrane recycling, extensive actin expansion or remolding provided an alternative way for adapting to an increased burden of particles. Once the membrane loss was too high to be compensated, the internalization was halted and showed the utmost level of particle burden.

For comprehensive understanding, we calculated the membrane loss index (S*, the ratio of total particle surface to cell membrane surface) and the volume magnification (V*, the ratio of total particle volume to cell volume) at the typical burden levels of low (L), middle (M), and high (H) (Fig. 3C). S* and V* were divided by a 6 μm radius spherical cell surface area and volume, respectively. As the membrane cover from the cell surface was indispensable for the completion of particle internalization, S* reflected the membrane loss at a specific V*. It was worth noting that the calculated S* for NPs (400 nm) was 19-fold above that for MPs (8 μm) at the same volume burden (V*). Although the total volume of the internalized NPs was only 0.88-fold of the cell volume, the theoretical membrane loss for NP entry was 26-fold (S* = 26) of the cell surface area. At the high burden level (V* = 2.64), S* was supposed to be 79.4 for NPs while only 4.0 for MPs. In this sense, the cell membrane failed to stretch or wrap all the nanoparticles, showing a much inferior level of NP burden. The completion of internalization was reported as unsuccessful when V* > 1 (particle volume was larger than cell volume), and researchers ascribe this event to the particle size.25 Herein, we propose a new mechanism of incomplete internalization referring to the membrane loss (S*) for particle entry. Meanwhile, we uncover the relationship between the particle burden level and acceptable loss of cell membrane. The quantification of membrane loss can also provide a theoretical basis for optimum dosage of particle carrier while cushioning the adverse effect.

To provide further verification, we examined the integrity and viability of cells after accommodating different particles. Lactic dehydrogenase (LDH) assay suggested that the leakage of the cellular enzyme was intimately related to the particle size and particle burden. Cells at a higher particle burden were inclined to leak larger amounts of LDH into the extracellular space (Fig. S8, ESI), indicating the compromised cell membrane integrity. However, exposure with MPs at the same burden (MP-M) resulted in significantly lower LDH leakage than that for NPs (Fig. 3E). According to the cell counting kit (CCK-8) assay, the viability of cells also decreased at middle NP incubation (Fig. 3D). In contrast, more than 95% of cells were still alive at high MP burden. The results not only supported the view of less membrane loss for MP entry, but also suggested a friendly coexistence between cells and MPs even at a high burden. In contrast, the rapid augment of NP dosage may not be ideal for biomedical use or daily exposure, owing to compromised cell function at higher NP burden.

Apart from the deformability of the cellular membrane, the mechanical property of the cell nucleus was also altered after particle internalization. Diverse nuclei morphologies rather than the regular oval shape were observed (Fig. 2) in CLSM and TEM imaging. With the increase of MP burden from low to high, irregular shapes including a polygon, bow tie, spindle, and even an embryo shape appeared. These shape alterations were possibly attributed to the passive adjustment of the nucleus, which utilized the flexibility of the nucleus to fill the accommodation space. In comparison, the morphology change was not evident even upon the middle burden of NPs, as the NPs with a small size were nimble enough to disperse and be accommodated instead of exerting pressure on the intracellular organelles. The mechanical stability of the cell nucleus is considered critical to many biological processes including DNA alignment, gene-level responses and cell migration dense tissue environments.26–29 However, most of the reported environments for nucleus deformation were constructed using micro/nano-pillars or other lithography methods, which were exterior and immovable. Instead, we provided a different avenue, which was named as the “particle intake method” (Fig. S9, ESI), to provide compensatory analysis inside the living cells. Despite being the stiffest organelle within the cell body,23 the nucleus was demonstrated to be capable of deformation once it encountered a higher burden of MPs. The deformation was also a size-associated event, as NPs were much weaker than MPs. In addition, membrane deformation is another important cell adjustment for accommodating MPs under a limited cell space. Furthermore, only a weak DNA degradation fragment appeared during DNA electrophoresis for cells at the MP-H burden level (Fig. S10, ESI), which confirmed the mild effect of physical burden on DNA integrity. Interestingly, MPs were in direct contact with the nucleus and exerted inward pressure, suggesting a potent avenue for gene/drug delivery to the nuclei targets. These findings highlight the advantages of the “intake method” for studying nuclear deformation. Such an interior manner provides a physical environment along with spontaneous particle arrangement rather than well-defined geometry, which reflects a natural dynamic niche for macrophage response.

The above results uncovered the uptake event of cell-sized particles and the effect of particle burden on the internalization completeness and subcellular mechanic deformation. Upon stimulation of exogenous substances, macrophages were prone to secrete numerous signalling molecules and trigger corresponding biological responses.30,31 To compare the cytokine secretion levels induced by particle burden, a series of cytokines including interleukin-2 (IL-12), tumor necrosis factor (TNF-α), interferon (IFN)-γ, monocyte chemotactic protein (MCP)-1, IL-6, and IL-10 were evaluated using the Cytometric Bead Array (CBA) on FCM (Fig. 4 and Fig. S11, ESI). In comparison with the blank control, the total fluorescence phycoerythrin (PE) signals of the cytokine profile were significantly enhanced in MP and NP groups, indicating that particles at both sizes could stimulate the cellular secretion of immune activation cytokines. However, exposure to MPs induced a lower level of cytokines (e.g. IL-6, MCP-1, and TNF-α) than that of NPs. This event was more evident when the particle burden was elevated to the middle level. It has been mentioned that the internalized MPs present a much smaller surface area than that of NPs at the same volume burden. In this sense, the interaction area between the MPs and the intracellular organelles was much smaller and led to weaker cytokine secretion. As TNF-α, MCP-1, and IL-6 are chemotactic factors for inflammation and cell recruitment, pleiotropic effects might be related.32,33 On the one hand, a self-activated response was mild for MP based adjuvant upon stimulation at a lower cytokine level. On the other hand, the MP carrier might be less active than that for the NPs in inducing a non-specific immune response, showing better biocompatibility.

image file: c7tb01673e-f4.tif
Fig. 4 Cytokine production of macrophages after exposure to MPs or NPs at different burdens.

Cell migration is a crucial process in the development and maintenance of multicellular organisms, and is involved in embryonic development, wound healing and immune responses.34,35 To examine the effect of physical burden on cell movement, the tailored trajectory of cells was recorded using spinning disk confocal microscopy. Owing to the bright autofluorescence of MPs/NPs, the particle loading cells could be easily distinguished. Representative macrophage trajectories (Fig. 5) showed that the cells moved a greater distance upon stimulation of MPs or NPs in comparison with the blank control. It was fascinating that the range of cell motion was promoted even when the cells bore a huge volume of exogenous particles. Although the MP group moved less actively in comparison with the NP group, the cell migration of the MP loaded group (MP-M) was around half the 20 μm spherical area, which was still above that of the control. This result is in line with a study regarding the enhanced migration of microparticle-stimulated cells.36 Furthermore, it is highly consistent with the finding in the aforementioned cytokine production, that MPs induce less chemotactic cytokines (e.g. MCP-1 and IL-6) than NPs. This result also recalls the attribute of macrophages that they can efficiently engulf senescent cells or cancer cells (size around 8 μm)37,38 before they generate danger signals to the body. In the case of targeted delivery, the flexible and quick movement of macrophages at high MP burden may largely prevent the toxicity of nonspecific particle distribution in tissues.

image file: c7tb01673e-f5.tif
Fig. 5 Spinning disk confocal analysis showing the tailored trajectory of cells upon stimulation of MPs or NPs (P* < 0.05, P** < 0.01).


In summary, we systematically investigate the macrophage response to cell-sized particles at different burden levels. The intake method together with the uniform autofluorescent particles ensure the achievement of distinguishable imaging and reliable quantification on aspects of cell responses to the physical signal. In detail, we uncover a formidable cell uptake capacity for MPs and a converse size-associated burden event where MPs outperformed NPs. The burden level for particles is highly related to the extent of membrane loss that is accompanied with particle entry. At the same particle burden, MPs induce mild responses on membrane integrity and cell viability (less LDH release) compared to NPs. Significant mechanic deformation of the cell membrane and nucleus synergistically endow the cell-sized MPs with the flexibility to be accommodated inside the macrophages. In addition, the MP loaded cells move slower than the NPs but quicker than the control, which is ascribed to the expression level of attractant cytokines. The present work highlights the biophysical signal obtained from the particle burden, expanding the applicability in basic and clinical research via the comprehensive design of particle size, dosage, and administration route. On the other hand, the insatiable uptake of MPs by macrophages may help to clear hazardous particles (PM2.5 to 10) or cells and thus reduce health risks.


Reagents and materials

Shirasu porous glass (SPG) membrane was obtained from SPG Technology. Chitosan (Mw = 200[thin space (1/6-em)]000) was bought from Jin Ke Co. Ltd. PO-500 (hexaglycerin penta ester) was purchased from Sakamoto Yakuhin Kogyo Co., Ltd (Japan). Paraffin and petroleum ether (boiling range 60–90 °C) were supplied by Sinopharm Chemical Reagent Co., Ltd. Acetic acid was provided by the Beijing Chemical Plant. All the other reagents were of analytical grade. Dulbecco's modified Eagle's medium (DMEM, Gibco) with 10% fetal bovine serum (FBS, Hyclone) was used. Rhodamine–phalloidin, 4,6-diamidino-2-phenylindole (DAPI), the CCK8 kit, and the LDH Assay kit were from Thermo Fisher Scientific. The Cytometric Bead Array (CBA) mouse inflammation kit was bought from BD Biosciences. All the reagents were of analytical grade.

Preparation and characterization of micro- and nano-particles

Chitosan micro- and nano-particles were prepared using an emulsification technique, and different diameters of particles (400 nm and 8 μm) were obtained by choosing specific pore size membranes. 2 wt% chitosan was dissolved in 1 wt% aqueous acetic acid as the water phase. A mixture of 3[thin space (1/6-em)]:[thin space (1/6-em)]1 (NPs) and 7[thin space (1/6-em)]:[thin space (1/6-em)]5 (MPs) (v/v) liquid paraffin/petroleum ether containing 4 wt% PO-500 emulsifier was used as the oil phase. For the microparticles, the water phase was directly permeated through the uniform pores of the SPG membrane into the oil phase under an appropriate pressure of nitrogen gas to form a W/O emulsion. For the nanoparticles, a process of low-speed homogenization was carried out, and then the nanodroplets were extruded from the uniform pores of the SPG membrane (1.4 μm). Under an appropriate pressure of nitrogen gas, this process was repeated 3 times. After that, glutaraldehyde was slowly added to either the micro- or nano-emulsion to solidify the chitosan droplets. Then, the particles were collected and washed with petroleum ether, acetone, ethanol, and deionized water under a specific centrifugation force. The morphology and size distribution of particles was determined on a JEM-6700F (JEOL) scanning electron microscope (SEM) and a Zetasizer ZS (Malvern) analyzer, respectively. As the particles were auto-fluorescent, the images of the particles were also directly observed using an SP5 laser confocal microscope (Leica) without adding any staining probe.

Cell culture

C57BL/6 mice were obtained from the Department of Laboratory Animal Science, Peking University Health Science Center (China). All animals were caged according to principles established for the care and use of laboratory animals. Mouse peritoneal macrophages (PMØ) were acquired from the C57BL/6 mice. The mice were stimulated referring to a typical protocol, and the collected cells were cultivated in DMEM medium with 10% (v/v) FBS at 37 °C, 5% CO2.

Cellular uptake study

Particles with different concentrations were added to the cell culture medium. After a desired incubation period, the cells were extensively washed with phosphate buffer solution (PBS). Samples were tested on a flow cytometer (CyAn ADP 9) through the FL-1 channel, and fluorescence intensity (FI) data were acquired from 20[thin space (1/6-em)]000 cells per sample.

The distribution of MP number (n) per cell at three burden levels was calculated from the FI data using the following equation. At different MP concentrations, the distribution of MP number (n) in the cells is proportional to the FI.

image file: c7tb01673e-t1.tif
FIcell means the detected fluorescence intensity per cell, which also reflected the total FI of all the particles in one cell; FIp means the fluorescence intensity per particle, which is 107.18 for 8 μm MP.

S* and V* were calculated using the following equations. V* is the magnification of the cell volume after particle internalization, which reflects the present particle burden. As the membrane cover from the cell surface was indispensable for the completion of particle internalization, S* reflected the membrane loss at specific V*.

image file: c7tb01673e-t2.tif

image file: c7tb01673e-t3.tif
VTp means the total volume of particles that are inside one cell, which can be calculated from the given FI (Y) using the linear equation YFI = 0.37 × X(V) + 9.41.

V cell means the volume of a natural cell (at an average diameter of 12 μm), which is 904.32 μm3.

S Tp means the total surface area of particles that are inside one cell.

S cell means the surface area of membrane for one cell (at an average diameter of 12 μm).

R cell means the average radius of a natural cell (6 μm).

R p means the average radius of a round particle (4 μm for MPs and 200 nm for NPs).

SEM, CLSM and TEM imaging of cells at different burdens of particles

PMØ were allowed to adhere for 24 h on slides which were inside a 6-well plate. Afterwards, micro or nanoparticles at low, middle, or high concentrations were added for a 2 h incubation period and rinsed with PBS. For the SEM images, cells were fixed in 2.5% glutaraldehyde (pH 7.4) at room temperature for 1 h, then serially dehydrated with acetonitrile. In order to maintain the original appearance, a K850 Critical Point Drier (EMITECH, American) was used to dry the samples. Subsequently, the samples were coated with platinum by ion sputtering (JFC-1600, JEOL), and then SEM was employed to observe the internalization process. For CLSM imaging, cells were fixed in 3.7% paraformaldehyde for 30 min and the cell membrane and nucleus were stained with rhodamine–phalloidin and DAPI, respectively. Images of the corresponding fluorescent cells were obtained using a TCS SP5 CLSM (Leica). For TEM (transmission electron microscopy) imaging, cells were fixed in 2.5% glutaraldehyde (pH 7.4) for 1 h at room temperature, and then were washed with PBS three times and soaked in PBS. Afterwards, serial sections were cut by a Reichert Ultracut microtome (Leica), and images were taken using a JEM-1400 (JEOL) TEM.

CCK8 and LDH release assay

The total release of cytoplasmic lactate dehydrogenase into the medium is a consequence of cellular integrity damage. Cells were seeded in a 96-well plate and treated with microparticles at different concentrations. The evaluation is based on a coupled enzymatic conversion from a tetrazolium salt into a formazan product, and the absorbance was measured at 450 nm by an Infinite M200 microplate spectrophotometer (Tecan).

Cytokine profile of cells upon exposure with particles

In order to address the effect of particle burden on the cytokine profile of macrophages, a CBA mouse inflammation kit was employed. After exposure to particles at different concentrations for 24 h, the cell supernatants were collected and the amounts of IL-6, IL-10, MCP-1, IFN, TNF and IL-12 were determined. Experiments were carried out following the manufacturer's instruction manual. In brief, 50 μL of sample was incubated for 1 h with the appropriate detection beads and analyzed by CyAn ADP9 color flow cytometry (FACS).

Trajectory analysis of cells

Cells were allowed to adhere to a petri dish for 24 h, and 8 μm microparticles and 350 nm nanoparticles at different concentrations were subsequently added. The interaction videos between the particles and cells were recorded by spinning disk confocal microscopy (SDCM, Leica) under a bright field. The trajectories of five different individual cells (n = 5) from numerous observed cells were provided. The trajectory images were quantitatively assessed using a 30 μm reference sphere and were aligned with the Ultraview analysis system.

Statistical analysis

All the experiments were represented as a mean ± standard deviation (SD). Statistical evaluations of the data were performed by the Student's t test for two groups, and the one-way ANOVA test for multiple groups, where P* < 0.05, P** < 0.01, P*** < 0.001 and P**** < 0.0001.

Conflicts of interest

There are no conflicts to declare.


This work was supported by the National Natural Science Foundation of China (51302265 and 21622608), the Major Project of the Ministry of Science and Technology of China (2014ZX09102045), Beijing Talents Fund (2015000021223ZK20), and the Youth Innovation Promotion Association, CAS (2013033).

Notes and references

  1. N. G. Sosale, K. R. Spinier, C. Alvey and D. E. Discher, Curr. Opin. Immunol., 2015, 35, 107–112 CrossRef CAS PubMed.
  2. M. A. Kafi, W. A. El-Said, T. H. Kim and J. W. Choi, Biomaterials, 2012, 33, 731–739 CrossRef PubMed.
  3. H. Yue and G. Ma, Vaccine, 2015, 33, 5927–5936 CrossRef CAS PubMed.
  4. S. Mitragotri and J. Lahann, Nat. Mater., 2009, 8, 15–23 CrossRef CAS PubMed.
  5. R. R. Shah, D. T. O'Hagan, M. M. Amiji and L. A. Brito, Nanomedicine, 2014, 9, 2671–2681 CrossRef CAS PubMed.
  6. M. O. Oyewumi, A. Kumar and Z. R. Cui, Expert Rev. Vaccines, 2010, 9, 1095–1107 CrossRef CAS PubMed.
  7. Y. F. Ma, Y. Zhuang, X. F. Xie, C. Wang, F. Wang, D. M. Zhou, J. Q. Zeng and L. T. Cai, Nanoscale, 2011, 3, 2307–2314 RSC.
  8. D. N. Nguyen, J. J. Green, J. M. Chan, R. Longer and D. G. Anderson, Adv. Mater., 2009, 21, 847–867 CrossRef CAS PubMed.
  9. Z. G. Yue, Z. X. You, Q. Z. Yang, P. P. Lv, H. Yue, B. Wang, D. Z. Ni, Z. G. Su, W. Wei and G. H. Ma, J. Mater. Chem. B, 2013, 1, 3239–3247 RSC.
  10. Y. Liu, Y. Yin, L. Y. Wang, W. F. Zhang, X. M. Chen, X. X. Yang, J. J. Xu and G. H. Ma, J. Mater. Chem. B, 2013, 1, 3888–3896 RSC.
  11. P. P. Lv, Y. F. Ma, R. Yu, H. Yue, D. Z. Ni, W. Wei and G. H. Ma, Mol. Pharmaceutics, 2012, 9, 1736–1747 CrossRef CAS PubMed.
  12. M. Kanapathipillai, A. Brock and D. E. Ingber, Adv. Drug Delivery Rev., 2014, 79–80, 107–118 CrossRef CAS PubMed.
  13. J. H. van den Berg, K. Oosterhuis, W. E. Hennink, G. Storm, L. J. van der Aa, J. F. Engbersen, J. B. Haanen, J. H. Beijnen, T. N. Schumacher and B. Nuijen, J. Controlled Release, 2010, 141, 234–240 CrossRef CAS PubMed.
  14. W. Miao, G. Shim, S. Lee, S. Lee, Y. S. Choe and Y. K. Oh, Biomaterials, 2013, 34, 3402–3410 CrossRef CAS PubMed.
  15. J. J. Moon, B. Huang and D. J. Irvine, Adv. Mater., 2012, 24, 3724–3746 CrossRef CAS PubMed.
  16. Z. Cheng, A. Al Zaki, J. Z. Hui, V. R. Muzykantov and A. Tsourkas, Science, 2012, 338, 903–910 CrossRef CAS PubMed.
  17. E. van Riet, A. Ainai, T. Suzuki, G. Kersten and H. Hasegawa, Adv. Drug Delivery Rev., 2014, 74C, 28–34 CrossRef PubMed.
  18. S. C. Balmert and S. R. Little, Adv. Mater., 2012, 24, 3757–3778 CrossRef CAS PubMed.
  19. J. A. Yang, H. T. Phan, S. Vaidya and C. J. Murphy, Nano Lett., 2013, 13, 2295–2302 CrossRef CAS PubMed.
  20. H. Yue, W. Wei, Z. G. Yue, P. P. Lv, L. Y. Wang, G. H. Ma and Z. G. Su, Eur. J. Pharm. Sci., 2010, 41, 650–657 CrossRef CAS PubMed.
  21. N. Darville, M. van Heerden, A. Vynckier, M. De Meulder, P. Sterkens, P. Annaert and G. Van den Mooter, J. Pharm. Sci., 2014, 103, 2072–2087 CrossRef CAS PubMed.
  22. M. Nikkhah, F. Edalat, S. Manoucheri and A. Khademhosseini, Biomaterials, 2012, 33, 5230–5246 CrossRef CAS PubMed.
  23. C. T. McKee, V. K. Raghunathan, P. F. Nealey, P. Russell and C. J. Murphy, Biophys. J., 2011, 101, 2139–2146 CrossRef CAS PubMed.
  24. W. Wei, L. Y. Wang, L. Yuan, Q. Wei, X. D. Yang, Z. G. Su and G. H. Ma, Adv. Funct. Mater., 2007, 17, 3153–3158 CrossRef CAS.
  25. J. A. Champion and S. Mitragotri, Proc. Natl. Acad. Sci. U. S. A., 2006, 103, 4930–4934 CrossRef CAS PubMed.
  26. P. Formentin, U. Catalan, M. Alba, S. Fernandez-Castillejo, R. Sola, J. Pallares and L. F. Marsal, New Biotechnol., 2016, 33, 781–789 CrossRef CAS PubMed.
  27. Z. Pan, C. Yan, R. Peng, Y. Zhao, Y. He and J. Ding, Biomaterials, 2012, 33, 1730–1735 CrossRef CAS PubMed.
  28. L. Hanson, W. Zhao, H. Y. Lou, Z. C. Lin, S. W. Lee, P. Chowdary, Y. Cui and B. Cui, Nat. Nanotechnol., 2015, 10, 554–562 CrossRef CAS PubMed.
  29. K. Wolf, M. te Lindert, M. Krause, S. Alexander, J. te Riet, A. L. Willis, R. M. Hoffman, C. G. Figdor, S. J. Weiss and P. Friedl, J. Cell Biol., 2013, 201, 1069–1084 CAS.
  30. R. Sridharan, A. R. Cameron, D. J. Kelly, C. J. Kearney and F. J. O'Brien, Mater. Today, 2015, 18, 313–325 CrossRef CAS.
  31. D. Nagorsen, S. Deola, K. Smith, E. Wang, V. Monsurro, P. Zanovello, F. M. Marincola and M. C. Panelli, Genome Biol., 2005, 6, R15 CrossRef PubMed.
  32. R. L. Coffman, A. Sher and R. A. Seder, Immunity, 2010, 33, 492–503 CrossRef CAS PubMed.
  33. H. Yue, W. Wei, Z. Yue, B. Wang, N. Luo, Y. Gao, D. Ma, G. Ma and Z. Su, Biomaterials, 2012, 33, 4013–4021 CrossRef CAS PubMed.
  34. M. Mak, F. Spill, R. D. Kamm and M. H. Zaman, J. Biomech. Eng., 2016, 138, 021004 CrossRef PubMed.
  35. C. Y. Tay, P. Cai, M. I. Setyawati, W. Fang, L. P. Tan, C. H. Hong, X. Chen and D. T. Leong, Nano Lett., 2014, 14, 83–88 CrossRef CAS PubMed.
  36. I. M. Meraz, B. Melendez, J. Gu, S. T. Wong, X. Liu, H. A. Andersson and R. E. Serda, Mol. Pharmaceutics, 2012, 9, 2049–2062 CrossRef CAS PubMed.
  37. A. Hochreiter-Hufford and K. S. Ravichandran, Cold Spring Harbor Perspect. Biol., 2013, 5, a008748 CrossRef PubMed.
  38. D. V. Krysko, G. Denecker, N. Festjens, S. Gabriels, E. Parthoens, K. D'Herde and P. Vandenabeele, Cell Death Differ., 2006, 13, 2011–2022 CrossRef CAS PubMed.


Electronic supplementary information (ESI) available. See DOI: 10.1039/c7tb01673e
These authors contributed equally to the work.

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