Controlling Surface-Induced Platelet Activation by Agarose and Gelatin-Based Hydrogel Films.

Platelet-surface interaction is of paramount importance in biomedical applications as well as in vitro studies. However, controlling platelet-surface activation is challenging and still requires more effort as they activate immediately when contacting with any nonphysiological surface. As hydrogels are highly biocompatible, in this study, we developed agarose and gelatin-based hydrogel films to inhibit platelet-surface adhesion. We found promising agarose films that exhibit higher surface wettability, better controlled-swelling properties, and greater stiffness compared to gelatin, resulting in a strong reduction of platelet adhesion. Mechanical properties and surface wettability of the hydrogel films were varied by adding magnetite (Fe3O4) nanoparticles. While all of the films prevented platelet spreading, films formed by agarose and its nanocomposite repelled platelets and inhibited platelet adhesion and activation stronger than those of gelatin. Our results showed that platelet-surface activation is modulated by controlling the properties of the films underneath platelets and that the bioinert agarose can be potentially translated to the development of platelet storage and other medical applications.

Introduction

Platelets are anucleated and discoid-shaped cells with an average diameter of 1–3 μm produced from the megakaryocyte cells in the bone marrow.[1] They play a vital role in hemostasis by clotting the blood at the sites of ruptured endothelium.[2] The blood of a healthy human contains an average of 150 000–400 000 platelets/μL.[3]

In blood circulation, the red blood cells, blood shear rate, immune system, and coagulation system have a great influence on the activation and adhesion of platelets. In vitro, platelets tend to activate immediately after a short contact with artificial surfaces, which is a drawback for many applications such as platelet storage and platelet-drug studies.[4] Activated platelets expose glycoprotein IIb/IIIa, which initiates the binding of fibrinogen and facilitates platelet aggregation,[5,6] and they also release prothrombotic substances from their granules.[7] The releasing proteins, on the one hand, cross-link and activate the surrounding platelets and, on the other hand, enhances thrombin generation together with plasma clotting factors. In serious cases, this leads to the formation of a hemostatic plug at the site of endothelial damage, which eventually results in blood vessel closure.[8,9] In implantation, blood proteins adsorb on the surface of an implant, thus allowing the adhesion of platelets, which can lead to thrombus formation or even escalate to stroke or cause extremity ischemia.[10,11] Hence, platelets play a key role in deciding the fate of an implant in the early stages post implantation.[12]

Biomaterials and biopolymers, in particular, are widely used in the development of various cardiovascular valves,[13] artificial blood vessels,[14] dialyzers,[15] and implants.[16] Especially, the aspect of coagulation, which involves protein adsorption followed by platelet adhesion, is of great relevance when dealing with applications involving blood purification like hemodialysis, plasmapheresis, or blood oxygenation.[17] Thus, it is important to control the biofouling of the surfaces by plasma proteins, which can ultimately lead to thrombosis.[18] Surface-related complications from the implant material can be minimized by developing effective surface coatings.[19] However, the development of an antithrombotic surface for implant continues to remain a challenge. Minimal platelet-surface activation has a major impact not only in implantation but also in platelet storage. Thus, the development of antithrombogenic surfaces with the aim to reduce the administration of anticoagulants is of utmost importance. However, the field of developing an antithrombotic surface, especially for platelets, requires further investigation. Additionally, the unavailability of a hemocompatible bulk biomaterial has encouraged the development of new surface coatings and surface engineering to tackle the problem of thrombosis.[19]

For platelet storage, the short lifetime (up to 5 days) complicates the management of the continuous demand for transfusion of platelets.[12] Patients with thrombocytopenia, with platelet defects, or suffering from bleeding while undergoing chemotherapy are in severe need of platelet transfusion.[20] Platelets are preserved at 22 °C to sustain their functionality,[12] and are regarded as fresh and young when they are stored for less than 3 days.[21] Stored platelets themselves, along with the storage medium, undergo changes that can cause platelet activation and dysfunction.[22] The short lifespan of the platelets is attributed to the activation of platelets in some cases and due to bacterial contamination in others.[23]

We have shown previously that the fabrication of nanotopography surface together with surface modification with collagen-G or laminin strongly reduced platelet-surface activation.[4,6] However, the fabrication of nanostructures requires advanced technologies,[6,24] while the collagen-G coating only allows inhibiting platelet-surface activation for a short time (∼15 min)[6] and, therefore, these methods are only suitable for research to understand the fundamental aspects of platelet interactions. These drawbacks limit many applications in medicine. It is extremely important to develop a simple and bioinert surface for the inhibition of platelet-surface activation.

Hydrogels are highly hydrated polymers, consisting of a polar polymer backbone and an enormous content of bound water.[25] Thus, hydrogels are promising candidates for any biomaterial and have the potential to mimic the native biological tissue microenvironment.[26] Hydrogels display antifouling activity, especially in the presence of hydrophilic hydroxyl groups and, therefore, are good candidates for developing blood-compatible materials.[27] It has been reported that the platelet density and degree of activation on the surface of synthetic hydrogel films can be controlled; thus, setting them one step further at being bioinert for platelet applications.[28]

Hydrogels containing agarose and gelatin find a wide range of applications in the field of tissue engineering and regenerative medicine.[29,30] Their unique properties, i.e., the capability to hold a high amount of water molecules (up to 40 fold of the dry weight) while maintaining the structural stability, give hydrogels an edge among other biomaterials.[31] The surface of a biomaterial plays a crucial role in determining the hemocompatibility of the material.[32] The steric repulsion by the hydrated chains in hydrogels contributes to their bioinertness.[28] Agarose, a polymer of natural origin, contains hydroxyl groups that contribute to the overall antifouling properties of the hydrogel.[33−36] Similarly, gelatin is also a biocompatible biopolymer with hydration properties and exhibits antimicrobial properties.[37] A reduction of pig platelet adhesion on agarose has been observed.[38] However, the response of human platelets on hydrogels, especially formed by agarose, has been insufficiently investigated. In this study, we filled up this gap by tracking the behavior of platelets on gels formed by agarose and also gelatin.

A novel approach to strengthen the hydrogel networks by incorporating nanoparticles (NPs) has been recently reported.[39−41] Mixing of the NPs to the existing polymeric network or blending them with polymers to cross-link the polymer chains are some of the developed procedures to fabricate nanocomposite hybrids.[39] Such hybrids are termed nanocomposite hydrogels. Nanocomposite hydrogels enhance the existing physical, chemical, and biological properties of the hydrogels.[31] NPs coated with carboxymethyl dextran (CMD) consists of a dextran backbone substituted with carboxymethyl groups that imparts a polyanionic character to the nanoparticle. Furthermore, the soft polymer coat that surrounds the NPs makes these systems less likely to aggregate due to the steric repulsions between particles.[42] In our study, we investigated the effect of Fe3O4 NPs on characteristics of agarose and gelatin gels and tracked if the nanocomposites reduce platelet-surface activation. Due to their superparamagnetic properties, Fe3O4 NPs are used for multiple biomedical applications ranging from magnetic resonance imaging (MRI), magnetic particle imaging (MPI) to drug delivery with certain coatings and, hence, are regarded as biocompatible.[43,44] Importantly, they also have antibacterial properties[45,46] that may become promising nanocomposites against bacterial contamination in platelet storage. We hypothesize that the nanocomposite hydrogels with defined characteristics can modulate the response of platelets toward a nonphysiological surface.

In this study, we fabricated agarose as well as gelatin hydrogels and extended to nanocomposite hydrogels by incorporating synthesized CMD-coated Fe3O4 NPs in the respective hydrogels. The influence of the various physicochemical properties of the gels on the platelet adhesion was examined. Multiple technologies such as dynamic light scattering (DLS), contact angle, nanoindentation, force spectroscopy, confocal laser scanning microscopy (CLSM), and scanning electron microscopy (SEM) were applied to characterize the fabricated gels and to study the response of platelets on them. We found that factors like stiffness, adhesion force, and wettability play a crucial role in developing a bioinert material. Agarose and its nanocomposite induced the strongest inhibition of platelet-surface adhesion and activation, followed by those of gelatin, and the weakest inhibition was observed on bare glass. Our results showed that agarose hydrogels have potential applications for material-based implants, improvement of platelet and blood storage bags, and biotechnology and pharmaceutical trials.

Results

Experimental Design

We first tracked the impact of hydrogels and nanocomposite films on the reduction of platelet-surface activation by fabricating films using bioinert materials, including gelatin and agarose, together with their nanocomposite gels (Figure ). The investigation of platelet response on gels formed by different concentrations of the polymer showed that gelatin of 10% and agarose of 1% provided the highest stability. Furthermore, the hydrogels were fabricated in the customized silicone molds and needed to be removed from the molds before characterization. During this process, the thin film was easy to be damaged. To identify the optimal thickness, we fabricated gels of different thicknesses and found that the film of 2 mm thickness is the most stable. Therefore, gelatin of 10%, agarose of 1%, and film thickness of 2 mm were selected for further investigation in this study. To form gels, gelatin and agarose were first melted at 60 and 95 °C, respectively, before leaving them to physically cross-link at 35 and 40 °C, respectively (Figure B–E). A cooling process allows the molecules to form stable networks of triple and double helix-coil transition arrangements of their polymer chains.[47,48]

Figure 1

Schematic representation of the polymer networks without and with NPs in the hydrogel. (A) Bare glass surface contains multiple hydroxyl groups and is used as a control. (B) Gelatin hydrogels form a triple helix-coil through hydrogen bonds. (C) Addition of NPs (green) further cross-links the negatively charged-carboxy groups of CMD coated on the particle surface and the amino groups on gelatin chains. (D) Agarose hydrogels possess the double helix-coil structure of their fibers. (E) Addition of NPs interferes during the gelation phase, thereby causing a poor matrix formation compared to the native agarose hydrogels.

To form the nanocomposite films, different concentrations of NPs in the films were tested to determine the threshold beyond which the particles caused aggregation of platelets on the surface. We observed that platelets aggregated from 3.2 mM concentration, while no aggregation was observed at ≤2 mM concentration (Figure S1). To gain the maximal effect of nanoparticles, a high concentration of particles should be added to the gels. Thus, 2 mM was identified as the optimum particle for the formation of nanocomposites (Figure S1). The electrostatic interaction between the carboxylic groups, present in CMD molecules coated on the NPs, and the amine groups, present in the gelatin, cross-link and stabilize the nanocomposite gels.

Characterization of Fabricated Hydrogel Films

Surface Wettability

Next, we determined the surface wettability of the fabricated films via static water contact angle measurements using the captive bubble method. For this, glass slides coated with the layer of hydrogel and the hydrogel nanocomposites were inverted and submerged in distilled water (Figure A). An air bubble was trapped, and the contact angle was measured using the ellipse fitting method (Figure B). The lower contact angles denote that the surface has higher hydrophilic properties. The contact angle was highest for the bare glass surface (51.1 ± 4.2°), followed by gelatin (22.6 ± 4.4°), and lowest for agarose (10.6 ± 6.8°) (Figure C). However, nanocomposites drive surface wettability differently. The contact angle for gelatin nanocomposites (46.1 ± 12.5°) was higher than that of the gelatin alone (22.6 ± 4.4°), whereas it showed a reverse trend for agarose nanocomposites (4.8 ± 6.6°) vs agarose (10.6 ± 6.8°). However, agarose gels showed much lower contact angles than gelatin gels, indicating higher hydrophilic properties of agarose gels.

Figure 2

Contact angle measurements using the captive bubble technique. (A) Schematic experimental setup for the measurement of gels on glass samples. (B) Optical image of the hydrogel sample and the air bubble (side view). (C) Graph shows the recorded values of the contact angle on different samples along with the standard deviation. *Statistically significant difference determined by the one-way ANOVA test (P < 0.05).

Water Retention Properties

The stability of the hydrogels and the nanocomposites was studied at room temperature (RT) by measuring their swelling and water retention properties. For this, we determined the weights of the fabricated gel (Wo), wet gel (WPBS), which was immersed in PBS, and gel stored in the dry environment (Wair) at different time points. The percentage of swelling or degradation at each time point was calculated as WPBS/Wo or Wair/Wo, respectively. The trends in degradation were almost similar irrespective of the polymer or the use of the NPs in the case of hydrogel degradation (Figure A). However, during swelling, there was a clear difference observed between the gelatin and agarose polymers (Figure B). While gelatin films showed a steeper incline in the weight gain, the increase in the weight of agarose films was nearly negligible.

Figure 3

Degradation and swelling behavior of the hydrogel and nanocomposite films. (A) No significant difference in water retention between the gelatin (green), gelatin nanocomposite (red), agarose (blue), and agarose nanocomposite (yellow) was obtained. (B) Gelatin (green) and gelatin nanocomposite (red) exhibited a continuous increase in their swelling behavior, whereas agarose (blue) and agarose nanocomposite (yellow) did not show a significant incline in swelling of the gels and reached equilibrium. n = 3 repetitions.

Gel Stiffness

We next determined the mechanical properties of the fabricated films using the AFM-based nanoindentation method with a colloidal probe of 3 μm diameter (Figure A). By applying an indentation force, the bead indents the underlying surface causing the cantilever to bend and indentation of the gel is detected in the force–distance curve. Typical approach curves showed the highest indentation depth on gelatin, followed by gelatin nanocomposites, lower indentation depth on the agarose nanocomposite, and lowest on agarose (Figure B). Fitting the indentation curve caused by the deformation with the Hertz model[49] allowed obtaining the Young’s modulus (E) of the films. Data analysis of F–D curves that were recorded at different locations on each gel from two independent times of gel preparation showed the E value of the gelatin film (Figure C) to be more than an order of magnitude lower than that of agarose (Figure D). However, NPs enhanced the E value of the gelatin film, whereas they reduced the E value of the agarose film (Figure C,D).

Figure 4

Determination of the stiffness of the fabricated films by nanoindentation. (A) Schematic illustration of a probe contacting the hydrogel and nanocomposite samples. Below: SEM image of the AFM cantilever with a gold colloidal particle attached. (B) Typical force–indentation curves on the four different films. (C) Young’s modulus was recorded on gelatin (green) and gelatin nanocomposites (red), (D) on agarose (blue), and agarose nanocomposites (yellow). *Significant difference determined by the one-way ANOVA test (P < 0.05). Note: the scale bar in the y-axis in (C) differs from (D).

Platelet Adhesion on the Fabricated Films

We seeded platelets on the four investigated film types and tracked the response of the platelets on those surfaces using confocal laser scanning microscopy (CLMS). The number of adhered platelets was highest on bare glass (Figure A), followed by gelatin (Figure B,C), and lowest on agarose surfaces (Figure D,E).

Figure 5

Confocal micrographs showing platelets stained with the anti-CD42a antibody dye on different surfaces after 2 h incubation time. (A) On the glass, a higher density of platelets and a higher degree of platelet activation as compared to (B) gelatin, (C) gelatin nanocomposites, (D) agarose nanocomposites, and (E) agarose were observed. (F) Average spreading area of the adherent platelets on the different substrates. *Statistically significant difference determined by the one-way ANOVA test (P < 0.05).

A change in platelet morphology along with the development of filopodia and lamellipodia was observed on the glass surfaces, whereas no significant activation of platelets was observed on all other surfaces (Figure B–E). The calculated spreading area of the platelets on bare glass (7.9 ± 3.0 μm2) was significantly higher than on other surfaces (Figure F). However, the size of platelets on gelatin (4.2 ± 1.7 μm2) is slightly higher than that on agarose (3.4 ± 1.6 μm2) in both cases, in the absence and presence of nanoparticles (Figure F). Adding NPs showed no change in platelet spreading on both gelatin (3.8 ± 1.7 μm2) and agarose gels (3.5 ± 1.5 μm2) (Figure F). The results indicate that both agarose and its nanocomposite gels inhibited platelet-surface adhesion and activation more than gelatin and its nanocomposite gels.

Adhesion Force between Platelets and Films

Next, we directly determined the adhesion force between single platelets and the fabricated gels using FluidFM technology (Figure A). A single platelet was picked up from the surface with a nanopipette (Figure B) and moved to the desired gels for measuring the adhesion force. A typical force–distance curve shows the adhesion events that occurred during the retraction of the platelet from the surface (Figure C).

Figure 6

(A) Schematic representation of FluidFM for measuring platelet-gel adhesion forces. (B) Simplified representation of picking platelets from a surface. (C) A typical retraction curve was recorded on the glass surface, showing rupture events that occurred while the adherent platelet disrupted from the surface. (D) Box plot representation of variations in adhesion forces at different setpoints (with contact time 0 s). (E) Box plot comparing the difference between the adhesion forces on glass surfaces for different contact times (with setpoints of 10 nN). (F) Typical representation of adhesion forces between single platelets and different surfaces presented in the form of box plots, from n = 3 independent platelet donors/conditions. *Statistically significant difference determined by the one-way ANOVA test (P < 0.01) for (E) and (P < 0.05) for (D) and (F). Independent platelet donors for (D) and (E).

To obtain comparable results, we first determined the dependency of the results on the chosen measurement parameters to find a parameter set suitable for comparative analysis. The parameters, with which the force–distance curves are recorded, can influence the final adhesion force values. Here, we also investigated the effect of measurement parameters, such as contact time and applied setpoint, on adhesion forces of platelets on glass surfaces. Higher setpoints also resulted in stronger adhesion forces. To overcome electrostatic interactions between the platelet and the glass surface, a setpoint of ≥5 nN must be applied in our experiment. Therefore, we investigated here the effect of the setpoint on the adhesion force between platelets and bare glass in the range from 5 to 20 nN, and an increase in the final adhesion force with an increasing setpoint was observed. The lowest adhesion force was observed with a 5 nN setpoint. A four times higher setpoint led to an almost doubling of the adhesion forces (Figure E). We assume that a higher applied force (=higher setpoint) forced more area of the platelets into the adhesion contact with the glass surface, leading to a larger adhesion force. At setpoints of 10 nN, the lowest variations (error bars see Figure D) and intermediate adhesion forces were observed. Thus, a 10 nN setpoint was identified as the best-applied force for our force spectroscopy measurements.

In addition to higher setpoints, longer contact times also resulted in stronger adhesion forces. The contact time, which is the time duration that the platelet stays in contact with the surface, was investigated between 0 and 7.5 s. The adhesion force increased with the increasing contact time. With 0 s contact time, a weak adhesion force of around 0.5 nN with small variation was observed, whereas it increased to about 4 nN with a large variation at 7.5 s contact time (Figure E), indicating a strong and complex response of the platelet on the glass as also seen in the CLMS image (Figure A). The variation of the adhesion force between 2.5 and 5 s contact time was low, indicating a stable range of the measured adhesion force, and we selected 5 s contact time for further experiments. The successful force curves obtained at 5 s contact time showed a yield of 91%, which is approximately equal to that at 0 s contact time. At too long contact time (7.5 s), the yield of successful force curves significantly reduced due to the strong adhesion of platelets on the glass as also observed previously.[4] A large variation of adhesion force also results from different platelet donors and platelet heterogeneity.[50]

We then applied the above conditions (10 nN setpoint, 5 s contact time) to measure the adhesion forces between platelets and gels. A clear difference in adhesion forces between glass and hydrogels was observed (Figure F). The recorded force values on the glass surface were up to 5 times higher than the ones recorded on the gels. The adhesion forces measured on gelatin were significantly higher than that on agarose, while gelatin nanocomposites showed higher adhesion forces compared to the agarose nanocomposites. However, no significant difference was induced by the nanocomposite gel as compared with their respective native hydrogels (Figure F).

Discussion

In this study, we found that agarose-based hydrogel and its nanocomposite are stable, inert, and can strongly reduce platelet-surface adhesion and activation. Gelatin nanocomposites enhance the gel stiffness as compared to the native gel but it shows lower stability than agarose gel. Though the gelatin film shows a significant reduction of platelet adhesion as compared to the bare glass, the degree of platelet adhesion and activation is slightly higher than that of the agarose film. Agarose and gelatin hydrogels as well as their nanocomposites exhibit a dissimilar effect on platelet adhesion and activation depending on multiple characteristics such as wettability, gel stiffness, water retention, and chemical function. The agarose and its nanocomposite gels are the most promising materials for the inhibition of platelet-surface activation.

The wettability of the biomaterials, known as surface hydrophilicity or hydrophobicity, is important, especially when the material is going to be in close contact with the blood. Our investigated agarose films show high hydrophilicity, indicating a surface with high resistance to unfavorable protein absorbance, which leads to a strong reduction of platelet adhesion. Consistently, it has been revealed that the degree of platelet-surface activation decreases with an increase in hydrophilicity of the surface,[51,52] whereas a hydrophobic surface facilitates absorbance of proteins and enhances platelet-surface activation.[53] Furthermore, surface chemistry is also known to contribute to surface wettability.[54] It is likely that the presence of multiple hydroxyl groups in agarose molecules makes them highly hydrophilic (Figure ). The degree of hydrophilicity is highest for agarose, followed by gelatin, and lowest for glass, indicating a possibility to tune the surface properties of glass or other hard metal surfaces by coating with these polymers. Our observation of a higher hydrophilic nature of agarose in comparison with the gelatin is consistent with the previous study.[55]

When adding NPs to the gels, the wettability of agarose did not significantly change, whereas gelatin nanocomposites showed a strong decrease in wettability (Figure C). This is due to the cross-linking of the CMD, coated on the Fe3O4 nanoparticles, with the gelatin. This reduces the number of free polar amino groups on the surface in the gelatin nanocomposite sample as compared to the case of native gelatin gel. In the case of agarose, the additional availability of −COOH groups on NPs in addition to the existing −OH groups contributes to making the gel surface slightly more hydrophilic. Consistently, our results showed a decrease in the contact angle on agarose after the addition of NPs, but it increased in the case of gelatin nanocomposites. The distinct surface characteristics such as hydrophilicity, among the investigated surfaces, probably control the degree of platelet adhesion and activation. We observed the highest contact angle (=lowest hydrophilicity) on bare glass, followed by the gelatin composites and gelatin, and the lowest on agarose composites and agarose. These lead to the highest density of platelet adhesion on the glass surface, followed by gelatin composite and gelatin, and lowest on agarose composite and agarose. Recently, several types of nanoparticles showed a potential impact on inhibiting bacterial growth.[56−58] As current platelet storage meets a serious drawback due to bacterial contamination, our nanocomposite gels may be a powerful tool to not only stabilize the gels but also provide antibacterial properties. However, this hypothesis requires further investigation.

Water retention and swelling of hydrogels are important to understand the stability of the hydrogels. Our results indicated that the evaporation of water molecules was low and irrespective of the composition of a hydrogel. The swelling studies for hydrogels performed at RT are of specific importance for the platelet storage since the storage itself is also carried out at room temperature.[59] Furthermore, it is reported that the swelling behavior in hydrogels is also associated with the cell adhesion properties of the hydrogels[60] and that the hydrogels with lower swelling degrees exhibit poor cell adhesion.[61] The more controlled-swelling behavior in the case of agarose-based gels contributes to restricting the platelet attachment to the surface. In contrast, both gelatin composite and gelatin show a linear increase in water retention over time that resulted in a slight increase in the platelet response on these surfaces.

The stiffness of the surface is also an important mediator in controlling the adhesion, activation, and spreading of the platelets on any surface.[62] We found that agarose gels showed higher stiffness than gelatin. Glycoproteins IIb–IIIa or αIIbβ3 integrins are key platelet adhesion receptors on the platelet surface and are responsible for platelet aggregation.[63] A stiffer substrate is known to induce higher resistance forces that lead to stronger platelet adhesion and outside-in signaling of αIIbβ3, which in turn generates a higher actomyosin mediated internal balancing force causing platelets to spread more.[62] However, both agarose and its nanocomposite gels strongly inhibited platelet adhesion and activation. We found a higher degree of platelet spreading on the glass, which is much stiffer (GPa range) than the soft hydrogel agarose (E = 181.2 ± 158.5 kPa) and gelatin (E = 1.72 ± 0.8 kPa). The mixture of NPs with agarose resulted in a softer film (E = 53.3 ± 68.4 kPa). Agarose undergoes gelation due to the extensive intermolecular hydrogen bonds, which eventually lead to a helix-coil structure.[64,65] Perhaps, the presence of NPs during this transition phase could have hampered the formation of the bonds and thereby weakened the polymeric network. However, in the case of gelatin nanocomposites, there was an increase in the Young’s modulus value (E = 3.84 ± 1.1 kPa) compared to the gelatin alone (E = 1.72 ± 0.8 kPa), indicating the formation of a tighter compact network. The reason for this is the electrostatic cross-linking between the amine groups of the gelatin and the carboxyl groups present within the CMD molecule coated on the NPs. The applied force for the indentation measurements was in the range of a few hundred piconewtons to mimic the traction forces generated by platelets on the surfaces. The indentation measurements are important in this study since the stiffness of the material can play a decisive role in determining the outcome of cell adhesion.[66−68]

The confocal micrographs of the platelets on the different surfaces prove yet again that the gel properties like wettability and surface stiffness play a crucial role in the development of biocompatible material. The highly hydrophilic agarose-based surfaces strongly inhibited the adhesion and activation of platelets. The trend was followed by less hydrophilic gelatin-based surfaces, which led to a weaker inhibition of platelet-surface activation. Though the number of platelets adhered onto the gelatin-based surfaces was slightly higher in comparison to agarose, it was significantly less when compared to the glass-control group. Minor activation of platelets was, however, still seen on the gelatin-based surfaces, which were completely absent in the case of agarose. It has been previously observed that the presence of certain functional groups within the polymers, such as hydroxyl, contributes to the antifouling properties of the hydrogels.[69] The overall results from the micrographs stress the importance of the antifouling properties of the surfaces since platelet adhesion is the first step that eventually leads to platelet activation cascades and aggregation.[70] Consistently, Oss et al. compared various casting techniques and found that the surface exposed to air during the casting process showed minimum adhesion of platelets.[38] The study also states that agarose, which is derived by removing the sulfate groups from the agar still shows anticoagulant activity like its source, which structurally resembles heparin.[38]

FluidFM force spectroscopy is a newly developed element integrated into the traditional AFM system to determine the adhesion force between single cells and surfaces. This technique allows picking up of single cells by applying negative pressure to a hollow cantilever, avoiding immobilization of cells on a colloid probe via chemical bonds as described in the traditional protocols, which potentially induce platelet-surface activation.[4] To date, an optimal protocol for measuring platelet adhesion using FluidFM is still missing. Here, we optimized the most important parameters in force spectroscopy measurement, including contact time and the setpoint, that directly influence the magnitude of the measured adhesion forces[71] before actually carrying out the force spectroscopy measurements between platelets and the investigated surfaces. We identified the most suitable contact time of 5 s and a setpoint of 10 nN for the platelet-surface adhesion force measurement. Other parameters such as applied pressure, z-length, and z-speed also influence the value of the measured adhesion forces in FluidFM, as described previously,[71,72] but we have kept these mentioned parameters constant throughout for platelet adhesion measurements.

FluidFM force spectroscopy results showed a clear difference between the control glass and other hydrogels or hydrogel-nanocomposite samples. The adhesion forces between single platelets and bare glass are up to 5-fold higher than their interaction force with the fabricated gels. The gelatin gel and its nanocomposite induced a higher adhesion force than those of agarose. These results are consistent with CLMS images in which only gelatin nanocomposite showed some weak platelet activation, whereas other gels did not. Thus, it is clear that the degree of platelet-surface activation strongly correlates with the adhesion forces between platelets and surfaces. We showed that several hydrogels, with their own set of properties like wettability, stiffness, and water content, can inhibit platelet activation substantially. The agarose hydrogels were able to inhibit the platelet adhesion largely. The adhesion forces between the platelets and surfaces governed the degree of platelet-surface activation.

The addition of nanoparticles did not have any significant negative effects in terms of platelet activation, indicating that these nanocomposites are safe to be used for platelet applications. Nanocomposites can be promising materials for antibacterial applications. Many efforts have been made to identify a suitable material that can inhibit platelet-surface activation for both fundamental studies and medical applications. However, to date, a stable and robust material is still missing. Previously, we found the collagen film to be able to reduce platelet-surface activation.[4] However, this film allows inhibiting platelets only up to ∼15 min,[4] or a bit longer on laminin-coated nanopatterns.[6] With agarose, a very low density of adhered platelets was observed, while we did not observe any changes in surface morphology of platelets up to 2 h of surface contact. Our present study clearly shows a potential application of the powerful agarose gel for the inhibition of platelet-surface activation as the agarose gel is easy to fabricate without the requirement of additional materials, highly stable, and extremely inert. For improvement of platelet storage bags and some specific implants, agarose could become a potential candidate. However, further investigation is still required, such as the stability of the gels in the presence of platelet concentrate and storage buffer/condition at longer times.

Methods

Synthesis of Fe3O4 Nanoparticles

Fe3O4 particles were synthesized under microfluidic conditions with a continuous flow mode. A mixture of FeCl3·6H2O and FeCl2·7H2O at a ratio of 1:2 was dissolved into deionized water, and a solution containing a coating agent was added and stirred together. A basic solution of ammonium hydroxide (NH4OH) was prepared in a different flask. The two solutions were filled in two 25 mL syringes, and a third one filled with 25 mL of deionized water was added. During the synthesis, the flow rate of each of the three syringes was set to 300 μL/min, at 70 °C. After mixing the reactants in the chamber, magnetite (Fe3O4) particles are formed. Fe3O4 suspension was transported via Teflon tubes through a stainless steel heating module, which approximately takes 20 min, followed by the collection of particles in the sample collector.

The size and the surface zeta potential of the synthesized NPs were determined by the Zetasizer (Zetasizer Nano ZS, Malvern Instruments Ltd., Worcestershire, U.K.). Dynamic light scattering (DLS) was used to determine the size of particles at 25 °C within 30 min in water using disposal cuvettes (Sigma-Aldrich, St. Louis). The measured average hydrodynamic sizes and their respective standard deviation values were analyzed. To determine the zeta potential, particles were diluted in water (pH 6.3, conductivity 0.25 mS/cm) to a concentration of 2 mM and measured in a folded capillary zeta cell at 25 °C with 10 repetitions, as previously described.[73] The particles are stable with an average size (Savg.) of approximately 255 nm and a zeta potential of around −56 mV (Figure ). For nanocomposites, the concentration of 2 mM was selected as the final concentration. Data analysis was performed using SigmaPlot (version 14.0).

Figure 7

Characteristics of synthesized NPs. Zeta Potential (blue) and the average size (Savg.) of nanoparticles (red) at 2 mM concentration measured by dynamic light scattering.

Hydrogel and Nanocomposite Fabrication

Agarose 1% (Lonza, Germany) and Gelatin 10% (Sigma-Aldrich, Germany) were added to the PBS solution (Thermo Fisher, Germany) at 95 and 60 °C under magnetic stirring, respectively. In the case of nanocomposites, particles were added to the respective solutions before they could form stable gels. The samples of the hydrogels or nanocomposites were prepared by pouring the above solution into the custom-made silicone molds. The samples with a diameter of 14 mm and a height of 2 mm were fabricated, and gels with a height less than 2 mm were found to reduce the stability of films while extracting them from the silicone molds. The surfaces exposed to air while casting were used for conducting all of the experiments. The gels were immediately used within 1 h of their fabrication.

Water Contact Angle

The water contact angle of the coatings was measured using the OCA 15+ system (DataPhysics Instruments GmbH, Filderstadt, Germany) by the captive bubble method. This method is particularly preferred while dealing with surfaces having high surface free energy and avoiding drying hydrogels during the measurement. In this method, a bubble of air is injected beneath the sample placed facing downward. The dosing volume was set to 3 μL, dosing rate as 1 μL/s, and the ellipse fitting method was chosen to compute the contact angle. The drop phase was selected as air, while the ambient phase was Milli-Q water (0.055 μS/cm). All of the surfaces were probed with five air bubbles at different positions on the coated surfaces. Calculation of contact angles was done by OCA 15+ software.

Hydrogel Swelling and Degradation

All gels were formed using the molds mentioned above and transferred to preweighed Petri dishes and weighed (Wo) using the weighing scale (Sartorius, Germany). To determine the wet weight (WPBS), the gels were immersed in 2 mL of PBS buffer, and measurements were taken after every hour, the extra PBS was removed before weighing. To determine the weight in a dry environment (Wair), the fabricated gels were dried at RT with the relative humidity level between 15 and 20%, and their weights were measured after every hour. The swelling at each time point was calculated as WPBS/Wo and degradation as Wair/Wo, as previously described.[74,75]

Nanoindentation

The mechanical properties of the fabricated hydrogels and nanocomposites were measured using the nanoindentation technique of atomic force microscopy. A gold bead is attached at the end of a cantilever (CP-CONT-AU-A, Nanoandmore GmbH, Germany) with a nominal bead diameter of 1.5–3 μm and nominal spring constant of 0.02–0.77 N/m. This bead was brought into contact with the gels to generate force vs displacement (F–D) curves. The F–D curves were obtained over the surface from a 10 × 10 μm2 area by subdividing the area into equal-sized 8 × 8 pixels for acquiring 64 F–D curves at tip velocity of 3 μm/s. After converting the force curve to the force–indentation curve, the elastic modulus of each sample was evaluated by fitting the corresponding approach curve to the Hertz model[49]where F is the applied force, δ is the indentation depth, R is a radius of the spherical tip, υ is the Poisson ratio, and E is Young’s modulus.

Only a single probe was used for nanoindentation experiments in this study and the bead diameter of 3 μm was determined accurately by SEM imaging. To obtain Young’s modulus values, the force curves recorded from different locations on the hydrogels/nanocomposite surfaces were fitted to the Hertz model using a spherical tip-shaped model. The final number of complete analyzable curves were 449 for gelatin, 239 for gelatin nanocomposite, 365 for agarose, and 335 for agarose nanocomposite. For each gel, all calculated Young’s modulus values from three repetitions were collected and further analyzed using SigmaPlot (version 14.0).

Isolation of Platelets

Human blood from healthy donors who were drug-free within the previous 10 days was collected into a tube of ACD-A 1.5 mL (BD-Vacutainer, Germany). The blood tube was sealed with the parafilm and rested at room temperature for 15 min. Platelet-rich plasma (PRP) was first obtained from the blood by centrifugation at 120g for 20 min at room temperature. Platelets were further isolated from PRP in the presence of 15% acid-citrate dextrose (ACD-A, Fresenius Kabi, Germany) and 2.5 U/mL Apyrase (grade IV SIGMA, Munich, Germany) by centrifuging at 650g for 7 min. The platelet pellet was resuspended in 5 mL of suspension buffer at pH 6.3 composed of 137 mM NaCl, 2.7 mM KCl, 11.9 mM NaHCO3, 0.4 mM Na2HPO4, 2.5 U/mL Hirudin and incubated 15 min, 37 °C before recentrifuging them at 650g for 7 min. Platelet pellets were again carefully resuspended in 2 mL of suspension buffer and the blood counter (pocH-100i, SYMEX, Germany) was used to count the platelets. Afterward, the platelets were incubated for 45 min, 37 °C before use.

Confocal Laser Scanning Microscopy (CLSM)

Platelets were stained at RT in the dark for 30 min with anti-CD42a FITC antibody dye (Dianova GmbH, Hamburg, Germany) with a final concentration of 0.1 μg/mL. After that, platelets were seeded at a concentration of 3 × 105 cells/μL on the gels and stored at room temperature for 2 h. Unbound platelets were removed by rinsing with PBS. Subsequently, 4% paraformaldehyde was used to fix platelets for 30 min at RT. The samples were examined using a confocal laser scanning microscope Zeiss LSM710 (Carl Zeiss, Gottingen, Germany) at RT in the dark. The red fluorescence signal was acquired using the excitation wavelength of 488 nm (15 mW argon laser) using a 63× objective and detection in a range of 500–550 nm. ImageJ software was used to further process the images and to quantify the spread area of the platelets. The analysis was performed with SigmaPlot (version 14.0).

Scanning Electron Microscopy (SEM)

To form agarose and gelatin films, round glass coverslips (Plano GmbH, Wetzlar, Germany) of 24 mm were cleaned with 80% ethanol prior to coating with 100 μL of gels. After that, platelets of 3 × 105/μL were seeded on the gels at RT for 15 mins before fixing with 4% paraformaldehyde. The samples were washed with PBS twice. This was followed by incubation for 10 min each in ascending isopropanol series (30, 50, 70, 90, and 100%) for dewatering. The samples were incubated with 50% hexamethyldisilazane (HMDS) + isopropanol 100% for 10 min. The final step included submerging the samples with 100% HMDS (Sigma-Aldrich, Germany). The samples were allowed to dry overnight before sputtering them with gold. SEM Evo LS10 (Carl Zeiss AG, Jena, Germany) was used to image the samples. Images were taken with a magnification of 1000× and a working distance of 12.5–14.5 mm.

Single-Platelet Fluid Force Microscopy

Fluid force microscopy measurements were performed with a JPK Nanowizard 4 (Berlin, Germany) assembled with a FluidFM add-on (Cytosurge, Switzerland), positioned under an acoustic hood, and mounted on an active vibration isolation system (Micro 40, Halcyonics, Germany) to minimize the effects of surrounding vibrations. An inverted microscope (Axio Observer, Zeiss, Germany) was used from beneath to observe the platelets and to approach the cantilever to the desired location. Force spectroscopy was performed using a FluidFM nanopipette (Cytosurge, Switzerland) with an aperture of 300 nm and a nominal spring constant of 2.09 ± 0.15 N/m. The reservoir was filled with suspension buffer, calibrated by the contact-free thermal noise method, and then approached the surface. The calibrated spring constant of the nanopipette before picking the platelet in the liquid environment was 0.42 ± 0.01 N/m. By applying a pressure of −800 mbar on a platelet for 4–8 min, a single platelet was drawn to the aperture of the cantilever. For subsequent force measurements, the pressure was reduced to −500 mbar.

For measuring the force–distance curves by the cantilever with a platelet, the cantilever was immersed in PBS liquid and a setpoint of 10 nN, a z-length of 3–5 μm, and a z-speed of 2.5 μm/s was used. A platelet was picked and force map measurements were carried out on each sample at three different places with 64 force–distance curves taken from each map (10 × 10 μm2). For each gel type, three repetitions were performed. The obtained force maps were then processed with the JPK data processing software (version 6.1.120). The mean values and corresponding error bars were analyzed with SigmaPlot (version 14.0).

Conclusions

We studied the influence of fabricated hydrogels and hydrogel-nanocomposite films on the inhibition of platelet-surface activation. We found promising agarose and agarose nanocomposite materials for human blood platelet applications. The agarose hydrogel and its nanocomposites exhibit higher surface wettability, better controlled-swelling properties, and greater stiffness than gelatin, resulting in a stronger reduction of platelet adhesion and spread. The observed behavior of platelets on the fabricated gels indicates how biomimetic surfaces with antifouling characteristics govern cellular responses. As several types of NPs have antibacterial properties, agarose nanocomposites have a powerful application in the fabrication of platelet storage bags that provide antibacterial functions. The final aim would be to implement such antithrombotic surfaces in blood-contacting medical devices and medical procedures. Our results open a new venue in the development of antithrombosis materials based on agarose hydrogels, which have potential applications in implantations, platelet/blood storage bags, as well as biotechnological and pharmaceutical trials.