1. Introduction
Tissue engineering has offered the tantalizing possibility to regenerate tissues and organs, allowing the treatment of a multitude of conditions and pathologies. Despite numerous significant progresses with in vitro and small animal studies, clinical applications have been scarce [
1]. Even the most advanced solutions delivered to physicians lack sufficient vascularization within the tissue engineered constructs [
2]. This is because the diffusion of oxygen and nutrient supply present major limits on the size and complexity of bioengineered scaffolds. For this reason, vascularization of biomaterials remains the highlighted focus in tissue engineering and regenerative medicines. In this context, one main current challenge in tissue engineering is the development of biomaterials that can promote angiogenesis, ultimately integrating with the host vasculature to form anastomosis.
Angiogenesis, the formation of new blood vessels from existing ones, is a complex process. During angiogenesis, quiescent endothelial cells (ECs) from pre-existing vessels are activated by the increase in concentration of pro-angiogenic factors induced by inflammation or by hypoxia [
3]. Activated ECs proliferate and differentiate into tip cells, which results in the elongation of new blood vessels in the direction of the pro-angiogenic stimulus. This sprouting process is modulated by the migration of ECs led by tip cells, characterized by lamellipodia and filopodia in their cytoskeleton, followed by stalk cells, which are found between quiescent cells and the tip cells. Stalk cells continue to proliferate and constitute the new endothelium, while ensuring a continuum with the original vessel through regulated proliferation [
4,
5]. Once the capillary is formed, ECs secrete attractant molecules to recruit perivascular cells, which migrate along the newly formed vessels and provide stability, support cell differentiation, and regulate vessel permeability [
6].
Angiogenesis is partially modulated by the ECM, which provides essential structural support and biochemical cues for cell morphogenesis and physiological functions [
7]. Numerous strategies employing hydrogels with functionalized pro-angiogenic molecules have been proposed to promote vessel formation. Most of these approaches are based on the delivery of growth factors (GFs), such as vascular endothelial growth factor (VEGF), to facilitate vascularization in vivo [
8]. Recently, pre-vascularization of biomaterials has been proposed as an approach to promote in vitro vessel formation prior to implantation. The idea is to stimulate in vitro vessel formation within 3D biomaterials which present pre-formed channels. Different techniques to develop hydrogels with pre-formed channels have been investigated. These include the use of syringes or glass micropipettes [
9], or sacrificial templates [
10,
11,
12,
13]. To promote cell adhesion, ECM proteins (e.g., collagen, fibrin, or fibronectin) and cell adhesive molecules (e.g., RGD, YIGSR sequences) are often incorporated into the hydrogel composition [
8,
14,
15]. Besides interaction with the ECM, angiogenesis also depends on spatial presentation of pro-angiogenic cues that direct vessel sprouting and maturation [
3]. Over the past decades, various approaches have attempted to fabricate hydrogels with spatial guidance either through direct patterning of vascular cells, or through spatial distribution of pro-angiogenic molecules (e.g., VEGF, FGF, angiopoietin, YIGSR) [
16,
17,
18,
19,
20]. The use of ECM molecules presents promising outcomes for in vitro and in vivo vascularization. Nevertheless, clinical translation still remains a hurdle due to high cost and immunogenic potential of animal-origin ECM molecules.
Several important factors must be taken into account when designing hydrogels that favor endothelialization for tissue engineering and regenerative medicines: (1) presence of interconnected pores favoring cell–cell interactions and migration; (2) presence of a hollow channel having a wide range of diameters to mimic native vessels; (3) ability to promote EC arrangement leading to the formation of microvessel-like networks; (4) biocompatible composition (pharmaceutical-grade materials); (5) integration of basement membrane proteins (BM), such as laminin and collagen type IV, and other ECM proteins to induce endothelial proliferation and differentiation during angiogenesis; (6) easy fabrication protocol; and (7) cost efficient.
For vascularization purposes, porous 3D hydrogels are widely employed due to their ability to facilitate nutrient and oxygen diffusion, thus enabling cell migration [
21,
22]. Additionally, the presence of channels within porous scaffolds has been reported to promote cell growth and rapid vascularization [
23,
24]. The channels in 3D hydrogels play a key role in guiding EC arrangement and should also be utilized to induce angiogenic behavior in ECs.
Polysaccharides are widely employed as tissue engineered biomaterials due to their physicochemical properties that can mimic the ECM [
25]. In this context, we utilized 3D porous hydrogels, composed of pullulan and dextran. Notably, our team has demonstrated in several studies the versatility of pullulan- and dextran-based hydrogels, where the scaffold geometry, mechanical properties, porosity, and swelling behavior of these hydrogels could be controlled [
25,
26,
27,
28]. The hydrogel crosslinking method was previously described in numerous publications and has been patented [
29,
30]. Thus, these hydrogels have been investigated in various in vitro and in vivo studies [
29,
31,
32]. Most recently, we have demonstrated the ability to guide EC arrangement based on channel curvature on the 3D polysaccharide hydrogels [
28].
In the context of promoting in vitro vessel formation, this study aimed to develop 3D porous hydrogels with different spatial presentation of pro-angiogenic signals to guide ECs towards angiogenic behavior. The challenge of this work was to functionalize the chemically crosslinked hydrogels to promote EC adhesion and to direct sprouting through spatial guidance using pro-angiogenic cues. Here, we present a simple method to produce biomimetic 3D porous hydrogels, made from pharmaceutical-grade pullulan and dextran, with preformed microchannels (
Figure 1). To provide cells with pro-adhesive and pro-angiogenic signals, the hydrogels were functionalized using a recombinant, engineered bacterial protein polymer called Caf1. Caf1 subunits assemble into long, highly stable and flexible polymers, which are bioinert, allowing bioactive peptide motifs from the ECM and growth factors to be inserted and hence provide exquisite control over the biological signals supplied to the cells [
33,
34,
35]. In this work, we demonstrate an innovative strategy to functionalize chemical hydrogels in a spatial-controlled manner. Capitalizing on the acidic pI of Caf1, we could functionalize hydrogels simply via electrostatic interactions induced by the coating method (
Figure 1b). Then, spatial cues of the pro-angiogenic motifs were modulated through a combination of hydrogel coating and a freeze-drying process (
Figure 1c).
The developed scaffolds were evaluated based on: (1) porosity; (2) presence of the hollow channels formed within the 3D scaffolds; (3) ability to promote EC cell adhesion as well as migration; (4) ability to induce pro-angiogenic behavior of ECs. Furthermore, our approach offers a facile protocol for both scaffold fabrication and functionalization. The use of Caf1 overcomes the high cost and immunogenic potential of traditional ECM molecules. The functionalized scaffolds exhibited good EC adhesion and proliferation. Scaffolds with different spatial distribution of pro-angiogenic moieties induced different EC behaviors. Based on the results obtained from this study, we report the first work, to our knowledge, in using animal-free ECM-like molecules to control the spatial cues of hydrogel-based scaffolds, which modulates EC behavior and guides them towards angiogenic sprouting.
3. Discussion
Porous hydrogels made of pullulan and dextran were synthesized by chemical crosslinking with sodium trimetaphosphate (STMP), as previously described [
29]. First, the hydroxyl groups in the polysaccharides were activated at basic pH using NaOH, resulting in the opening of cyclic STMP and crosslinking between the polysaccharides, leading to hydrogel formation [
36]. Previous studies have demonstrated the potential of these biocompatible hydrogels as scaffolds for 3D cell culture, tissue engineering, and cell therapy applications [
25,
27,
37,
38]. However, due to the high water content (~93%) and the chemical structures of pullulan and dextran, endothelial cells do not adhere spontaneously to these hydrogels [
28,
38], as shown in
Figure 7a (NC sample). The neutral polysaccharides were cationized by incorporating diethylaminoethyl dextran (DEAE–Dex) to facilitate electrostatic interactions with the negatively-charged ECM-like molecules (Caf1-YIGSR and Caf1-VEGF). The shape and diameter of the channel remained 100 ± 20 µm before and after swelling. These observations are in correspondence with the swelling behavior observed in all formulations. Here, we demonstrated the ability to form straight microchannels with circular cross-section and controlled diameter.
Cell analysis of HUVECs seeded on functionalized scaffolds with various coating methods (SFD vs. DFD) and spatial distribution of the two Caf1 motifs (YIGSR and VEGF) confirmed that the Caf1 solution (p.I. = 4.6; 1 mg/mL; ɀ = −23.6 mV) was adequate to facilitate electrostatic interactions with cationized hydrogels (PUDNA-D20; ɀ = +29.5 mV). Through the addition of microchannels within the polysaccharide scaffolds and sufficient surface functionalization via electrostatic interactions, ECs were able to adhere, leading to good cell proliferation and cell spreading within the microchannel. In this work, spatial control of the ECM-mimicking moieties was shown to induce different EC behaviors that could be interesting for vascularization applications. An in-depth discussion on this part is presented in
Section 3.2,
Section 3.3 and
Section 3.4.
3.1. The Impact of DEAE–Dex Concentration on Hydrogel Opacity and Functionalization
As seen in
Figure 3, an increase in the concentration of DEAE–Dex added to the polysaccharide solution contributed an increase in hydrogel opacity, which limited sample visibility under the microscope. Even with a multiphoton microscope or a high-resolution confocal laser scanning microscope and sufficient image treatment and analysis, it was very difficult to locate the microchannel embedded in the middle of the sample depth. Consequently, these observations suggest that a balance between hydrogel transparency and the cationic polymer concentration need to be considered to ensure sample visibility for microscopy analysis, which is essential to monitor cell behavior in the scaffolds. Moreover, this balance must also allow sufficient interactions between the charged materials in order to facilitate cell adhesion on the functionalized scaffolds.
After a series of optimization work, by synthesizing cationized hydrogels with varying DEAE–Dex concentrations (5–100% DD:Dex
w/
w), we were able to determine the best cationic parameters to yield optimal hydrogel opacity and favorable EC behavior outcomes. At 20% (DD:Dex
w/
w), the scaffold surface and microchannel were still visible under the CLSM (
Figure 3a). More importantly, ECs were also observable after 7 days in culture (
Figure 4a). On day 7, PUDNA-D20C scaffolds facilitated better EC adhesion, where more elongated cells were present inside the channel (
Figure 4a) and cell metabolic activity was statistically higher than the metabolic activity on the rest of the other conditions (
Figure 4c). These results strongly suggest that PUDNA-D20C was the optimal hydrogel condition for most favorable EC adhesion and morphology.
3.2. Caf1-YIGSR Induced Cell Adhesion on SFD Hydrogels and Enhanced Cell Proliferation on DFD Hydrogels
Cell morphology and behavior on SFD scaffolds using Caf1-YIGSR (
Figure 5a) confirmed that the YIGSR sequence could be used as a cell-adhesive coating material.
Recent works on functionalized biomaterials have demonstrated the ability to modulate cell behavior by varying the concentration of cell-adhesive ligands in the scaffolds, with an increase in ligand concentration leading to an improvement in cell adhesion, spreading, and proliferation [
39]. The results obtained from this study are in accordance with these findings. When both the channel and pores were coated (DFD coating method: YIGSR-DFD), the amount of Caf1 protein detected on the SgC-DFD scaffolds was higher than that on the SgC-DFD scaffolds, suggesting an increase in bulk ligand concentration (
Figure 9). As a result, cell morphology on the DFD scaffolds greatly improved (
Figure 5a) and greater cell metabolic activity was observed after 7 days in culture (
Figure 5b). The decrease in cell metabolic activity on day 9 could be due to high cell confluency. This is supported by the fact that no signs of cell death were observed after 9 days. In conclusion, the increase in spatial distribution of the YIGSR sequence, contributed to an increase in ligand bulk concentration on the scaffold, leading to an enhancement of EC morphology and behavior.
3.3. Caf1-VEGF Induced Cell Migration and Angiogenic Sprouting Depending on Its Spatial Presentation on Porous Hydrogels
When Caf1-VEGF was used alone as the coating material in the DFD method, few cells adhered inside the channel and did not completely line the channel edge. Adhered cells showed polarization characteristics and sprouting-like behaviors. It is well known that VEGF is a pro-migratory factor that induces filopodia elongation in ECs during angiogenesis [
5]. This explains why ECs in VEGF-DFD scaffolds showed filopodia structure resembling migration behaviors (
Figure 5a). Additionally, cells on the top side of the channel protruded and connected to cells on the bottom side (
Figure 6a). Due to lack of cell-adhesive moieties (i.e., Caf1-YIGSR), not enough cells adhered inside the channel, resulting in lower cell proliferation, as demonstrated by the lower cell metabolic activity (
Figure 6b). This drop in metabolic activity could also be linked to cell confluency on day 9.
Taking these results into consideration, CoC scaffolds were prepared, where two different Caf1 proteins were presented on the hydrogels in different spatial distribution. First, the channel was coated with Caf1-YIGSR, then the pore-filled region was coated with Caf1-VEGF (
Figure 1c). On co-coating scaffolds, the cell morphology and behaviors significantly altered. Adhered cells inside the channel started to migrate outwards to the pore-filled region. Some cells even exhibited filopodia structure. These results strongly suggest that different spatial presentations of Caf1-VEGF on porous hydrogels drive distinct cell behaviors.
During the last decades, cell–ECM interaction research has shown that when cell-adhesive molecules were spatially presented to the cells in different manners, they induced different patterns of cellular behavior [
40,
41]. In the case of CoC hydrogels, cell adhesion was achieved thanks to the contribution of the Caf1-YIGSR coating in the channel during the first coating step (
Figure 4). The presence of Caf1-YIGSR facilitated proper cell adhesion, where cells could form a strong anchor to the substrate at focal complexes [
4]. The presence of Caf1-VEGF promoted protrusion formation of ECs and transformed protrusion into forward movement. This explains the observation of filopodia structure, stress fibers, and polarization of HUVECs seeded on the CoC hydrogels (
Figure 7a). The adhered cells sensed migratory signals from the VEGF sequence, which stimulated cell migration processes. In other words, our results suggest that the presence of Caf1-VEGF moieties in the pores created cell directionality, leading to cells moving from the channel outwards to the porous region (exterior of the channel).
3.4. Synergistic Effects of Caf1-YIGSR and Caf1-VEGF on EC Morphologies and Behaviors
Taken the outcomes discovered from CoC hydrogels, a question regarding Caf1-YIGSR and Caf1-VEGF spatial distribution on hydrogel was considered. What will be the effect of these two Caf1 proteins on cell morphology and cell behavior, if they were both presented on the hydrogel in similar spatial organization? This question led to the creation of CoCmx hydrogels, where Caf1-YIGSR and Caf1-VEGF solutions were mixed in equal parts (50:50
v/
v) and used to coat the scaffolds via the DFD method. Here, both the channel and the pores were functionalized with Caf1-YIGSR and Caf1-VEGF at the same time. Initial inspection of cell morphology on CoCmx scaffolds showed good cell adhesion (where cells fully lined the channel) and elongated filopodia (which indicated cell sprouting and migration) (
Figure 7a, top panel). Moreover, migrating cells connected with non-migrating cells both inside the channel and outside the channel (
Figure 7a, bottom panel). These observations suggested the synergistic effect of Caf1-YIGSR and Caf1-VEGF. Both VEGF and YIGSR are known to play a role in angiogenesis, with YIGSR contributing to cell adhesion, cell–cell interactions, and tubule formation, while VEGF stimulates cell migration [
4,
42,
43]. The presence of both Caf1-YIGSR and Caf1-VEGF inside the channel induced a stabilizing adhesive effect on ECs. These ECs then migrated towards the VEGF stimulus that was also available in the pores of the scaffolds. Consequently, the dual presence of YIGSR and VEGF sequences, both exhibiting angiogenic effects, promoted greater EC proliferation. These ECs possibly produced their own ECM, which further stabilized the vessel-like network and induced EC differentiation towards angiogenic phenotypes. This explains why elongated migrating cells were observed in both the channel section and the porous regions outside the channel only on CoCmx scaffolds (
Figure 8).
In other words, the dual presence of Caf1-YIGSR and Caf1-VEGF functionalized on our 3D porous hydrogels created a synergistic effect on seeded HUVECs. Previously, a Caf1 mosaic co-polymer containing two pro-osteogenic motifs was seen to promote the early stages of bone formation in primary human mesenchymal stromal cells in a 2D system [
35]. The synergistic effect described here further demonstrates the benefits of the Caf1 system, where bioactive peptides can be easily introduced and placed in close proximity in a single material, allowing these synergistic effects to take place. Thus, these effects mimicked the in vitro angiogenesis, where ECs adhered and became activated, then proliferated and differentiated into tip cells, resulting in elongation in the direction of the VEGF stimulus.
3.5. Comparison of the Developed Method with Current Vascularization Strategies
Over the past decades, numerous attempts have been made to develop vascularized constructs using three main strategies: microfluidic-based approaches, 3D bioprinting, and organoids/spheroids-based techniques. The readers are invited to read more on this topic in the published review [
2].
The use of ECM-based membranes integrated in microfluidic platforms has allowed researchers to develop more physiologically relevant models thanks to the ability to perfuse the systems. However, most models require soft lithography for materials fabrication, which is expensive and is difficult to be used by a wide end-user’s range.
The use of 3D additive manufacturing, such as fuse deposition modeling (FDM), facilitates printing of sacrificial components that better mimic in vivo vasculature. However, these techniques often require several manufacturing steps and still present major issue in terms of resolution. Most vessel geometries remain relatively simple and the vessel diameters are in the range of hundreds of microns. Channels obtained using co-axial bioprinting or with sacrificial bioinks remain in the same range. More recently, the use of laser-assisted bioprinting (LAB) offers high resolution (5–10 µm) of printed channels, automation, reproducibility, and high throughput. Similarly, the use of Vat photopolymerization-based bioprinting opens new possibility to create complex vascular patterns with high precision and high resolution. However, these approaches are still far from translation due to the high cost of equipment, and the need to work with photosensitive materials and photoinitiators further limit their application for therapy.
Spheroids and organoids are another alternative approach to promote the vascularization of hydrogel constructs. They offer the ability to recapitulate the microenvironment, thus present great potential as vascularized models. However, to reach a substantial quantity of tissue, a large number of cells are needed. The use of ECM proteins with heterogeneous composition and high immunogenic potential (e.g., collagen and Matrigel), further prevents translation of these models in the industry and clinical settings.
In this work, we employed a simple method to form microchannels at the microcapillaries range (≥100 µm). Although the filament templating/removal technique is limiting in terms of producing complex designs, it enables high reproducibility and facile fabrication. Our system, porous hydrogels with channels, functionalized in a spatial-controlled manner, present several advantages compared to other aforementioned vascularization strategies.
Compared to other hydrogel-based vascularization strategies, our polysaccharide-based hydrogels support long-term cell culture of up to 9 days, as demonstrated in this study, and could be kept up to 14 days in other studies without being degraded [
27]. With a small amount of protein (~0.25–1.8 µg/mg hydrogel), we were able to induce initial cell adhesion, followed by cell proliferation and migration on functionalized scaffolds. Thus, the spatial cues (e.g., YIGSR and VEGF) further direct cell migration mimicking the first step of sprouting angiogenesis. Even though the electrostatic bonds are weaker than covalent bonds, our functionalization method was stable enough to enable observation of grafted Caf1 on the hydrogels (as shown in
Figure S5). Moreover, the concomitant presence of channels and pores offers the possibility to promote vascularization of tissue constructs, while enabling co-culture with other cell types for the development of different bioengineered models. Caf1 molecules are manufactured in vitro using bacterial expression systems in high quantities and with a lower cost [
33]. Thus, the animal-free origin of Caf1 would reduce immunogenic potential, making them ideal materials for implantable constructs. Our coating method based on ionic interactions is performed in a one-step process and uses green chemistry. In this study we focused on YIGSR and VEGF, but in the future, it will be possible to use the same strategy to incorporate other Caf1 peptides to confer new properties to the material. Finally, from an industrial point of view, our fabrication technique and the choice of materials are highly beneficial: The production method is simple and can be easily scaled-up and the freeze-dried hydrogels allow for long-term storage, all contributing to low-cost production and maintenance.
4. Materials and Methods
4.1. Materials
Pullulan (Mw 200 kDa) and dextran (Mw 500 kDa) were obtained from Hayashibara Inc. (Okayama, Japan) and Pharmacosmos (Holbaeck, Denmark), respectively. FITC-dextran (dextran labeled with fluorescein isocyanate, TdB consultancy® (Prince George, BC, Canada)) was used to label the hydrogels. All other chemicals were obtained from Sigma-Aldrich® (Saint-Quentin-Fallavier, France). Caf1-YIGSR and Caf1-VEGF as freeze-dried powder were provided to us by Newcastle University (Newcastle, UK).
4.2. Hydrogel Synthesis: 3D Porous Polysaccharide-Based Hydrogel with Microchannel
Briefly, a solution of pullulan and dextran (75:25
w/
w) and NaCl was prepared in ultrapure water. This solution is referred to as PUDNA. Then, NaOH 10M was added to the PUDNA solution to activate the hydroxyl groups before reacting with the crosslinker STMP (sodium trimetaphosphate) (3%
w/
v) at room temperature under magnetic stirring. The crosslinked solution was poured in between two glass slides, separated by polypropylene suture filaments ø 70 µm (6.0, Ethicon
®) (Raritan, NJ, USA) and two spacers of 250 µm thickness, before crosslinking in an oven at 50 °C for 20 min. This incubation step was carried out to facilitate the crosslinking reaction and to form microchannels within the hydrogel. Afterwards, the hydrogels were cut into discs of 5 mm in diameter using a biopsy disc-cutter from Harris Uni-Score (Sigma-Aldrich
®) (
Figure 1a).
Hydrogels were neutralized in PBS 10X and washed in distilled water until equilibrium (ca. 15 µS/cm). The conductivity was measured with an Orion 145 A+ conductivity meter purchased from Thermo Fisher Scientific (Asnières-sur-Seine, France). A second wash was performed in NaCl 0.025% (Sigma-Aldrich®) until equilibrium (ca. 500 µS/cm). Finally, the hydrogels were freeze-dried to promote pore formation.
The freeze-drying protocol consisted of three stages: freezing under atmospheric pressure from 15 °C to −20 °C at a constant rate of −0.1 °C/min, followed by a phase at constant temperature of −20 °C for 90 min. Primary drying was performed at low pressure (0.001 mbar) and −5 °C for 8 h and secondary drying at 30 °C for 1 h [
26].
4.3. Hydrogel Characterization
4.3.1. SEM
The topography of freeze-dried hydrogels was observed using the JEOL JSM-IT100 system (software InTouch Scope v.1.060) under low-vacuum conditions. The SEM system was located at the Institute Jacques Monod (Paris, France).
4.3.2. Porosity
The porosity of hydrogels was determined based on a published protocol which calculates the water amount absorbed in the hydrogel before and after manual squeezing tests [
44]. Experiments were performed by soaking 5 samples in PBS 1X in a 24-well cell culture plate (Corning
®) (Corning, NY, USA) for 2 h under mechanical shaking. Samples were then weighed after removing the excess liquid by placing them on the plastic lid. This was considered the weight of the swollen gel (M
swollen, mg). Following this step, samples were weighed again after squeezing out the remaining liquid using tissue paper and gentle pressing using a spatula. This was considered the “squeezed” weight (M
squeezed, mg). The porosity calculated by this method corresponds to the large pores that entrap water molecules free or weakly bound to the polysaccharide matrix that are release by gentle mechanical compression. The pore volume percentage was calculated using Equation (1). At least three scaffolds were analyzed per condition. Results were expressed as mean values ± SD.
4.3.3. Swelling Ratio
Scaffolds were weighed before (M
dry) and after (M
swollen) rehydration in PBS 1X for 48 h. The swelling ratio was determined using Equation (2). At least three scaffolds were analyzed per condition. Results were expressed as mean values ± SD.
4.3.4. Water Content (WC)
The water content was calculated by using the sample weight after 48 h post-rehydration (M
swollen) and the sample weights before rehydration (M
dry). The water content was calculated using Equation (3). At least three scaffolds were analyzed per condition. Results were expressed as mean values ± SD.
4.4. Hydrogel Functionalization via Electrostatic Interactions
4.4.1. Caf1 Solution Preparation
To assure cell adhesion onto the polysaccharide-based hydrogels, recombinant, engineered Caf1 proteins displaying pro-adhesive and pro-angiogenic peptide motifs were used to functionalize the hydrogels. Briefly, the sequence encoding the peptide was inserted into the
caf1 gene, present on a standard expression plasmid, and the protein was expressed and purified from an
E. coli culture using tangential flow filtration and size exclusion chromatography [
33,
35].
The Caf1 proteins with cell-adhesive motifs are called Caf1-YIGSR and Caf1-VEGF. Solutions of 1.0 mg/mL (ɀ = −23.6 mV for Caf1-YIGSR and ɀ = −21.7 mV for Caf1-VEGF) were prepared by diluting the freeze-dried powder in miliQ water at room temperature and stored at −20 °C. These solutions were then thawed on the day of hydrogel coating and allowed to cool to room temperature, before being used.
4.4.2. Cationization of Polysaccharide Hydrogel
Briefly, a predetermined amount of diethylaminoethyl (DEAE)–dextran (Mw 500 kDa) from Pharmacosmos (Holbaeck, Denmark) was added into the standard hydrogel solution to obtain a solution at various concentrations: 5–20% (DD:Dex w/w; ɀ = +29.5 mV) and mixed well at room temperature (RT) until fully dissolved. The hydrogel precursor solution was degassed overnight at RT and used for hydrogel synthesis the next day.
4.4.3. Spatial-Controlled Hydrogel Coating
To facilitate electrostatic interactions, positive charges were added to the hydrogel network (by incorporation of DEAE–Dextran) to react with the negatively charged protein solution (pI = 4.46). Once the hydrogels were synthesized and rinsed thoroughly (
Section 4.1), they were immediately coated via the syringe vacuum-induced method (
Figure S1) (100 µL/ gel) and incubated for 2 h at RT. This coating step was performed either only before, or both before and after the freeze-drying step to coat the gel channel only (SFD coating method) or both the channel and pores (DFD coating method) (
Figure 1).
4.5. Cell Culture and Cell Seeding
Human umbilical vein endothelial cells (HUVECs) (ATCC-CRL-1730) purchased from ATCC® (Manassas, VA, USA) were maintained and subcultured in T75 surface-treated flasks (Corning®) in complete endothelial growth medium (EGM-2) (Lonza) following the manufacturer’s recommendations. To prevent bacterial contamination, 1% antibiotic-antimyotic (AA 100X) (Gibco™) purchased from Thermo Fisher Scientific, was also added to the complete growth medium. Cells splitting was performed according to manufacturer and kept in an incubator prior to use (37 °C, 5% CO2).
Prior to cell seeding, hydrogels were sterilized under UV light for at least 1 h. Cells were first detached with 1 mL of Trypsin solution (1X, Gibco) at 37 °C for 5 min. Trypsin was inactivated by performing cell dispersion in EGM-2, followed by centrifugation and cell counting. Cell dilution in cell culture medium was conducted to reach the desired concentration. Cell loading was performed via the syringe vacuum-induced method to ensure cell seeding only inside the preformed microchannel. Briefly, hydrogels and cell suspension were introduced in a 10 mL syringe barrel. A 3-way stopcock was used to close the system and the plunger was pulled to make cell suspension circulate inside the channels. Then, cell-loaded hydrogels were placed in a 24 well-plate (Corning®), complete cell medium was added, and the plates were placed in an incubator.
The optimal seeding density was determined to be 5.0 × 10
6 cells/mL. Culture medium (EGM-2) was refreshed every 2–3 days. To facilitate cell lining of the channels, the hydrogels were turned 180° twice following the protocol described in
Figure S2.
4.6. Cell Metabolic Activity
Cell metabolic activity was determined using the In Vitro Toxicology Assay Kit (Resazurin-based, TOX8-1KT, Sigma-Aldrich, France). Briefly, cells were cultured as previously described (
Section 4.3). On day 2, 4, 7, and 10, cell medium was removed and 0.5 mL of fresh culture medium containing 10% resazurin solution was added. After 3 h of incubation (37 °C, 5% CO
2), 100 μL (in triplicates per sample) of the supernatant was transferred to a 96-well plate. Fluorescence was measured using an Infinite M200 Pro microplate reader (TECAN
®) at 560Ex/590Em. All samples were analyzed in triplicate, in three different experiments. Results were expressed as mean values ± SD.
4.7. Cell Staining for Confocal Microscopy
Cellularized hydrogels were fixed with paraformaldehyde 4% (Sigma-Aldrich®) in PBS for 1 h at 4 °C. After rinsing with PBS, membranes were permeabilized with Triton X-100 (Sigma-Aldrich®) 0.1% in PBS for 1 h at RT. Actin filaments were labeled by incubation with TRITC-conjugated phalloidin (Sigma-Aldrich®) (1/200, 1 h incubation time at RT) and nuclei were stained with DAPI (1/2000). Samples with FITC (λex 488 nm) and cellularized samples stained with phalloidin actin marker (λex 561 nm) and DAPI nuclear marker (λex 405 nm) were observed using a Leica SP8 confocal microscope. Images were acquired using the LSA-X software (LAS X Core 3.7.6) and image analysis was performed with ImageJ/Fiji software (Window, version 153, Java8).
4.8. Immunofluoresence Staining of the Caf1 Protein Polymers
In order to confirm the presence of Caf1 protein polymer functionalized on the hydrogel channel (SFD coating method), we conjugated Caf1 with fluorescent markers. Briefly, the primary antibody YPF19 (Yersinia pestis F1 antigen antibody, mouse monoclonal, GTX28275) from GeneTex (Irvine, CA, USA) was prepared in PBS (1/200) to conjugate the Caf1 presented on hydrogels. The functionalized, freeze-dried hydrogels were incubated overnight at 4 °C. After thorough washing in PBS, the samples were incubated with a secondary antibody (Alexa Fluo 647, goat anti-mouse, 1/1000) for 45 min at 37 °C. Finally, the samples were washed in PBS several times for at least 30 min. Then, the samples were observed using CLSM.
5. Conclusions
In this study, 3D porous polysaccharide-based hydrogels made of pullulan and dextran that do not promote cell adhesion, were functionalized with animal-free ECM-like molecules via electrostatic interactions promoted by the incorporation of cationized dextran (DEAE–dextran). Although the cationization resulted in slightly opaque samples, we were still able to visualize cell morphology and evaluate in vitro cellular behaviors using 3D microscopy. Our work has demonstrated that electrostatic bonding between the charged hydrogels and Caf1 molecules was stable enough to induce adequate cell adhesion and proliferation. The spatial cues on these scaffolds were controlled through a combination of hydrogel coating and a freeze-drying step. On one hand, ECs adhered and showed sprouting according to how they exposed the cell-adhesive Caf1-YIGSR. On the other hand, the VEGF-like molecule (Caf1-VEGF) functioned as a migratory factor in the presence of the adhesive moiety (Caf1-YIGSR). When ECs were exposed to both Caf1-YIGSR and Caf1-VEGF, they exhibited angiogenic behavior. These results strongly suggest that our functionalized polysaccharide-based hydrogels can provoke different EC behaviors thanks to spatially controlled presentation of these ECM-like, animal-free, pro-angiogenic molecules. Moreover, we also demonstrated that scaffold functionalization via electrostatic interactions was sufficient to promote cell adhesion and cell proliferation for a week, which allowed ECs to further differentiate into their angiogenic phenotypes when exposed properly to the different Caf1 moieties.
The novel approach described here represents an advance in the study of the effect different peptide sequences of the ECM have on ECs behavior. This work represents a proof of concept and opens the door to future studies to determine the effect of other spatial combinations using different Caf1 motifs in different cell types. The pro-angiogenic materials prepared here could be implanted in vivo for regenerative medicine applications. Furthermore, previously in the team, we have demonstrated the formation of soft tissue constructs (e.g., liver spheroids) using the non-functionalized polysaccharide hydrogels [
27,
45] (Le Guilcher et al., 2022 under revision). These 3D hepatic constructs showed long-term liver functions, including biliary functions, holding promise to be used as 3D models of the liver for theragnostic purposes. The developed polysaccharide hydrogels could be further optimized and integrated with the aforementioned hepatic constructs to build better organ-specific in vitro models. In the near future, we hope to contribute to the translation of vascularized constructs towards clinical applications and drug development.