Controlling placental spheroid growth and phenotype using engineered synthetic hydrogel matrices

Abstract

The human placenta is a complex organ comprised of multiple trophoblast subtypes, and inadequate models to study the human placenta in vitro limit the current understanding of human placental behavior and development. Common in vitro placental models rely on two-dimensional culture of cell lines and primary cells, which do not replicate the native tissue microenvironment, or poorly defined three-dimensional hydrogel matrices such as Matrigel™ that provide limited environmental control and suffer from high batch-to-batch variability. Here, we employ a highly defined, synthetic poly(ethylene glycol)-based hydrogel system with tunable degradability and presentation of extracellular matrix-derived adhesive ligands native to the placenta microenvironment to generate placental spheroids. We evaluate the capacity of a hydrogel library to support the viability, function, and phenotypic protein expression of three human trophoblast cell lines modeling varied trophoblast phenotypes and find that degradable synthetic hydrogels support the greatest degree of placental spheroid viability, proliferation, and function relative to standard Matrigel controls. Finally, we show that trophoblast culture conditions modulate cell functional phenotype as measured by proteomics analysis and functional secretion assays. Engineering precise control of placental spheroid development in vitro may provide an important new tool for the study of early placental behavior and development.

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1. Introduction

The human placenta provides nutrients and immune protection to the developing fetus, but inadequate research models have limited our understanding of the behavior and development of this complex organ. Disordered placental development can lead to life-threatening complications for both the mother and fetus. These aberrant placental behaviors are challenging to study both in situ and in animal models,¹ which are poorly homologous to human placental development.² Trophoblast cells form the outside of the blastocyst and develop into the complicated three-dimensional (3D) architecture of the placenta, comprised of three primary trophoblast subtypes with varying functions:³ cytotrophoblasts (CT), syncytiotrophoblasts (ST), and extravillous trophoblasts (EVT). Trophoblasts are known to express pan-trophoblast markers (GATA3) and cytokeratin-7 (KRT7).^4,5 CT are the proliferative progenitor to the ST and EVT subtypes and express integrin alpha-6 ⁶ (ITGA6) and e-cadherin-1 (CDH1).^7–9 ST line the intervillous space, express human chromogranin beta-3 (CGB3) and syndecan-1 (synd-1),^9–11 and secrete a majority of the human chorionic gonadotropin beta (hCGβ) produced by the placenta.^8,9 EVT invade into the maternal decidua, express integrin alpha-5 ¹² (ITGA5) and human leukocyte antigen-G (HLA-G),¹³ while also secreting hCGβ¹⁴ and matrix metalloproteinase-2 (MMP2).¹⁵ The placental architecture and trophoblast function are critical to maintaining a healthy pregnancy.^16,17

The complicated placental architecture is not well recapitulated with current in vitro models which are commonly trophoblast-like cell lines grown in two-dimensional (2D) systems.¹⁸ Though term placentas are widely available as discarded tissues, they do not reveal the dynamic changes of the placenta throughout pregnancy, and studying some trophoblast subtypes are difficult due to their low numbers at term.^12,13,19,20 As there are ethical and technological barriers to studying early human placental dynamics and animal models do not readily replicate human placental physiology,²¹ better in vitro models are needed to study the early human placenta. In place of hard-to-access primary early gestation trophoblasts, trophoblast-like cell lines, such as choriocarcinoma lines JEG-3, JAR, and BeWo, are commonly used to model pregnancy in vitro. JEG-3 cells have been shown to exhibit EVT-like phenotypic behavior, while JAR cells exhibit villous trophoblast-like behavior, and BeWo cells are typically used to model the ST phenotype.¹⁸

Trophoblast cells and placental spheroids are commonly grown in 2D on polystyrene culture systems that do not recapitulate native extracellular matrix (ECM) cues and the native 3D placental environment.²² To enhance these systems, groups often use ECM like collagen I,^5,9 decellularized placenta,²³ Matrigel™,²⁴ or fibronectin^25,26 layered on 2D surfaces; however, 2D ECM systems do not provide 3D signaling and mechanical forces, which impact cell morphology and function. 3D cell culture systems, such as Matrigel,^27–29 Geltrex™,³⁰ and gelatin methacrylate,^31–34 have been employed to create villous-like structures and spheroids using primary isolated trophoblasts; however, these naturally derived hydrogel systems have high variability in their ECM components and limited control over the environmental cues presented, which may lead to poor experimental reproducibility.

Here we design a library of synthetic hydrogels to enable precise control over 3D matrix cues to influence placental spheroid proliferation and differentiation in an in vitro culture system (Fig. 1A). We hypothesized that a synthetic 20 kDa 4-arm poly(ethylene glycol)-maleimide (PEG-mal)-based hydrogel platform modified with ECM-derived adhesive ligand sequences and crosslinked with protease-sensitive linkers would enable the expansion of trophoblasts in 3D and potentially modulate trophoblast function. PEG is a synthetic polymer that provides minimal batch-to-batch variability and can provide precise control over matrix cues such as ECM composition and growth factors. This system has been used to culture primary islet spheroids,^35–37 intestinal spheroids,³⁸ and mesenchymal stem cells,^39,40 among others.^41–45 The maleimide functional group enables a rapid Michael-type addition reaction with free thiols, which can be introduced by the inclusion of terminal cysteines in peptide sequences. We introduce adhesion ligand sequences derived from ECM components present in the placenta such as RGD (fibronectin), GFOGER (collagen), YIGSR (laminin), and IKVAV (laminin). Protease-sensitive crosslinkers VPM, GDQ, and GPQ-W and nondegradable dithiothreitol (DTT) and PEG-dithiol (-DT) crosslinkers were evaluated to determine the influence of matrix protease sensitivity on placental spheroid growth within a 3D matrix. We also compared our synthetic hydrogel platform against natural protease-degradable ECM-rich matrices Matrigel and collagen IV,⁴⁶ and non-protease-degradable, non-cell adherent agarose and alginate. We demonstrate that degradable synthetic hydrogels support placental spheroid development with superior viability and function relative to Matrigel controls; further, we find that environmental cues within the 3D culture environment modulate trophoblast phenotype and placental spheroid architecture and function.

Fig. 1 Tunable synthetic PEG-based hydrogel library composition and degradation dynamics. (A) schematic of 4-arm poly(ethylene glycol)-maleimide (PEG-mal) modified with ECM-derived adhesive ligand sequences and crosslinked with protease-sensitive linkers to form a hydrogel matrix for trophoblast culture. (B) Hydrogel degradation images over 8 hours when cultured with 0.005 U ml⁻¹ collagenase A and (C) image quantification normalized to 0 h gel area (n = 3). Data shown as mean ± SEM. Degradation analyzed by two-way ANOVA with the Geisser–Greenhouse correction and Dunnett's multiple comparisons test to Matrigel; ns = not significant, * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

2. Materials and methods

2.1. Materials

Crosslinker and adhesion ligand sequences were purchased from GenScript as outlined in Table S1.† Cell culture reagents were purchased from Fisher Scientific unless otherwise noted. DTT was purchased from ThermoFisher (R0861), and PEG-DT (M_n = 1000) was purchased from Sigma (717142). Matrigel (Corning™ 356237) and human collagen IV (Corning™ 354245) were purchased from ThermoFisher Scientific. Agarose (A9045), calcium carbonate, and glucono-delta-lactone (GDL) were purchased from Sigma. Sodium alginate was purchased from Novamatrix (4200101).

2.2. Cell lines

Human choriocarcinoma cell lines, JEG-3 (HTB-36) and JAR (HTB-144) were purchased from the American Type Culture Collection (ATCC). BeWo cells were kindly gifted from Dr Rachel Riley at Rowan University. JEG-3 and JAR cells were cultured in MEM (11095098) and RPMI 1640 (11875119) respectively, supplemented with 10% fetal bovine serum (FBS, 26140095), 1× antibiotic–antimycotic (15240062), 1× non-essential amino acids (11140050), 1 mM sodium pyruvate (11360070), and 10 mM HEPES (15630080). BeWo cells were cultured in Ham's F-12K (Kaighn's, 21127030) media supplemented with 15% FBS, 1× antibiotic–antimycotic, 1× non-essential amino acids, 1 mM sodium pyruvate, and 10 mM HEPES. Cells were maintained and assayed under standard conditions at 37 °C with 5% CO₂ in a humidified incubator.

2.3. Hydrogel preparation

JEG-3, JAR, and BeWo cells were passaged with trypsin around 60–80% confluent from 2D culture in standard conditions and cultured for up to 7 days in 3D hydrogel matrices (Fig. 1A) made of 5% (w/v) 20 kDa 4-arm poly(ethylene glycol)-maleimide (PEG-mal) macromer with 1 mM adhesive ligands (RGD, RDG, GFOGER, YIGSR, IKVAV) and degradable (VPM, GPQ-W, GDQ) or nondegradable (DTT, PEG-dithiol) crosslinkers as previously described.⁴⁷ Briefly, PEG-mal, adhesive ligands, and crosslinkers were resuspended in DPBS, without magnesium or calcium chloride, except for IKVAV which was resuspended in 10% DMSO in DPBS, and the pH of all reagents was adjusted to 7 with sodium hydroxide as necessary. Matrigel was diluted to 5 mg mL⁻¹ with cell-specific media, and placenta-derived human collagen IV was used as provided. These gels were made on culture plates heated to 37 °C for faster gelation. For 2% sodium alginate gels, 120 mM sodium alginate and 20 mM glucono-delta-lactone (GDL) solutions were made using DPBS without magnesium and calcium chloride and incubated at 4 °C for >12 hours for homogenization. 60 mM calcium carbonate was added to the sodium alginate solution before use, and this solution was mixed at a 1 : 1 ratio with the GDL solution. Low-gelling temperature agarose was resuspended in DPBS with magnesium and calcium chloride for a 2% solution and heated to 100 °C for 30 minutes before use. These gels were made on culture plates cooled to −20 °C for faster gelation. Single cells were resuspended in the respective gels with 100 000 cells in 15 μL gels or 100 000 cells plated on a 24 well culture-treated plate for 2D conditions. 1 mL of cell-specific media was added to the wells and collected and replaced on days 1, 3, and 7.

2.4. Degradation studies

1 mM RGD was resuspended in 95% DPBS with 5% Alexa Fluor™ 647 NHS Ester (Succinimidyl Ester) (A37573) and incubated on a vortex for 1 hour. Hydrogels were made as described above with the labeled RGD and washed in DPBS to remove any unbound RGD. The gels were incubated with 0.005 U mL⁻¹ collagenase A (Sigma # 10103578001) at 37 °C. These gels were imaged over 8 hours on the EVOS FL auto live cell imaging system and Nikon Eclipse TE300 inverted microscope at 0, 2, 4, 6, and 8 hours. The collagenase was replaced before each time point. Image analysis was performed on gel total area using FIJI and normalized to the 0-hour time point.

2.5. Cell viability

Cell viability in the various hydrogels was assessed on days 1, 3, and 7 through confocal imaging on a Leica SP8 white laser confocal staining with 2 μM CalceinAM (Invitrogen C1430) and 1 μM ethidium homodimer (Invitrogen E1169) in DPBS or cell-specific media for 30 minutes at 37 °C. Percent viability image analysis and live average size was performed on max-projected z-stacks using FIJI analysis of the total area of the CalceinAM channel over the total area of CalceinAM and ethidium homodimer for percent viability or average size of the CalceinAM channel for average live size, respectively, using the same threshold, size, and circularity constraints for all images.

2.6. Cell metabolism

AlamarBlue (Invitrogen DAL1100) was used following the manufacturer's instructions for 90 minutes at 37 °C with triplicate technical replicates per gel or 24 well plate for 2D cultured cells on days 1, 3, and 7. Fluorescent values were read using the BioTek Synergy H1 Plate Reader at ex/em 560/590 using a gain of 50. Cell-specific cell-free media was used as a background control and subtracted from each sample.

2.7. Cytohistochemistry

Hydrogels were fixed with 4% paraformaldehyde (AAJ19943K2) at room temperature for 20 minutes on day 7 and stored at 4 °C in DPBS or cell-specific media until staining. 0.25% Triton-X 100 diluted in DPBS was used for permeabilization for 30 minutes at room temperature. Gels were washed in DPBS + 1% BSA and blocked with Power Block™ Universal Blocking Reagent (HK085-5K) for 90 minutes then 2–4 drops of goat serum (Biogenex Laboratories HK1129K) for 30 min at room temperature. Gels were incubated for 90–120 minutes at room temperature in primary antibodies (rat anti-human integrin alpha 6, 1 : 50, R&D systems #MAB13501; mouse anti-human SDC1 sydecan-1, 1 : 1000, biorbyt #orb153547) or isotype controls (Rat IgG2a, R&D systems #MAB006; mouse IgG1, Novus #NBP196983) diluted in DPBS + 1% BSA, washed 3 times for 5 minutes with DPBS + 1% BSA, and incubated in secondary antibodies AF488 goat anti-mouse IgG (ab150113) or AF680 goat anti-rat IgG (ab175778) diluted 1 : 500 in DPBS + 1% BSA for 1 hour at room temperature. Gels were washed as above and transferred to slides with ProLong™ Gold Antifade Mountant with DAPI (Invitrogen #P36931) for 24 hours to cure at room temperature, then kept at 4 °C until imaging on a Leica SP8 white laser confocal microscope. Images were analyzed with Fiji using the analyze particles function with the same threshold, size, and circularity constraints for all images.

2.8. ELISA

Media was collected after incubation with the hydrogels on days 1, 3, and 7 and stored at −80 °C until use with any cell debris discarded. Human chromogranin beta (hCGβ) (R&D systems; DY9034) and matrix metalloproteinase-2 (MMP2) (R&D systems; DY902) secretion into media at each time point was measured using the R&D system DuoSet ELISA kit and corresponding ancillary reagent kit according to the manufacturer's instructions. Analysis was performed in duplicate. Corresponding concentrations were calculated using a four-parameter logistic curve from MyAssays.com.

2.9. Western blotting

JEG-3, JAR, and BeWo cells were cultured in 2D conditions or encapsulated as above in PEG-RGD-VPM and Matrigel hydrogels and lysed in 1× RIPA lysis buffer (Millipore 20-188) diluted in DI water with 1 : 50 Halt™ protease & phosphatase inhibitor cocktail (1861284) for 10 minutes on ice and subsequently centrifuged at 13 000g for 10 minutes for supernatant collection. Samples were stored at −80 °C until the electrophoresis run. Pierce™ BCA protein assay kit (23225) was used to quantify protein concentration. 10 μg of protein from each sample was combined with 1× Bolt LDS sample buffer (1 : 4, Invitrogen B0007) and 1× Bolt reducing agent (1 : 10, Invitrogen B0009) according to the manufacturer's instructions. Samples were incubated at 100 °C for 10 minutes to denature proteins. Electrophoresis was performed in NuPAGE MES SDS running buffer (Invitrogen NP0002) on Bolt BisTris 4–12% 1.00 mm mini gels (NW04120BOX) for 32 min at 200 V. The proteins were transferred using iBlot transfer stacks (Invitrogen IB23001) to a nitrocellulose membrane with a 0.2 μm pore size using the iBlot 2 P0 preset method for 1 minute at 20 V, 4 minutes at 23 V, and 2 minutes at 25 V. The membrane was stained with Ponceau S (Cell Signaling #59803) according to the manufacturer's instructions to confirm protein transfer. The membrane was washed in TBST (VWR 10791-794 with added 0.05% Tween 20: final 0.1% Tween 20) and blocked in TBST with 5% (w/v) nonfat milk (Amazon, 138 Foods) for an hour, then incubated at 4 °C overnight in TBST with 5% (w/v) BSA with anti-HLA-G (Cell Signaling #79769, 1 : 1000) and α/β-tubulin (Cell Signaling #2148, 1 : 1000). The membrane was washed with TBST and incubated in TBST + 5% (w/v) nonfat milk and anti-rabbit IgG HRP-linked antibody (Cell Signaling #7074, 1 : 1000) for 1 hour then washed in TBST. Bio-Rad clarity western ECL substrate (1705061) was used according to the manufacturer's instructions for a 5-minute incubation. The membrane was imaged with Analytik JenaTM UVP ChemStudio with an optimized exposure time using VisionWorks software.

2.10. Proteomics

100 000 JAR cells were encapsulated as stated above in degradable synthetic (PEG-RGD-VPM) hydrogels, degradable natural (Matrigel) hydrogels, or 2D-cultured with media changed on days 1 and 3. On day 7, samples were washed with PBS and lysed with 1× RIPA lysis diluted in DI water with 1 : 50 Halt™ protease & phosphatase inhibitor cocktail for 10 minutes on ice and subsequently centrifuged at 13 000g for 10 minutes for protein collection. Samples were stored at 80 °C until subsequent analysis.

Samples were processed at the Biosciences Mass Spectrometry Core Facility (https://cores.research.asu.edu/mass-spec/) at Arizona State University using the Protifi S-trap Micro Columns as per manufacturer instructions with the below modifications (using S-trap Ultra High Recovery Protocol). Samples were quantified for protein concentration using the EZQ protein quantification kit (ThermoFisher #R33200). 10 μg of total protein was taken from each sample for processing and solubilized in equal volumes of 10% SDS/100 mM TEAB (pH 7.55). A final concentration of 50 mM dithiothreitol (Sigma-Aldrich #43816-10ML) was added and vortexed with the sample, then incubated for 10 minutes at 95 °C. Proteins were alkylated with a 40 mM final concentration of freshly prepared iodoacetamide (Thermo Scientific #A39271) and incubated at room temperature for 30 minutes in the dark. Samples were then acidified by adding 55% phosphoric acid to a final concentration of ∼5% phosphoric acid and mixed thoroughly. 2.0 μg of sequencing grade modified trypsin (Promega #V5111) was immediately added to the acidified sample, mixed, combined with 7× S-trap protein binding buffer (90% methanol, 100 mM TEAB, pH 7.1), and allowed to permeate into the S-Trap Micro columns (Protifi #C02-micro-80). Columns were spun at 4000g for 1 minute and repeated until all the solution passed through. Columns were washed 4× with S-trap protein binding buffer. An additional 0.5 μg of trypsin in 25 μL of 50 mM TEAB (pH 8) was added to the top of each column, allowed to diffuse into the column matrix, and incubated for 90 minutes at 47 °C. Samples were first eluted off the S-trap columns using 50 mM TEAB and centrifuged for 1 min at 4000g. Next samples were eluted with 0.2% formic acid in water and centrifuged, and lastly with 50% acetonitrile/50% water/0.2% formic acid and centrifuged. All elutions were combined and dried down for 2 hours via speed vac and resuspended in 30 μL of 0.1% formic acid.

Liquid-chromatography tandem mass spectrometry was collected in positive mode using an Orbitrap Fusion Lumos mass spectrometer (Thermo Scientific) coupled with an UltiMate 3000 UHPLC (Thermo Scientific). One μL of peptides was fractionated using an Easy-Spray LC column (25 cm × 75 μm ID, PepMap C18, 2 μm particles, 100 Å pore size, Thermo Scientific) equipped with an upstream 300 μm × 5 mm trap column. Electrospray potential was set to 1.6 kV and the ion transfer tube temperature to 300 °C. The mass spectra were collected using the “Universal” method optimized for peptide analysis provided by ThermoScientific. Full MS scans (375–1500 m/z range) were acquired in profile mode with the Orbitrap set to a resolution of 120 000 (at 200 m/z), cycle time set to 3 seconds, and mass range set to “Normal”. The RF lens was set to 30% and the AGC was set to “Standard”. Maximum ion accumulation time was set to “Auto”. Monoisotopic peak determination (MIPS) was set to “peptide” and included charge states 2–7. Dynamic exclusion was set to 60 s with a mass tolerance of 10 ppm, and the intensity threshold set to 5.0e3. MS/MS spectra were acquired in a centroid mode using a quadrupole isolation window was set to 1.6 (m/z). Collision-induced fragmentation (CID) energy was set to 35% with an activation time of 10 milliseconds. Peptides were eluted during a 120-minute gradient at a flow rate of 0.250 μL min⁻¹ containing 2–80% acetonitrile/water as follows: 0–3 minutes at 2%, 3–30 minutes 2–15%, 30–90 minutes at 15–30%, 90–110 minutes at 30–35%, 110–120 minutes at 35–90%.

Raw spectral files were imported into Proteome Discoverer v2.5 using standard processing and consensus methods as provided by ThermoScientific. A minimum peptide length was set to 6aa and up to 2 missed cleavage sites were allowed. Sequest HT was used to identify peptide spectral masses (PSMs) and a fixed-value PSM validation method was employed. Parameters were set as follows: database used set to Uniprot Homo sapiens (Tax ID 9606), precursor mass tolerance set to 20 ppm, and fragment mass tolerance 0.5 Da, static modifications used were carbamidomethyl on cysteines (+57.021 Da).

2.11. Statistics and analysis

Background-corrected metabolic activity, protein secretions, and normalized abundance of individual proteins of interest were analyzed and visualized using GraphPad Prism 9. Ordinary one-way ANOVA with multiple comparison tests was used to show statistical significance. Data shown are individual replicates with mean and SEM for error bars.

Normalized abundance values for each sample were used as a measure of relative expression for each protein. Principal components analysis with all quantified proteins was performed using the prcomp function in R (version 4.2.1 base package stats). Markers of placental cell types as determined by published single-cell studies were examined in our data set.³ ANOVA (analysis of variance) in R (version 4.2.1) was used to assess which proteins had abundance that differed significantly between hydrogel/matrix conditions. Proteins with ANOVA p-value of less than 0.05 after Benjamini–Hochberg multiple hypothesis correction were clustered by correlation and visualized using pheatmap (R package pheatmap v 1.0.12). Proteins assigned to 3 main clusters which matched with high expression in one of the three growth conditions were extracted using the R cutree function (k = 3). Proteins in each cluster were analyzed for pathway enrichment using Metascape.⁴⁸ Significantly enriched pathways were summarized according to shared hits and specific pathways were chosen for discussion.

3. Results and discussion

3.1. Synthetic hydrogel library design for placental spheroid generation in 3D culture

We first generated a synthetic hydrogel library (Fig. 1A) consisting of PEG functionalized with adhesive ligands representative of fibronectin, collagen, or laminin peptide sequences and crosslinked with either MMP-sensitive degradable peptides or nondegradable synthetic linkers. We characterized the sensitivity of these matrices to degradation by cell-released proteases by evaluating their relative degradation rates in collagenase type A, as measured by the change in hydrogel area over time (Fig. 1B and C). Nondegradable PEG (5% w/v) hydrogel crosslinkers DTT and PEG-DT were compared to degradable peptide crosslinkers VPM, GDQ, and GPQ-W (Table S1, ESI†) which possess varying protease sensitivity. We found that DTT, PEG-DT, and GPQ-W crosslinked hydrogels exhibited little change in area over 8 hours (Fig. 1B and C). By contrast, GDQ, Matrigel, and VPM crosslinked hydrogels were completely degraded within 2, 4, and 8 hours, respectively (Fig. 1C), with no significant differences compared to Matrigel at those time points, while the hydrogel areas of the remaining conditions (DTT, PEG-DT, DTT-VPM, GPQ-W) were significantly higher than Matrigel at each time point. Hydrogels crosslinked with a mixture of VPM and DTT exhibited an increase in size over 8 hours, demonstrating hydrogel swelling as the matrix network is partially degraded. These data demonstrate the generation of a library of synthetic hydrogels with a range of protease sensitivities, including conditions (GDQ, VPM) comparable to the degradation profile of Matrigel, a common naturally derived matrix used for 3D in vitro cell culture.

3.2. Synthetic hydrogel degradation characteristics modulate trophoblast viability and proliferation

We next evaluated the influence of hydrogel degradability on encapsulated trophoblast growth and morphology over 7 days in culture (Fig. 2 and S1, S2A–C, ESI†). We assessed the viability and metabolic activity of trophoblast cell lines JEG-3, JAR, and BeWo^18,49 within our library of degradable and nondegradable synthetic hydrogels, as well as common natural degradable and nondegradable 3D hydrogel matrices. We evaluated synthetic PEG hydrogels containing fibronectin-derived adhesion ligand RGD and crosslinked with degradable (VPM, GDQ, GPQ-W) and/or nondegradable (DTT, PEG-DT) linkers and compared our synthetic platform against naturally derived hydrogel controls: protease-degradable Matrigel and human placenta-derived collagen IV and non-protease-degradable alginate and agarose. Confocal imaging on day 7 of culture demonstrated a stark difference in trophoblast morphology and viability between degradable and nondegradable hydrogels (Fig. 2A), where large cell clusters (∼100 μm diameter) were evident in degradable PEG hydrogels for all three cell lines and a large, uniform cell growth area was evident in degradable natural matrices collagen and Matrigel. By contrast, limited expansion of cells was observed in nondegradable PEG, agarose, and alginate matrices, except for JAR-seeded alginate hydrogels, where large clusters were observed. Cell death was evident in nondegradable-crosslinked hydrogels by day 1 for all cell lines, while cells in Matrigel and collagen densely and rapidly populated the gel area (Fig. S1, ESI†).

Fig. 2 Varying hydrogel degradability and composition alters trophoblast viability and growth. (A) Cell viability and morphology through z-maximum projected confocal microscopy images (green – live, magenta – dead) in degradable and nondegradable synthetic hydrogels with RGD adhesion ligand or Matrigel, collagen IV, agarose, and alginate controls on day 7 and corresponding viability (B–D) and size analysis (E–G) on JEG-3 (B and E), JAR (C and F), and BeWo (D and G) trophoblast-like cell lines (n = 5–12). scale bar = 200 μm. Cell metabolic activity via AlamarBlue assay on day 7 for JEG-3 (H), JAR (I), and BeWo (J) cell lines (n = 7–16). Data shown as mean ± SEM. Data analyzed by ordinary one-way ANOVA with Tukey's (E–G) or Dunnett's (B–D and H–J) multiple comparisons test (B–D and H–J compared to Matrigel only); * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

Trophoblast viability was highest in degradable hydrogels over 7 days, with degradable peptide-crosslinked synthetic hydrogels preserving the most viability in all cell lines (JEG-3: 84 ± 4%, JAR: 94 ± 1%, BeWo: 88 ± 3%), followed by naturally-derived matrices (JEG-3: 69 ± 4%, JAR: 81 ± 4%, BeWo: 63 ± 4%), and nondegradable synthetic matrices (JEG-3: 33 ± 9%, JAR: 48 ± 5%, BeWo: 34 ± 8%) (Fig. 2B–D). JEG-3 and JAR cells cultured in Matrigel had higher viability than when cultured in hydrogels crosslinked with nondegradable PEG-DT (JEG-3: p < 0.01, JAR: p < 0.0001) and DTT (JAR: p < 0.01) (Fig. 2B and C). BeWo cells cultured in hydrogels crosslinked with degradable VPM and GDQ and partially degradable DTT-VPM had significantly higher viability than Matrigel-cultured BeWo cells (DTT-VPM: p < 0.01, VPM: p < 0.001, GDQ: p < 0.01) (Fig. 2D).

Spheroid-like cell clusters formed and expanded in area out to day 7 for all cell lines in the degradable synthetic hydrogels, with significantly larger sizes of JEG-3 and BeWo cell clusters observed compared to nondegradable crosslinkers, while JAR cell clusters were significantly larger in degradable VPM versus nondegradable DTT hydrogels (Fig. 2A and E–G). Hydrogels made with an equimolar degradable and nondegradable crosslinker (DTT-VPM) resulted in JEG and JAR cell clusters of intermediate area between degradable and nondegradable synthetic hydrogels, whereas BeWo cell clusters grown in this condition were significantly larger than BeWo clusters cultured in nondegradable controls (p < 0.01) (Fig. 2E–G). JAR cells formed the largest overall clusters in naturally derived alginate, with significantly larger clusters than in PEG-DTT and agarose hydrogels (Fig. 2F). Consistent with these observations, the degree of circularity of live cells in nondegradable synthetic hydrogels was higher than in degradable synthetic hydrogels, indicating less cell expansion and interaction with the surrounding matrix (Fig. S2A–C, ESI†). Additionally, the feret diameter of cells in degradable hydrogels trended greater than nondegradable hydrogels for all cell types, consistent with overall area measurements (Fig. S2D–F, ESI†).

The metabolic activity of all cell lines was higher in degradable PEG hydrogels compared to nondegradable PEG hydrogels and significantly higher than the Matrigel control (JEG-3: VPM, GDQ, and GPQ-W p < 0.0001; JAR: VPM and GDQ p < 0.0001, GPQ-W p < 0.01; BeWo: VPM and GPQ-W p < 0.0001, GDQ p < 0.01) for all cell lines on day 7 (Fig. 2H–J and S3A–C, ESI†). Matrigel produced significantly higher cell metabolic activity compared to agarose and alginate with JEG-3 (p < 0.01 and p < 0.0001, respectively) (Fig. 2H), JAR (p < 0.01 and p < 0.05, respectively) (Fig. 2I), and BeWo (p < 0.0001, both) (Fig. 2J) cells. Metabolic activity for collagen IV hydrogels was not significantly different from Matrigel-cultured JEG-3 and JAR cells but significantly lower for the BeWo cells (p < 0.0001). Interestingly, JAR metabolic activity was significantly higher in alginate than in Matrigel controls, which corresponded to large cluster sizes and high viability (Fig. 2A, C, F, and I). 2D cultured cells exhibited significantly higher metabolic activity than degradable PEG hydrogels for all cell lines (JEG-3: VPM p < 0.0001, GDQ p < 0.001, GPQ-W p < 0.001; JAR: GDQ p < 0.001, GPQ-W p < 0.0001; BeWo: VPM, GDQ, GPQ-W p < 0.0001), except for JAR cells cultured in VPM crosslinked PEG gels. Overall, synthetic PEG degradable hydrogels consistently resulted in higher viability and metabolic activity compared to nondegradable synthetic and natural gels, comparable to 2D controls where cells have less growth restriction than 3D hydrogel conditions.

Trophoblast cell lines generally grew best in degradable synthetic hydrogel matrices as seen by increased viability and metabolic activity (Fig. 2), possibly due to their ability to grow in clusters (Fig. 2A–D) and remodel their environment. Matrigel is the most common matrix used to generate placental organoids/spheroids in vitro,^28,29,50,51 and several groups have successfully generated placental organoids using this platform. In our system, Matrigel resulted in rapid cell proliferation out to day 3 when cells within the hydrogel matrix became confluent and significantly degraded the ECM, resulting in a reduction in metabolic activity and proliferation observed on day 7 (Fig. S1 and S3, ESI†). This may be due to differences in seeded cell number and a rapid degradation of the Matrigel matrix, as well as the plethora of soluble signals and growth factors present in Matrigel, which may promote rapid short-term cell growth.

3.3. ECM-derived adhesive ligand presentation in synthetic hydrogels influence placental spheroid morphology and proliferation

We next explored whether modifying the adhesion ligand sequence tethered within our synthetic PEG matrices would influence encapsulated trophoblast cell viability, morphology, and function (Fig. 3 and S3D–F, S4, S5, ESI†). As the degradable crosslinker VPM produced consistent high viability, cluster size, and metabolic activity amongst the tested crosslinkers for the 3 cell lines (Fig. 2), VPM was used for testing adhesions ligands. We modified degradable (VPM-crosslinked) PEG hydrogels with adhesive ligand peptide sequences derived from several ECM components found in the placenta, such as the ubiquitous motif RGD (originally identified in fibronectin, but present across several ECM components),⁵² collagen I and II-specific motif GFOGER,⁵³ and laminin-specific motifs YIGSR (β chain)⁵⁴ and IKVAV (α chain).⁵⁵ Of note, the native placenta exhibits primarily collagens I, III, IV, and VI, laminin, and fibronectin.^23,56 Cytotrophoblasts present integrin α6β4, a laminin-binding integrin motif, on their surface throughout pregnancy,^57,58 which has been suggested to play an important role in stabilizing trophoblast adhesion.⁶ Consequently, we hypothesized that trophoblast-like cell lines may prefer adhesive ligands sourced from laminin ECM. A scrambled non-adherent fibronectin motif (RDG) and PEG hydrogels containing no adhesive ligand (“none”) were used as control groups.

Fig. 3 Adhesion ligand presentation in degradable synthetic hydrogels influences placental spheroid size and metabolic activity. (A) Cell viability and morphology through z-maximum projected confocal microscopy images (green – live, magenta – dead) in PEG-based hydrogels crosslinked with degradable VPM on day 7 with corresponding viability analysis (B–D) and live cell size analysis (E–G) on JEG-3 (B and E), JAR (C and F), and BeWo (D and G) trophoblast-like cell lines (n = 6–12). Scale bar = 200 μm. (H–J) Cell metabolic activity via AlamarBlue assay on day 7 for JEG-3 (H), JAR (I), and BeWo (J) trophoblast-like cell lines (n = 12). Data shown as mean ± SEM. Analyzed by ordinary one-way ANOVA with Tukey's multiple comparisons test; ns = not significant, * p < 0.05, ** p < 0.01.

Imaging across 7 days revealed increases in cell cluster size over time across all groups, including controls (Fig. S4, ESI†). Varying adhesion ligand or excluding it all together did not have a significant impact on viability for any cell line (Fig. 3A–D); however, adhesive ligand inclusion impacted cluster area on day 7 after encapsulation (Fig. 3E–G) with RGD producing significantly larger cell cluster areas compared to GFOGER (p < 0.05) and YIGSR (p < 0.01) in JEG-3 spheroids (Fig. 3E), IKVAV (p < 0.05) in JAR spheroids (Fig. 3F), and YIGSR (p < 0.05) in BeWo spheroids (Fig. 3G). Consistent with these observations, circularity of RGD clusters was lower than other adhesive ligand groups, and ferret diameter was larger for RGD adhesive ligand with the only the BeWo cell line (Fig. S5, ESI†). Interestingly, despite the strong presence of laminin in the placenta, the inclusion of laminin-derived adhesive ligands led to reduced cluster size in JEG-3 (YIGSR; Fig. 3E), JAR (IKVAV; Fig. 3F), and BeWo cultures (YIGSR; Fig. 3G).

Metabolic activity was similar across adhesion ligands and control groups (Fig. 3H–J and S3D–F, ESI†), with JAR spheroids exhibiting greater overall average activity (∼4500 RFU, Fig. 3I) relative to JEG-3 (∼3000 RFU, Fig. 3H) and BeWo (∼2500 RFU, Fig. 3J). JAR spheroids show no significant differences in metabolic activity with varying adhesion ligands or controls (Fig. 3I), while metabolic activity in JEG-3 spheroids was significantly higher in RGD than YIGSR (p < 0.05) (Fig. 3H). BeWo metabolic activity was significantly higher in response to RGD than RDG (p < 0.01) and IKVAV (p < 0.05) ligands with RDG cultured spheroids displaying significantly lower metabolic activity than the no adhesion ligand control (p < 0.05) (Fig. 3J). Overall, varying the adhesion ligands had minor effects on placental spheroid viability, cluster size, circularity, and metabolic activity. We hypothesize that this may be due to choriocarcinoma cells remodeling and laying their own ECM in degradable environments; however, adhesion ligands may be more influential in future studies with primary cells that may be more sensitive to environmental ECM signaling.

3.4. Composition of 3D culture matrix modulates trophoblast protein expression

We next investigated how 3D culture conditions and bioactive signaling in a 3D environment altered biological processes in trophoblasts. Shotgun proteomics was used to assay global protein expression in JAR cells cultured for 7 days in standard 2D culture conditions compared against a highly defined 3D hydrogel condition (degradable synthetic hydrogel PEG-VPM-RGD) and Matrigel (Fig. 4 and S6, ESI†). As these conditions showed significant differences in metabolic activity in JAR cells (Fig. 2I), we assayed protein expression to determine possible molecular mechanisms that underly differences in cell behavior when grown in 2D or 3D conditions. 2998 proteins were quantified across the three culture conditions using high-throughput mass spectrometry (Table S2, ESI†). Principal component analysis demonstrates that trophoblasts grown in each of the three culture conditions have a distinct protein expression profile (Fig. 4A), with synthetic hydrogels and 2D conditions clustering separately from Matrigel on the first dimension, and the three groups clustering separately on the second dimension, indicating that Matrigel exhibits the most distinct profile of the compared groups. Hierarchical clustering of proteins differentially abundant across culture conditions shows that protein expression in cells grown in synthetic hydrogels is more comparable to cells grown in 2D culture, while cells grown in Matrigel have more distinct protein expression patterns (Fig. 4B). Overall, this indicates greater similarity in protein expression between 2D and synthetic hydrogel culture conditions with Matrigel protein expression varying significantly from the other two conditions.

Fig. 4 Proteomic analysis of JAR cells cultured in 2D, Matrigel, and PEG-RGD-VPM. (A) Principal component analysis of global protein abundance shows distinct separation on culture medium after 7 days of culture. (B) Hierarchical clustering of proteins with significantly different abundance (ANOVA adjusted p-value <0.05). Summary of significantly enriched pathways in proteins in each of the major expression clusters are listed. (c) Protein abundance of markers of placental cell types (syncytiotrophoblasts (ST), cytotrophoblasts (CT), extravillous trophoblasts (EVT)), and common to all trophoblast types (PAN-TB), from placenta cells observed at 8 weeks (8w) or term in RNA-seq analysis. Abundance of PAN-TB maker (D) KRT7, CT markers (e) CDH1 and (f) ITGA6, EVT marker (G) ITGA5, and (H) tight junction protein 2 (TJP2) (n = 3). Data shown as mean ± SEM. D–H Analyzed by ordinary one-way ANOVA with Tukey's multiple comparisons test; * p < 0.05, ** p < 0.01, *** p < 0.001.

We next investigated overall pathway enrichment to identify patterns in protein expression changes across culture conditions (Fig. 4B). 1512 proteins showed differential protein abundance across the three culture conditions (ANOVA adjusted p < 0.05). Proteins that have higher abundance in synthetic hydrogel and 2D culture are associated with pathways involved in translation, mRNA metabolism, cell cycle, DNA replication, and mitochondrial activity, consistent with increased cell proliferation observed in these conditions (Fig. 2I and 4B; full list of enriched pathways provided in Table S3, ESI†). Proteins with higher expression in Matrigel are enriched for extracellular matrix reorganization, as well as many other proteins that are involved in cell adhesion (Fig. 4B), such as laminin, fibulin, and vitronectin (Fig. S6A, ESI†).

We next sought to evaluate how culture conditions affect trophoblast phenotypic protein expression and polarity. We evaluated normalized protein abundance of markers specific to primary human CT, ST, and EVT phenotypes broadly, as well as phenotypic markers identified in 8-week gestational age primary trophoblasts in an RNA-seq dataset³ (Fig. 4C–G and S6, ESI†). Most trophoblast phenotype markers had elevated expression in the 2D and 3D synthetic hydrogel conditions and lower expression in the Matrigel condition (Fig. 4C). Normalized abundance of pan-trophoblast marker KRT7 (Fig. 4D), CT marker ITGA6 (Fig. 4F and S6B, ESI†), and tight junction protein 2 (TJP2) (Fig. 4H) were significantly higher in 2D cultured cells compared to synthetic hydrogel (p < 0.01, p < 0.001, p < 0.05, respectively) and Matrigel (p < 0.05, p < 0.001, p < 0.001, respectively) conditions, with TJP2 expression in synthetic hydrogel also higher compared to Matrigel (p < 0.05). CT marker CDH1 (Fig. 4E) and EVT marker ITGA5 (Fig. 4G and S6B, ESI†) were significantly higher in Matrigel relative to 2D (p < 0.01, p < 0.01, respectively), with ITGA5 expression in Matrigel also higher compared to the synthetic hydrogel (p < 0.001), but all other CT and ST markers showed increased abundance in synthetic hydrogel and 2D cultured conditions (Fig. 4C). Apoptosis proteins such as programmed cell death protein-5 (PD-5) and PD-6 were significantly higher in Matrigel relative to 2D controls (p < 0.0001 and p < 0.05, respectively), while PD-2 had significantly higher expression in 2D than Matrigel (p < 0.05; Fig. S6C, ESI†). Other proteins of interest, such as MMP14 (Fig. S6D, ESI†) which is involved in trophoblast invasion, TJP1 gene (Fig. S6E, ESI†) which encodes for ZO-1 that is downregulated in 3D culture,^59,60 and TJP3 (Fig. S6F, ESI†), showed no significant differences between culture groups. Overall, these data demonstrate that 2D and synthetic hydrogel cultured cells support a protein expression profile more consistent with characteristic CT and ST phenotypes, whereas Matrigel results in reduced proliferation at this timepoint and an EVT-like phenotype. These data illustrate how culture conditions can modulate trophoblast cellular phenotype.

3.5. Synthetic hydrogel degradation characteristics influence placental spheroid phenotype and function

As proteomics analysis indicated that 3D culture conditions induced significant alterations to trophoblast phenotype, we next sought to evaluate how the degradability of 3D culture matrices influence placental spheroid phenotype via trophoblast marker expression and functional secretion (Fig. 5 and S7–S15, ESI†). Trophoblast marker expression was assessed to evaluate changes in expression of CT (ITGA6) and ST (synd-1) specific markers via cytohistochemistry, and EVT (HLA-G) specific marker via western blotting for JEG, JAR, and BeWo cell lines. JEG-3 cells (Fig. S7A and S11, ESI†) express all three of these markers, while JAR cells (Fig. 5A and S11, ESI†) are known to be HLA-G negative^18,61 and BeWo cells (Fig. S7B and S11, ESI†) have conflicting reports of HLA-G expression.^18,61,62 By western blot analysis, we show JEG-3 cells to be HLA-G positive in PEG-VPM-RGD, Matrigel, and 2D culture conditions on day 7, while JAR and BeWo remain HLA-G negative in all conditions (Fig. S11, ESI†). Decreased expression of the ITGA6 and synd-1 surface markers in the nondegradable PEG-based hydrogels is likely due to decreased viability and proliferation at day 7 (Fig. 2B–D and H–J), and robust expression of these two markers were observed in degradable hydrogel groups across all three cell types (Fig. 5A and S7, S10, ESI†).

Fig. 5 Hydrogel composition and degradability influences trophoblast phenotypic secretion. (A) JAR cells stained with ITGA6 (yellow) and synd-1 (syndecan-1, magenta) with DAPI (blue) nuclear stain after culture in PEG-RGD hydrogels with varying crosslinkers or hydrogel controls for 7 days (n = 3–9). Scale bar = 100 μm. Secretion of (B and C) MMP2 and (D–F) hCGβ from (d) JEG-3, (B and E) JAR, and (C and F) BeWo cells on day 7 (n = 8). Data shown as mean ± SEM. Analyzed by ordinary one-way ANOVA with Dunnett's multiple comparisons test to Matrigel; * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

To further investigate the influence of culture conditions on trophoblast phenotype and function, we examined the secretion levels of human chorionic gonadotropin beta (hCGβ), a hormone associated with the ST phenotype, and matrix metalloproteinase-2 (MMP2), a necessary factor in normal placenta remodeling and a protease associated with the EVT phenotype⁶³ (Fig. 5B–F). Cell lines demonstrated similar secretion profiles over the first 3 days of culture in varying matrices (Fig. S9A–C and G, H, ESI†), with an increase of total hCGβ and MMP2 secretion in degradable PEG hydrogels compared to nondegradable PEG hydrogels on day 7 across all three cell types, most likely due to the decreased viability and proliferation of cells in nondegradable environments (Fig. 2). Surprisingly, EVT-like JEG-3 spheroids did not secrete detectable levels of MMP2 (0.6 ng mL⁻¹), whereas JAR and BeWo spheroids exhibited comparable levels of MMP2 secretion (Fig. 5B and C), with secretion significantly impaired in nondegradable synthetic hydrogel environments. JAR and BeWo cells cultured in degradable PEG-based hydrogels secreted significantly higher levels of MMP2 than Matrigel cultured cells (JAR: VPM p < 0.0001; BeWo: VPM p < 0.01, GDQ and GPQ-W p < 0.05). JAR and BeWo cells cultured in degradable PEG-based hydrogels also secreted significantly more total hCGβ than Matrigel (p < 0.0001) (Fig. 5E and F). Notably, in 2D culture, BeWo cells tend to exhibit the most ST-like phenotype of the three cell lines,¹⁸ as evidenced by 10- and 20-fold higher hCGβ secretion in BeWo cells (Fig. 5F) relative to JAR (Fig. 5E) and JEG-3 lines (Fig. 5D), respectively. Secretion of hCGβ in degradable hydrogels was higher than in 2D-cultured cells, while the total secretion of MMP2 was comparable between degradable hydrogels and 2D-cultured cells. Generally, MMP2 secretion, which corresponds to an EVT phenotype, correlated to observations of increased EVT-like phenotype in 2D culture conditions in proteomics analysis relative to synthetic hydrogel or Matrigel conditions (Fig. 4C and G). Additionally, hCGβ secretion, indicative of ST-like phenotype, was generally higher in degradable synthetic hydrogel and 2D relative to Matrigel conditions in JAR and BeWo cells, a phenotype also observed in proteomics analysis on JAR trophoblasts (Fig. 4C). Surprisingly, the secretion of both MMP2 and hCGβ was generally high in alginate culture groups in JAR and BeWo cells, despite a lack of physiological cues to support phenotypic differentiation. This observation warrants further investigation but may be due to the relatively greater stiffness of alginate (∼5000 Pa)⁶⁴ over PEG (1000 Pa)⁶⁵ and Matrigel (44 Pa),⁶⁶ which greater approximates the native placental mechanical environment (∼1000–5000 Pa).⁹

Previous work generating placental spheroids using trophoblast stem cells seeded in Matrigel has consistently noted inverted architectures, where ST cells cluster in the center of organoids surrounded by CT while EVT line the periphery.^28,29 In our hands, trophoblast cell lines seeded in Matrigel did not form organoid-like clusters, but rather uniform sheets of cells without any notable organization to CT, ST, and EVT phenotypic staining, which may be impacted by cell seeding density. Functionally, JAR and BeWo trophoblast cell lines in Matrigel secreted generally lower levels of hCGβ than those cultured in 2D or synthetic hydrogels. Trophoblast cell lines seeded in synthetic degradable hydrogels demonstrated localization of ST marker synd-1 at the periphery of spheroids (Fig. 5A and S7, S14, ESI†), indicating an organization homologous to the native placenta, whereas spheroid structures in nondegradable synthetic (DTT, PEG-DT) and natural (alginate, agarose) hydrogels display a less organized or even inverted phenotype. This correlated to overall higher hCGβ secretion in synthetic degradable hydrogels than in Matrigel or 2D controls (Fig. 5E and F), suggesting that culture environments that support organization typical to the native placenta can enhance functional outcomes. Additionally, these functional outcomes correlate to observations in our proteomic analysis, which identified upregulation of ST markers in 2D and degradable synthetic hydrogel groups and downregulation of ST proteins in Matrigel-cultured JAR cells (Fig. 4C). Interestingly, proteomics analysis identified CGB3 expression only in the degradable synthetic hydrogel group and not the Matrigel or 2D control groups, which further supports that the synthetic hydrogel composition promotes a functional ST phenotype (Table S4, ESI†). Conversely, while alginate promoted spheroid generation comparable to degradable synthetic hydrogel environments in JAR cells, these spheroids exhibited stronger synd-1 staining in the center of the spheroid (Fig. 5A). Functionally, alginate-generated JAR spheroids demonstrated elevated MMP2 secretion, indicative of an EVT-like skewed phenotype relative to Matrigel and synthetic degradable hydrogel controls, further supporting the premise that spheroid architecture may influence function, and that this architecture may be controlled by physical cues in the 3D culture environment.

3.6. Synthetic hydrogel adhesion ligand signals have limited influence on placental spheroid phenotype

Finally, we evaluated the influence of varying adhesion ligands within synthetic hydrogels on placental spheroid phenotype (Fig. 6 and S9, S13–16, ESI†) via CT, ST, and EVT-associated surface marker expression and secreted factors hCGβ and MMP2. Variation of adhesion ligand within synthetic hydrogels did not noticeably alter the expression of ITGA6 and synd-1 markers across JEG-3, JAR, and BeWo-derived spheroids (Fig. 6A, C, F and S14, ESI†) and did not alter hCGβ secretion in JEG-3 spheroids (Fig. 6B) even when normalized to the metabolic activity of each group (Fig. S13A, ESI†), which could account for differences due to total cell number on day 7 of culture. In JAR spheroids, no adhesion ligand (“none”) produced significantly more hCGβ secretion cells compared to the incorporation of GFOGER (p < 0.01), YIGSR (p < 0.05), and IKVAV (p < 0.0001) (Fig. 6D), and IKVAV presentation resulted in significantly less total MMP2 than RGD (p < 0.01), RDG (p < 0.0001), GFOGER (p < 0.01), YIGSR (p < 0.001), and no adhesion ligand (p < 0.001) (Fig. 6E). Normalization of secretion to metabolic activity yields similar results where IKVAV produces significantly less hCGβ secretion in JAR cells than RGD (p < 0.01), RDG (p < 0.001), YIGSR (p < 0.05), and no adhesion ligand (p < 0.0001) (Fig. S13B, ESI†), and IKVAV produces significantly less MMP2 secretion in JAR spheroids than RGD (p < 0.001), RDG (p < 0.0001), GFOGER (p < 0.01), YIGSR (p < 0.001), and no adhesion ligand (p < 0.01) (Fig. S13C, ESI†). Total secretion of hCGβ and MMP2 from BeWo spheroids was not significant between groups (Fig. 6G and H); however, normalization to metabolic activity produced significant differences, where YIGSR produced the highest average hCGβ secretion relative to RGD (p < 0.0001), GFOGER (p < 0.001), and no adhesion ligand (p < 0.01) (Fig. S13D, ESI†), and YIGSR produced the highest average MMP2 secretion relative to RGD (p < 0.0001), GFOGER (p < 0.0001), and no adhesion ligand (p < 0.0001) (Fig. S13E, ESI†).

Fig. 6 Synthetic hydrogel adhesion ligand functionalization influences placental spheroid phenotype. (A) JEG-3, (C) JAR, (F) BeWo cells stained with ITGA6 (yellow) and synd-1 (sydecan-1, magenta) with DAPI (blue) nuclear stain after culture in degradable PEG-VPM hydrogels with varying adhesion ligands for 7 days (n = 3). Scale bar = 100 μm. Secretion of (B, D and G) hCGβ and (E and H) MMP2 from (B) JEG-3, (D and E) JAR, and (G and H) BeWo cells in varying adhesion ligands in PEG-based hydrogels on day 7 (n = 8). Data shown as mean ± SEM. Analyzed by ordinary one-way ANOVA with Tukey's multiple comparisons test; ns = not significant, * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

As adhesion ligand variation resulted in limited effects on the growth and metabolic activity of placental spheroids, it is unsurprising that we observed limited significant effects on protein expression and secretion between adhesion ligand groups. However, one interesting observation is that laminin adhesive ligand IKVAV, but not YIGSR, lead to reduced hCGβ and MMP2 secretion in JAR spheroids, suggesting that this ligand is not supportive of ST or EVT phenotypes. This is surprising given that laminin represents a major component of the placenta ECM, along with collagens I, III, IV, and VI.²³ Our proteomic analysis of JAR cells demonstrated the presence of integrin α1, α5, α6, and αV, and β1, β4, and β5 (Fig. S6B and Table S2, ESI†). Given that integrin α1, α6, β1, and β4 are associated with laminin binding, and YIGSR is known to bind integrin α4 β1,⁵⁴ it is interesting that laminin-derived peptides resulted in less hCGβ and MMP2 secretion in JAR spheroids, suggesting that specificity of the ECM ligands presented to placenta spheroids can influence functional outcomes. While adhesive ligand variation demonstrated limited effects in placental spheroid function with trophoblast-like cell lines, we expect that these effects may be more pronounced with primary cytotrophoblast cells, as primary cells can be more sensitive to environmental cues than transformed cell lines.⁶⁷

JEG-3, JAR, and BeWo cells lines originate from human choriocarcinoma and while they are often used to model invasion, migration, and syncytialization of the human placenta, these cells are unlikely to recapitulate the behavior of healthy tissue. The synthetic hydrogel 3D culture model described in this work can be applied to more physiologically relevant cells to study placental development and microenvironment, such as induced pluripotent stem cell-derived trophoblasts^29,68 and primary trophoblast stem cells.^69–71 Cell processes such as invasion, migration, and signaling rely heavily on dynamic protein expression, interactions with ECM, and integrin expression, and our data with JEG-3, JAR, and BeWo demonstrate that a 3D synthetic hydrogel can provide these necessary cues. We expect the 3D environment provided by hydrogel matrices to better support primary trophoblast phenotypes’ physiological functions, such as the ability of CT to proliferate and differentiate and ST to secrete hCGβ.

4. Conclusions

Here, we employed a highly defined, synthetic poly(ethylene glycol)-based hydrogel system with tunable degradability and containing extracellular matrix-derived adhesive ligands native to the placenta microenvironment to investigate whether a 3D culture environment influences trophoblast spheroid growth and phenotype. We evaluated the capacity of this material library to support the viability, function, and phenotypic protein expression of three trophoblast cell lines with features of CT, ST, and EVT phenotypes over 7 days and found that degradable synthetic hydrogels support the greatest degree of placental spheroid viability, proliferation, and function relative to gold standard Matrigel controls. Finally, we show that 3D culture conditions modulate trophoblast cell functional phenotype as measured by proteomics analysis and functional secretion assays. Overall, we demonstrated the suitability of biomimetic synthetic hydrogel matrices as a 3D culture system for trophoblast-like cells and show that hydrogel composition influences trophoblast spheroid function and phenotype. Future studies will evaluate primary human placental trophoblasts within our synthetic hydrogel system, which may enable the generation of placental organoids that can be used to study trophoblast-matrix interactions and trophoblast cell–cell interactions. This hydrogel system may enable the study of human placental physiology, function, and development in a superior manner than traditional 2D culture systems.

Data availability

The data that support the findings of this study are available in the ESI† and from the corresponding author upon request.

Author contributions

E. M. S. and J. D. W. conceptualized and planned the experiments. E. M. S., S. R. B., and S. C. H. performed the experiments. E. M. S., S. R. B., S. B. P., and J. D. W. analyzed the data and composed the figures. All authors contributed to the writing of the manuscript.

Conflicts of interest

E. M. S., S. R. B., S. B. P., and S. C. H. have no conflicts to declare. J. D. W. is the cofounder of, and holds equity in, ImmunoShield Therapeutics.

Acknowledgements

We acknowledge funding through the Juvenile Diabetes Research Foundation Innovator Award (1-INO-2020-915-A-N), the Arizona Biomedical Research Center New Investigator Award, and the National Institutes of Health Director's New Innovator Award (DP2AI169476). We thank the Regenerative Medicine and Bioimaging Facility at ASU for use of the Leica SP8 confocal microscope system, which was acquired by the NIH SIG Award 1 S10 OD023691-01, the Bioscience Mass Spectrometry Core Facility at ASU for running the mass spectrometry samples, and the Kuei-Chun Wang Lab at ASU for use of their Analytik JenaTM UVP ChemStudio for the western blot imaging.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: https://doi.org/10.1039/d3bm01393f

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