Templated Macroporous Polyethylene Glycol Hydrogels for Spheroid

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Templated Macroporous Polyethylene Glycol Hydrogels for Spheroid and Aggregate Cell Culture Mozhdeh Imaninezhad, Lindsay Hill, Grant Kolar, Kyle Vogt, and Silviya Zustiak Bioconjugate Chem., Just Accepted Manuscript • DOI: 10.1021/acs.bioconjchem.8b00596 • Publication Date (Web): 18 Dec 2018 Downloaded from http://pubs.acs.org on December 19, 2018

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Bioconjugate Chemistry

Templated Macroporous Polyethylene Glycol Hydrogels for Spheroid and Aggregate Cell Culture 1Mozhdeh

Imaninezhad, 1Lindsay Hill, 2Grant Kolar, 1Kyle Vogt, 1,*Silviya Petrova Zustiak

1Department

of Biomedical Engineering, Saint Louis University, Saint Louis, MO 63103

2Department

of Pathology, Saint Louis University, Saint Louis, MO 63104

To whom correspondence should be addressed: Silviya Petrova Zustiak Department of Biomedical Engineering Saint Louis University Saint Louis, MO, USA, 63103 Tel: 314-977-8331 Fax: 314-977-8403 E-mail: [email protected]

Key words: Macroporous hydrogel, cell encapsulation, multicellular spheroid, droplet generator, cell scaffold, degradable microspheres, microfluidics

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ABSTRACT Macroporous cell-laden hydrogels have recently gained recognition for a wide range of biomedical and bioengineering applications. There are various approaches to create porosity in hydrogels such as lyophilization or foam formation, to name a few. However, many do not allow a precise control over pore size or are not compatible with in situ cell encapsulation. Here, we developed novel templated macroporous hydrogels by encapsulating uniform degradable hydrogel microspheres produced via microfluidics into a hydrogel slab. The microspheres degraded completely leaving macropores behind. Microsphere degradation was dependent on the incubation medium, microspheres size, microsphere confinement in the hydrogel as well as cell encapsulation. Uniquely, the degradable microspheres were biocompatible and when laden with cells, the cells were deposited in the macropores upon microsphere degradation and formed multicellular aggregates. The hydrogel-encapsulated cell aggregates were used in a small drug screen to demonstrate the relevance of cell-matrix interactions for multicellular spheroid drug responsiveness. Hydrogel-grown spheroid cultures are increasingly important in applications such as in vitro tumor, hepatocellular, and neurosphere cultures and drug screening; hence, the templated cell aggregate-laden hydrogels described here would find utility in various applications.


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1. Introduction Hydrogels, due to their high water content, similarity to the native extracellular matrix (ECM), biocompatibility, and tunable mechanical, chemical and physical properties are attractive as cell scaffolds for tissue engineering, cell delivery, and drug screening applications.1 However, a major limitation of hydrogels is their nano- to micro-porosity, which could be restrictive for oxygen and nutrients diffusion, as well as cell infiltration, migration and proliferation.2, 3 Consequently, strategies have been sought after for introducing macroporosity, while capitalizing on the inherently beneficial properties of hydrogels.4 Improved porosity and pore interconnectivity has led to improved angiogenesis in vitro and in vivo,5 improved cell growth and ECM secretion for tissue engineered cartilage,2 ability to fabricate organoid cultures,6 improved scaffolds for stem cell propagation and differentiation,7, 8 and fabrication of effective wound healing matrices,9 among others. Macroporous hydrogels have been created by approaches such as phase separation,10 lyophilization,7, 11 foam formation and cryogelation.12-14 While useful, such materials are hampered by non-uniform pore sizes, toxic byproducts or leachables, and fabrication conditions that are not compatible with cell encapsulation. Cell seeding in pre-fabricated macroporous scaffolds is achieved by cell infiltration, which could result in low cell seeding efficiency, heterogeneous cell distribution and lengthy seeding periods.4 To achieve in situ macropore formation in the presence of encapsulated cells, pore formation methods should be compatible with high cell viability, while resulting in desired hydrogel physical and mechanical properties. A few recent reports have focused on developing macroporous hydrogels in the presence of cells by utilizing biocompatible gelatin microspheres as a sacrificial porogen.7, 15 Such hydrogels can be prepared at room temperature in the presence of cells and then incubated at 37oC to liquidify 3 ACS Paragon Plus Environment


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and leach out the gelatin, leaving pores behind.15 However, this approach is limited to seeding the cells in the surrounding bulk gel as the gelatin microsphere preparation techniques are not compatible with cell encapsulation. Here, we explored a similar principle of embedding degradable microspheres as porogens in a bulk hydrogel slab to create a templated scaffold. The difference in our approach is that cells were encapsulated in biocompatible and biodegradable microspheres, which were then embedded in a hydrogel slab. The cell-laden microspheres were produced with a narrow size distribution and in a wide range of sizes (120 - 480 μm pre-swelling) via a simple microfluidic technique. Upon microsphere degradation, cells were deposited directly in spherical pore openings inside the hydrogel slab enabling cells to aggregate and form aggregate spheroid cultures. These spheroid cultures would then be limited to the size of the pore created upon microsphere degradation. Hydrogel-grown spheroid and aggregate cultures are increasingly important in applications such as in vitro tumor,16 hepatocellular,15 neurosphere cultures,17 and drug screening applications.18 For example, hydrogel grown tumor spheroids have been found to exhibit more uniformly spherical morphology, reproducible drug responses and different gene expression profiles compared to liquid-grown spheroids.18 Confinement and mechanical cues from the surrounding matrix in particular have been shown to influence cellular density, motility and invasiveness in growing cell aggregates.19 Hence, the templated spheroid-laden hydrogels described here would find a broad utility. Microfluidic techniques to generate microencapsulated tumor spheroids have been described previously. For example alginate-encapsulated breast cancer spheroids were generated and trapped in a microfluidic device and subsequently used in drug screening applications.20, 21 Microfluidics has been utilized to also produce leukemia cell-loaded alginate microbeads, where


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CaCO3 nanoparticles were used to combine internal gelation with droplet formation for the production of cell-loaded monodisperse microbeads of tunable sizes.22 In yet another microfluidic flow focusing device, alginate was combined with matrigel to obtain human cervical carcinoma spheroids.23 In our approach the microfluidic device was used to fabricate cell-laden degradable polyethylene glycol (PEG) microspheres used as porogens in the developed templated hydrogel. We focus on the synthetic hydrogel PEG for both the hydrogel slab and the degradable microspheres, albeit of two different gelation chemistries. PEG is biocompatible, inert, highly swollen and allows tight control and independent tunability of mechanical, biochemical and physical properties.24, 25 To encapsulate cells in the PEG microspheres, we used Michael-type addition reaction between an acrylate and a thiol moiety due to the reaction’s mildness and selectivity.26 Furthermore, our group has identified multiple dithiol crosslinkers that can be used to control the rate of microsphere degradation from hours to days or weeks.24, 27 While here we utilized a single degradable crosslinker, such versatility would provide flexibility in applications that require microsphere degradation tuned to cell proliferation or native matrix synthesis. Finally, to demonstrate the significance of hydrogel encapsulation and subsequent cell-matrix interactions, we conducted a small drug screen with the developed aggregate-laden hydrogels.

2. Results Here, we developed templated PEG hydrogels with cell-laden macropores. This was accomplished by encapsulating hydrolytically degradable PEG hydrogel microspheres prepared from 4-arm polyethylene glycol acrylate (4-arm PEGAc) and a 2-amino butane dithiol (DTBA) crosslinker in a non-degradable PEG diacrylate (PEGDA) bulk hydrogel slab. The microspheres 5 ACS Paragon Plus Environment


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were pre-loaded with cells, which were then deposited in the pores left behind in the hydrogel slab upon microsphere degradation. 2.1. Fabrication of Hydrogel Microspheres via Microfluidics PEG hydrogel microspheres in varying sizes and with a narrow size distribution were produced by a custom T-junction microfluidic device (Figure 1). Microspheres were then used to prepare templated hydrogels as described in Figure 2. The microfluidic set-up was chosen for its simplicity and cost effectiveness. The microspheres were produced as droplets inside the microfluidic device, collected in an olive oil bath and allowed to gel. Gelation was achieved by a Michael-type addition reaction, which is base-catalyzed and strongly affected by the pH of the reaction buffer.27 Since droplet generation by microfluidics requires time, we first aimed to determine reaction buffer conditions that modulate gelation time, while being compatible with cell encapsulation. We chose to investigate piperazineethanesulfonic acid (HEPES) and triethanolamine (TEA) buffers, which are biocompatible,27 at a near neutral pH range of 6.5 to 7.4. Gelation time was measured by the inverted tube method (Figure 3A) and by rheology for one buffer condition, namely 0.3 M TEA (Figure 3B). A good agreement between the two methods was found. As expected, reaction buffer pH was inversely correlated with gelation time (Figure 3).28, 29

Interestingly, the effect of pH was more pronounced for the TEA buffer as compared to the

HEPES buffer and was also more pronounced at higher buffer strength for TEA (0.3 M vs 0.1 M). Overall, there were no significant differences in gelation time between the different buffer conditions at the highest pH of 7.4. However, at acidic pH of 6.9 and 6.5, gelation time decreased slightly but significantly with decrease in TEA buffer strength and was lower for the HEPES buffer compared to the TEA buffer of the same strength. The tested conditions resulted 6 ACS Paragon Plus Environment

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in a gelation time range of ~2 – 13 min. TEA 0.3 M at pH 7.4 with a gelation time of ~2 min was used for most subsequent experiments, primarily to ensure high cell viability. Overall, gelation time limits the number of microspheres that can be produced per run. Note that change in buffer pH resulted in change in microsphere size, where faster gelation (i.e. higher pH) was associated with larger microsphere sizes and more polydispersity (Table S1), possibly due to increase in polymer precursor solution viscosity (due to initiation of gelation) during the run.30, 31 Next, to control microsphere size, we modulated the flow rate of the PEG hydrogel precursor solution (discrete phase) and the olive oil (continuous phase) (Table 1, Figure 4 Figure S3). Microspheres were imaged and measured immediately upon fabrication (in the oil bath). Note that microspheres increased in size 20-30% upon washing and swelling in an aqueous buffer, however, the distribution as measured by percent coefficient of variance (%CV) increased only slightly (Table S2). We noted that decrease in PEG (disperse phase) flow rate resulted in smaller microsphere sizes. For example, microsphere size changed significantly from 480 ± 108 μm to 327 ± 63 μm (47% change) when PEG flow rate decreased from 60 to 40 μL/min (1 mL/min oil flow rate), and it changed from 327 ± 63 to 181 ± 34 μm (45% change), when the PEG flow rate was changed from 40 to 20 μL/min (1 mL/min oil flow rate). Overall, the change was more pronounced for higher PEG flow rates: ~2% increase in microsphere size per 1 μL/min decrease in PEG flow rate for PEG flowrates above 20 μL/min and ~1 - 1.5% increase in microsphere size per 1 μL/min decrease in PEG flow rate for PEG flow rates below 20 μL/min. Conversely, decrease in oil (continuous phase) flow rate resulted in a slight increase in microsphere size. For example, microsphere size increased from 125 ± 18 μm to 153 ± 35 μm (18% change) when oil flow rate decreased from 1 to 0.5 mL/min (10 μL/min PEG flow rate). When considering the



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ratio of flow rates - Qoil/QPEG, where Qoil stands for oil flow rate and QPEG stands for PEG flow rate, we noted a sharp initial decrease in microsphere diameter with increase in Qoil/QPEG (Figure 4). However, we noted minimal change in microsphere diameter for Qoil/QPEG above 50. Note that the same ratios produced similar size microspheres, regardless of the individual oil or PEG flow rates. For example, a Qoil/QPEG of 50, achieved through 0.5 mL/min Qoil and 10 μL/min QPEG and 1 mL/min Qoil and 20 μL/min QPEG led to 153 ± 35 μm and 181 ± 34 μm size microspheres, respectively. No clear trend was noted for the change in %CV, where %CV varied between 12 and 23 for the various combinations of PEG and oil flow rates.

2.2. Degradation of Hydrogel Microspheres as a Function of Medium, Cell Loading and Confinement Microsphere degradation was followed for both microspheres in medium directly (nonconfined) and microspheres loaded in a hydrogel slab (confined). The confined condition represents microspheres loaded in a bulk hydrogel slab as depicted in Figure 2. Figure 2D shows a representative compressed z-stack fluorescent image of microspheres embedded in a hydrogel slab, where fluorescent beads were used to aid visualization. The image shows that microspheres were randomly dispersed within the slab and were also visible in multiple planes, indicating that not all microspheres settle to the bottom of the slab during gelation. Microscopy images of microspheres incubated at 37°C and 5% CO2 in 1X phosphate buffered saline (PBS), serum medium, serum-free medium and with encapsulated cells in serum medium for 2, 24, 72, and 168 h are shown in Figure 5A (non-confined) and Figure 5B (confined). The microspheres were loaded with fluorescent beads, which helped identify the contours of the microspheres prior to degradation, but were released in the surrounding medium 8 ACS Paragon Plus Environment

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or inside the pores of the hydrogel slab upon microsphere degradation. For non-confined microspheres, degradation was noted by increase in microsphere size and the “loss of contour” due to fluorescent bead release in the medium upon degradation (Figure S2A). Microspheres have been shown to increase in size upon degradation due to random network scission, which leads to decrease in crosslink density and increase in swelling.32, 33 For confined microspheres swelling was hampered by confinement; thus, degradation was monitored by decrease in microsphere size and “loss of sphericity” as determined by volumetric microscopy images (Figure S2B). Upon hydrolysis, polymer degradation products diffused out of the templated hydrogels and the fluorescent beads, which were previously suspended in the nanoporous structure of the microsphere network, accumulated at the bottom of the macropores due to the force of gravity. The change in microsphere shape (i.e. sphericity) upon degradation is illustrated in Video S1 for a confined microsphere in PBS at 7 d, where degradation was minimal, and in Video S2 for a confined microsphere in serum medium at 7 d, where degradation was extensive. Overall, we noted that non-confined microspheres degraded faster than confined ones, which corroborated prior research.34 When non-confined, hydrogel microspheres degraded completely at 24 h in serum medium and at 168 h in PBS and serum-free medium. Microspheres degraded in 168 h in serum medium when cells were encapsulated within, indicating that cell loading slowed the degradation considerably. For confined microspheres, we noted decrease in microsphere size predominantly for microspheres in serum medium, where decrease in size was indicative of degradation as described above. To quantify degradation, we monitored microsphere size as a function of time. Again, microsphere size (as tracked by the contour of the fluorescent particles encapsulated in the microsphere) increased upon degradation for non-confined microspheres (Figure 5C) and



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decreased (due to fluorescent beads settling to the bottom of the macropores) for confined ones (Figure 5D). For non-confined microspheres we noted similar size increase in PBS (22%), serum-free medium (24%), and serum medium with loaded cells (19%), as measured by the difference in microsphere size between the first time point (2 h) and the last time point prior to complete degradation (72 h) (Figure 5C). Overall, in non-confined conditions, the microspheres exhibited a degradation time of ~5 d in PBS, serum-free medium, and serum medium with loaded cells. In serum medium in the absence of cells, the microspheres degraded completely in a little over 8 h and exhibited only minimal increase in size at that time point (11%). Slower degradation was noted for microspheres confined in a bulk slab hydrogel (Figure 5D). By comparing the microsphere size at 2 and 168 h, we noted the most pronounced decrease in size in serum medium (56%), followed by serum medium with loaded cells (43%), and followed by PBS (34%). A minimal decrease in size was noted for confined microspheres in serum-free medium (18%). Overall, the highest decrease in confined microspheres was noted for the microspheres in serum medium, while the lowest was noted for microspheres in serum-free medium. Microsphere swelling and degradation rate as a function of confinement, medium, and cell loading are summarized in Table 2. 2.3. Cell Viability and Multicellular Aggregate Formation in Templated Hydrogels We investigated the potential of the developed templated-hydrogels to support glioblastoma multicellular aggregate formation. Note that the bulk hydrogel slab had mechanical properties (Figure S4) compatible with the chosen application. The bulk PEGDA hydrogel had a storage modulus of 14.37 ± 0.21 kPa, which is similar to the reported modulus of glioblastoma tumor tissue.16 However, the mechanical strength of the gel could be further modulated to suit a chosen application by manipulating the polymer concentration and molecular weight or by using 10 ACS Paragon Plus Environment

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a different biomaterial. The initial storage modulus of the degradable microspheres was 6.83 ± 0.01 kPa. For both materials (slab and microspheres), the storage modulus was significantly higher than the loss modulus and was not affected by increase in frequency from 0-10 Hz, indicating the formation of a stable crosslinked hydrogel. First we established hydrogel biocompatibility by culturing U251 glioblastoma cells in microspheres only and in templated hydrogels and following cell viability via live/dead staining for 7 days (Figure 6). Representative live/dead images, where all cells were stained with 3,3’dioctafecyloxacarbocyanine perchlorate (DiOC) (green) and dead cells were stained with propidium iodide (PI) (red) are shown in Figure 6A,B. Overall, we noted excellent cell viability of >90% for all conditions at all time points, demonstrating the suitability of the hydrogel microspheres and the templated hydrogels for prolonged cell culture (Figure 6C). The high cell viability at 2 h in non-confined microspheres (94.3 ± 2.4%) and in templated hydrogels (94.6 ± 1.7%), also demonstrated that the microsphere or hydrogel bulk encapsulation process did not have an adverse effect on the cells. We also noted that by day 7, dead cells were mostly present at the center of the cell aggregates (Figure 6B), which could be related to oxygen and nutrient diffusion limitations. Next, we monitored the cells in the templated hydrogels, where cells were cultured for 14 d to ascertain multicellular aggregate formation (Figure 7). We used a membrane stain (WGA) and a nucleus stain (DAPI) to visualize the cells via confocal microscopy. We noted an even cell distribution at day 1 of culture (Figure 7A, B), indicating that the cells were evenly dispersed in the hydrogel microspheres inside the templated hydrogel. At day 14 the cells formed a multicellular aggregate, indicating complete microsphere degradation (Figure 7C, D). The average aggregate size was 153.7 ± 61.7 μm, where most of the aggregates were in the size rage



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of 180-200 μm. There was however some size distribution – the aggregates did not seem to grow beyond 220 μm at 14 days of culture, while the smallest aggregate measured was 54 μm. To confirm that the encapsulated cells formed an aggregate due to microsphere degradation and not cell movement inside the microsphere, we also cultured the templated hydrogel with fluorescent microbeads, where the beads showed even dispersion in the microspheres at day 1 and aggregation at day 14 (Figure 7E, F). We further examined the cells for aggregation and morphology at days 1, 7 and 14 (Figure 8). We noted individual cells at day 1, small cell aggregates and some individual cells at day 7 and larger cell aggregates with no individual cells at day 14. The cells retained their spherical morphology during degradation and did not seem to interact with the templated hydrogel, which could be attributed to the inert nature and lack of attachment sites of the PEGDA hydrogel slab.16, 35 2.4. Drug Screening To demonstrate the utility of the developed multicellular aggregate-laden templated hydrogels for drug screening applications, we conducted a small drug screen with five FDAapproved anti-cancer drugs: paclitaxel (PTX), doxorubicin (DOX), camustine (BCNU), lomustine (CCNU) and temozolomide (TMZ) (Figure 9A). All of the chosen drugs have been tested or approved for use against glioblastoma.36, 37 All drugs were used at a single bolus dose as explained in the methods. DMSO was used as a vehicle only control. Furthermore, to determine whether the properties of the surrounding matrix affect cell response to drugs, we used inert PEG only or cell-adhesive PEG-RGDS (0.8 mmol in RGDS) as the bulk hydrogel slab into which multicellular aggregates were templated. Our results indicated that at the concentrations used in this study, DMSO had no negative impact on cell viability. As expected, all tested drugs led to reduced cell viability compared to no 12 ACS Paragon Plus Environment

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drug control. The viability of U251 cells seeded in inert PEG templated gels was 54, 41, 45, 15 and 48% for PTX, DOX, BCNU, CCNU and TMZ, respectively. For most of the drugs tested, cell viability was overall slightly, but insignificantly higher for cell aggregates seeded in the modified PEG-RGDS templated hydrogels compared to the unmodified PEG hydrogels. The exception was the cell aggregate response to CCNU and TMZ, where a significant 35% and 28% increase in cell viability, respectively, was noted in the presence of integrin-mediated cell-matrix interactions in the PEG-RGDS gels compared to cell aggregates in the inert PEG gels. There was no statistical difference in cell viability between cell responses to the different drugs when multicellular aggregates were encapsulated in the PEG-RGDS templated hydrogels. Confocal imaging of the U251 multicellular aggregates revealed differences in cell morphology for cells seeded in the inert PEG versus cell-adhesive PEG-RGDS templated hydrogels (Figure 9B). Specifically, aggregates exhibited a smooth surface in the inert PEG gels but had a ruffled appearance in the PEG-RGDS gels, indicative of integrin-mediated cell-matrix interactions.

3. Discussion In this study we designed and characterized templated PEG hydrogels for multicellular aggregate encapsulation (Figure 2). While macroporous hydrogels in the presence of cells have been recently created, the authors utilized gelatin microspheres as porogens, which cannot be prepared in the presence of cells; hence, cells were encapsulated in the hydrogel slab rather than in the microspheres.7, 15 Here, we report a technique for macroporous hydrogels preparation in the presence of cells, where cells could be encapsulated in the microspheres or in the hydrogel slab. We only discuss depositing cell-laden microspheres in a cell-free hydrogel slab for the purpose of creating hydrogel-grown multicellular aggregate and spheroid cultures. However, our



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approach could be extended to the creation of physically segregated co-culture systems where one cell type is trapped inside the macropores and the other cell type is trapped inside the bulk of the hydrogel, as both are biocompatible. The approach also does not prevent creating additional micro and macroporosity in the bulk of the gel by using cell-free degradable microspheres. We chose PEG because of our ability to finely and reproducibly control its properties, including degradability as well as PEG’s excellent biocompatibility.27, 38 Not surprisingly, PEG hydrogels have broad utility and are some of the most commonly investigated materials for biomedical applications.38-40 Specifically, we encapsulated cell-laden biodegradable 4-arm PEGAc/DTBA microspheres in PEGDA hydrogel slabs. Upon microsphere degradation, cells were deposited in the PEGDA macropores of controlled sizes and formed multicellular aggregates. To fabricate biocompatible and biodegradable microspheres, we utilized a Michael-type addition reaction between an acrylate (4-arm PEG-Ac) and a dithiol crosslinker (hydrolytically degradable DTBA), which is a mild and specific gelation chemistry suitable for in situ cell encapsulation.27 In this reaction, a thiolate ion (in DTBA) forms in the presence of bases such as TEA or HEPES and attacks the unsaturated acrylate moiety (in 4-arm PEG-Ac) leading to the formation of a degradable thioester bond.41, 42 We used a single crosslinker as a proof-ofprinciple. A greater system versatility can be achieved by choosing hydrolytically degradable dithiol crosslinkers of varying degradation times, which we have previously synthesized and characterized, 27, 38 or enzymatically degradable peptide crosslinkers that would allow cellcontrolled degradation and microsphere remodeling.26 Such versatility will enable tailoring degradation to match cell proliferation and ensure cell-matrix interactions during aggregate and spheroid formation.


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To produce uniform cell-laden hydrogel microspheres in a variety of sizes, we used a custom T-junction microfluidic set-up (Figure 1). This microfluidic setup was chosen for its simplicity and because it enabled us to produce microspheres of diverse sizes (118 – 480 µm) in a narrow size distribution for most conditions (%CV of 12 – 23) (Table 1). Diverse size microspheres in turn would allow the production of a templated hydrogel of varying macropore sizes targeting multiple applications. For example, a macroporosity of 160 – 270 μm has been shown beneficial for vascularization,43 while 300 - 500 μm would be suitable for tumor spheroid formation.44 To control microsphere size, we manipulated the PEG microsphere precursor solution (disperse phase) flow rate and the olive oil (continuous phase) flow rate (Table 1). Corroborating previous studies,45 we determined that increasing the continuous phase flow rate and decreasing the disperse phase flow rate resulted in smaller microspheres (Table 1). Higher continuous phase flow rate has been shown to result in smaller microspheres due to increased friction between the phases, contributing to droplet detachment. 31 Importantly, for most of the conditions studied, microsphere polydispersity was low (≤10% CV), which could be attributed to the higher viscosity of olive oil (continuous phase) used in the T-junction set-up compared to the viscosity of the disperse phase;31 the viscosity of oil is approximately 4-times higher than the viscosity of PEG.46,47 Note that a %CV of 15% or below is preferred because it signifies relatively monodisperse microspheres.48 Monodisperse microspheres at this stage would lead to a templated scaffold with uniform size macropores. Next, we tested microsphere swelling and degradation in various environments as microsphere degradation would lead to macroporosity (or templating) of the bulk PEG hydrogel (Figure 5, Table 2) and noted several trends. First, microsphere degradation was accelerated in the presence of serum in the media, possibly due estereases degrading the ester bonds in the



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hydrogel as shown by Martens et al.,49 previously. Previous studies have also shown that certain serum proteins, such as bovine serum albumin (BSA), increase the rate of hydrolytic degradation in poly(lactic acid) materials due to BSA’s esterase-like activity.50, 51 Degradation was also affected by cell loading, where cell-laden microspheres degraded slower than cell-free ones. Our results corroborate findings from Bryant et al.,52 who demonstrated that the degradation rate of PEG hydrogels was lower in the presence of cells due to differences in the water content in the cell-laden hydrogels compared to hydrogels alone. Lastly, degradation of microspheres was slower in a confined environment compared to a non-confined one. These results could be attributed to the lower water accessibility in the microspheres in the confined environment as well as the inability of the microspheres to swell upon degradation and, hence, to imbibe more water. Differences in hydrogel swelling due to confinement, where confinement leads to reduced swelling, have been reported previously.34, 53 The microspheres described here degrade by hydrolysis due to the presence of a thioester bond (Figure S1).27 A thioester is formed at every crosslink site and then randomly hydrolyzed to give an alcohol (4-arm PEG-OH) and an acid degradation products. When the degradation products become sufficiently small, they are able to diffuse through the bulk hydrogel material. Since scission eventually happens at every ester bond (at complete degradation), the degradation products have a molecular weight very similar to that of the original 4-arm PEGAc polymer, which is 10 kDa. Dynamic light scattering measurements in water for various PEG polymers have shown that star polymers have smaller hydrodynamic radii, Rh, than linear polymers of the same molecular weight.54 Rh of the degradation products for the PEG microspheres described here should be 90% viability up to 7 days in the microspheres alone or in the templated hydrogels (Figure 6). This result was expected as we have previously demonstrated that our materials, microsphere fabrication methods, and degradation products are compatible with in situ cell encapsulation.27, 55 Also, the hydrogel slab had ~1.4 mm thickness when equilibrium-swollen in serum medium, hence, major diffusion limitations for oxygen and nutrients into the PEG hydrogel were not expected.56 Importantly, we demonstrated that cells were able to form multicellular aggregates (through aggregation) in the templated hydrogels upon microsphere degradation (Figure 7, Figure 8). Note that the cells did not completely fill the void left behind upon microsphere degradation even after 14 d of culture. Here, we used cell densities of 1x106 cells/mL (degradation studies) or 3x106 cells/mL (cell viability and spheroid studies), which are typically used for 3D cell culture experiments.38, 57 However, a higher initial cell density could be used to produce larger aggregates in a shorter time frame.44 For example, we have shown that cells can be encapsulated in biodegradable PEG hydrogel microspheres at a cell density of up to 1x108 cells/mL without adverse effect on cell viability.55 Note that even cancer cells that are dispersedly seeded in a PEG hydrogel will eventually form aggregates or spheroids.58-60 However, there are some limitations to such approaches. First, in a non-degradable scaffold such aggregates or spheroids would be formed mostly by clonal



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expansion and not by aggregation as in our approach, which would not be representative of the cell heterogeneity present in multicellular spheroids. Second, aggregates or spheroids may overlap during growth, which would be undesirable for single aggregate or spheroid analysis.61 Third, such aggregates or spheroids would be of diverse sizes, which would be a limitation for certain high-throughput applications, such as drug screening and therapy development,60 or for assessing other cell outcomes, where differences in size would lead to irreproducible outcomes. For example, multicellular spheroid size has been shown to affect cell metabolic activity, diffusion processes as well as albumin production and cytochrome P450 activity, which are key parameters of cellular physiology in hepatocyte cultures, of HEPG2 hepatocytes.62 Overall, multicellular aggregates and spheroids are becoming a useful tool in drug screening and drug development applications,63 as well as building blocks of various complex 3D tissues.64 At the same time, emerging research has demonstrated differential cell behavior for multicellular aggregates and spheroids grown in a liquid versus those cultured in a hydrogel.18 Hence, our approach represents an important step towards enabling the use of hydrogel-grown multicellular aggregates and spheroids in various biomedical applications. Lastly, as an example application we tested the hydrogel-encapsulated glioblastoma aggregates in a small drug screen. We used a single bolus dose of several FDA-approved chemotherapeutics all of which have been tested against glioblastoma, namely PTX, DOX, BCNU, CCNU and TMZ. PTX binds to tubulin and inhibits the disassembly of microtubules, thereby resulting in the inhibition of cell division.65 DOX intercalates between base pairs in the DNA helix, thereby preventing DNA replication and ultimately inhibiting protein synthesis.66 BCNU alkylates and crosslinks DNA during all phases of the cell cycle, resulting in disruption of DNA function, cell cycle arrest, and apoptosis.67 CCNU alkylates and crosslinks DNA, thereby


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inhibiting DNA and RNA synthesis.36 TMZ is a DNA alkylating agent known to induce cell cycle arrest at G2/M and to eventually lead to apoptosis.68 To determine if hydrogel properties could affect multicellular aggregate drug responsiveness, we used an inert PEG hydrogel and a cell-adhesive RGDS-modified PEG hydrogel (Figure 9). Note that our ability to create hydrogelgrown aggregates or spheroids of controlled sizes is critical for drug screening, as aggregate or spheroid diameter has been shown to affect cell responsiveness to drugs due to cell heterogeneity and differences in drug, oxygen and nutrient diffusion with aggregate or spheroid diameter.69 Cell aggregate-matrix interactions with the PEG-RGDS hydrogel was confirmed by the ruffled spheroid appearance in the presence of the ligand, compared to a smooth aggregate surface in the inert hydrogel (Figure 9B). For two drugs - CCNU and TMZ - cell aggregates appeared more susceptible to the drugs when encapsulated in the inert PEG hydrogel in the absence of cell-matrix interactions. This result was not surprising as integrin binding has been previously implicated in conferring drug resistance of dissociated cells, including glioblastoma cells in particular.58, 70, 71 Corroborating current findings, we have previously shown that when dissociated glioblastoma cells were seeded in a soft alginate hydrogel, cells were significantly less susceptible to toxins in the RGDmodified gels, compared to cells in the inert unmodified gels.70 Currently integrin signaling is being widely explored as a therapeutic strategy.72, 73 For example, RGD-integrin antagonists that inhibit cell adhesion and induces anoikis in glioblastoma cells have been suggested as a therapeutic approach to reduce malignancy.74 Hence, our results demonstrate that the aggregateladen templated hydrogels developed here could be successfully used in a drug screening application to parse the effect of matrix properties on multicellular aggregate drug responsiveness.



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4. Conclusion Here we developed a templated PEG hydrogel by incorporating cell-laden biocompatible and biodegradable PEG microspheres in non-degradable PEG hydrogels. The microspheres were prepared via a Michael-type addition between a 4-arm PEG-Ac and a dithiol crosslinker, namely DTBA. While we demonstrated the developed technology using a single crosslinker, there is a wide variety of hydrolytically or enzymatically degradable dithiol crosslinkers described in the literature, which makes our approach highly tunable. The microspheres were prepared in a range of sizes (118 - 480 μm) and with a low polydispersity by a simple microfluidic technique. Microsphere degradation was dependent on the incubation medium, where the presence of serum resulted in faster degradation than all other conditions. Microsphere degradation was slowed by the presence of cells in the microspheres or by sphere confinement in a bulk hydrogel scaffold. The developed templated hydrogel was compatible with in situ cell encapsulation, where >90% cell viability was noted over 7 days in culture. Multicellular aggregates formed in the macropores left behind in the templated hydrogel upon cell-laden microsphere degradation. The hydrogel-encapsulated glioblastoma aggregates were tested in a small drug screen, which demonstrated that cell-matrix interactions affected cell drug responsiveness and hence, underscored the importance of multicellular aggregate encapsulation in a physiologically relevant scaffold. The developed templated hydrogel would be useful in a variety of biomedical applications, such as drug screening or as building blocks of complex 3D tissues.

5.

Materials and Methods

5.1. Materials 20 ACS Paragon Plus Environment

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4-arm polyethylene glycol acrylate (4-arm PEGAc; 10 kDa) was obtained from Jenkem Technology USA Inc. (Plano, TX). The chemotherapeutics paclitaxel (PTX), doxorubicin (DOX), carmustine (BCNU), lomustine (CCNU) and temozolomide (TMZ), as well as triethanolamine (TEA) and 2-amino butane dithiol (DTBA) were purchased from Sigma Aldrich (St. Louis, MO). Polyethylene glycol diacrylate (PEGDA; 5 kDa) and polyethylene glycoldithiol (PEG-diSH; 3.4 kDa) were obtained from Laysan Bio Inc. (Alabama, USA). Phosphate buffered saline (PBS, 10X, pH 7.4), fluorescent polystyrene beads with diameter of 2 μm (Fluoro-MaxTM Fluorescent Red, 542/612 nm), hydrochloric acid (HCl), and wheat germ agglutinin (WGA) were purchased from Thermo Scientific (Waltham, MA). Irgacure 2959 was purchased from BASF Corporation (Florham Park, NJ). 4-(2-hydroxyethyl)-1piperazineethanesulfonic acid (HEPES) was purchased from Acros Organics (Pittsburgh, PA). Gas-tight syringes (500 μl, Model 1750 LT) were purchased from Hamilton (Reno, NV). Homo sapiens brain glioblastoma U251 cells were purchased from ATCC (Manassas, VA). Cell stains 3,3’-dioctafecyloxacarbocyanine perchlorate (DiOC) and 4’ and 6-diamidino-2-phenylindole (DAPI) were purchased from Life Technologies (Carlsbad, CA). Propidium iodide (PI) was purchased from MP Biomedical LLC (Solon, OH). Teflon tubing was purchased from Scanivalve Corp (Liberty Lake, WA). Olive oil was purchased from a local grocery store. Cell strainers of mesh size 100 μm were purchased from Greiner bio-one (Kremsmünster, Austria). Silicone isolator sheets (0.5 mm thick) were purchased from Grace Bio Labs (Bend, Oregon) and were used as spacers between glass slides. Fetal bovine serum (FBS) and penicillin/streptomycin (pen/strep) were purchased from Hyclone (Logan, UT). Roswell Park Memorial Institute (RPMI)-1640 medium and 0.05% trypsin/0.02% ethylenedinitrilotetraacetic acid (EDTA) were purchased from Coring (Coring, NY). Glycine-Arginine-Cysteine-Aspartic Acid-Arginine-



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Glycine-Aspartic Acid-Serine (GRCD-RGDS) peptides were obtained from Peptide 2.0 (Chantilly, VA). PEEK MicroTee T-junction and tubing were purchased from VICI Valco Instruments (Houston, TX). 5.2. Preparation of 4-arm PEG-RGDS 4-arm PEG-RGDS was mono-functionalized with 80% modification efficiency following a previously developed protocol.38, 75 Briefly, 8% w/v GRCD-RGDS was dissolved in 5% acetic acid in de-ionized (DI) water and 11.5% w/v 4-arm PEG-Ac was dissolved in 0.3 M TEA buffer. Both solutions were combined in a 1:4 v/v ratio of RGDS:Ac and reacted with stirring for 30 min at room temperature. The concentrations were chosen such that, assuming an ideal reaction, one of the acrylate groups of each 4-arm PEG-Ac macromere will be conjugated with RGDS. Upon reaction the solution was placed at -80°C for 15 min and then lyophilized (VirTis Sentry 2.0 Lyphilizer) overnight. The 4-arm PEG-RGDS product was purged with argon and stored in a desiccated environment at -20°C until use. 5.3. Fabrication of Degradable Microspheres via Microfluidics To prepare the microspheres, a 20% w/v stock solution of 4-arm PEG-Ac was prepared in 0.3 M TEA buffer of pH 7.4, unless otherwise noted. Immediately prior to use, a 5% w/v stock solution of the DTBA crosslinker was also prepared in 0.3 M of TEA buffer of pH 7.4, unless otherwise noted. To make a 10% w/v gel precursor solution, 4-arm PEG-Ac and DTBA were combined in a 1:1 molar ratio of Ac:SH and TEA buffer was added to achieve the desired polymer concentration. In certain cases, red-fluorescent polystyrene beads were added at a final concentration of 0.01% w/v for subsequent microsphere visualization under a fluorescent


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microscope. The precursor solution was vortexed for 30 sec and loaded in a gas-tight syringe for microsphere fabrication. To fabricate hydrogel microspheres, a custom T-junction droplet generator was developed (Figure 1). The droplet generator consisted of a 100 μL gas-tight syringe, a 60 mL syringe, a mixing tee with a bore diameter of 0.5 mm, two programmable single-syringe micropumps (NE-1000, Farmingdale, NY), female luer locks, and tubing with an inner diameter of 0.02”. To fabricate microspheres, the 100 μL gas-tight syringe was loaded with PEG precursor solution (disperse phase), placed on a micropump and was connected to the top port of a mixing tee via a luer lock and tubing. The 60 mL syringe loaded with olive oil (continuous phase) was placed on a micropump and connected to the side port of a mixing tee via a luer lock and tubing. Tubing was also connected to the remaining side port of the mixing tee and placed in an olive oil bath for droplet collection. To control microsphere size we varied the PEG precursor solution flow rate (5 - 60 μL/min) and the olive oil flow rate (0.5 - 1 mL/min). Hydrogel droplets were collected in an olive oil bath positioned on a shaker platform (Chemglass Life Science, Vineland, NJ. Upon collection, the droplets were left in the oil bath for ~15-30 min to complete gelation, collected by centrifugation, and washed with sterile 1X PBS on a cell strainer to remove residual oil. 5.4. Templated Hydrogel Preparation Schematic describing the preparation of templated hydrogels as well as the chemical structure of the polymers used to prepare the degradable microspheres and the bulk hydrogel slab is presented in Figure 2.



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Bulk hydrogel slabs were prepared from PEGDA via ultraviolet (UV) crosslinking as described previously.76, 77 Briefly, stock solution of 1% w/v Irgacure 2959 was prepared by dissolving Irgacure 2959 in de-ionized (DI) water and sonicating the solution in a bath sonicator (Branson, 2800, Fisher Scientific, Waltham, MA) for 90 min. The Irgacure 2959 solution was stored shielded from light at 4°C until use. Stock solution of 20% w/v PEGDA was prepared in 1X PBS pH 7.4 and stored at 4°C for up to one week or used immediately. A 10% w/v PEGDA hydrogel precursor solution containing 0.1% w/v Irgacure 2959 was prepared in 1X PBS pH 7.4 and vortexed for 30 s. To prepare the templated hydrogel, the PEGDA precursor solution was mixed with degradable 4-arm PEG-Ac/DTBA microspheres (50-100 microspheres per 50 µL of gel precursor solution). The microsphere-containing solution was sandwiched between two glass slides pre-coated with Rain-X® and separated by 1 mm silicone spacers. Hydrogels were formed by exposure to UV light (365 nm; 4.81 mW/cm2) for 10 min. In some cases, 4-arm PEG-RGDS was added at 1% w/v (i.e. 1% w/v 4-arm PEG-RGDS + 9% w/v PEGDA) to the bulk hydrogel precursor solution to elicit cell attachment. 5.5. Cell Maintenance and Encapsulation in Degradable Microspheres U251 cells were cultured in RPMI-1640 medium supplemented with 10% v/v FBS and 1% v/v pen/strep in a humidified incubator at 37°C and 5% CO2. Medium was replaced every other day until cell confluency was reached. To subculture, confluent cells were harvested by a 5 min exposure to 0.05% trypsin/0.02% EDTA. To encapsulation in microspheres, cells were added directly to the hydrogel precursor solution at 1x106 cells/mL (for degradation studies) or 3x106 cells/mL (for cell viability and 24 ACS Paragon Plus Environment

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spheroid formation studies). Microspheres were then prepared as described above under sterile conditions inside a tissue culture hood. Only cells up to passage 20 were used for experiments. 5.6. Gelation Time Gelation time was measured by the inverted tube method.78 Each hydrogel sample was prepared at room temperature in a microfuge tube, vortexed, and monitored by repeated inversion. The gelation time was noted as the time when the precursor solution was no longer flowing upon inversion, indicating the formation of a hydrogel network. 5.7. Rheology All rheology experiments were conducted using an AR-2000 ex rheometer (TA Instruments) with 20 mm parallel plate geometry. The absence of slip was verified by running experiments with various gap heights. 79 A frequency of 1 - 10 rad/s and a constant strain of 1%, which was within the linear viscoelastic region, were used for mechanical testing of all hydrogels. For rheology testing, hydrogels were prepared as 20 mm diameter discs of thickness ~500 μm and equilibrium swollen overnight in 1X PBS, pH 7.4. Prior to measurements, excess water from the hydrogel surface was blotted carefully with a KimWipe®. For gel point studies by rheology, a time sweep was performed at a constant frequency of 0.5 Hz and a constant strain of 0.05%. The evolution of storage modulus, G’, was followed. For each experiment, the hydrogel precursor solution was pipetted between the two rheometer parallel plates and the gap was adjusted to 0.1 mm. Precursor solution evaporation was prevented by a custom-made rheometer attachment, which ensured a humidified environment during measurements. 5.8. Microsphere Size and Polydispersity



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To determine size and polydispersity, microspheres were imaged and analyzed immediately upon collection (before washing) unless otherwise noted. The microspheres were imaged using an inverted microscope (Zeiss Axiovert 200) at 5x magnification. Microsphere diameter was analyzed from phase contrast images via Image J (freely from the NIH at http://rsweb.nih.gov/ij/) using the Line Analyzer plug-in. Polydispersity was calculated as percent coefficient of variance (%CV) calculated as the standard deviation divided by the mean. 5.9. Microsphere Degradation Microspheres prepared from 4-arm PEGAc and DTBA crosslinker degraded by ester hydrolysis (Figure S1). Microsphere degradation (cell-laden and cell-free) was assessed by following the changes in microsphere diameter at pre-selected time points. Specifically, microspheres were imaged under an inverted fluorescent microscope (Zeiss, Axiovert 200M, Oberkochen, Germany) at 10X zoom and microsphere diameter was analyzed with ImageJ. Degradation was followed for microspheres immersed in an aqueous medium directly (nonconfined) or encapsulated in a hydrogel slab and then immersed in an aqueous medium (confined) (Figure S2). Degradation was assessed in the following aqueous media: 1X PBS, serum-free medium, serum medium (10% FBS and 1% pen/strep), and serum medium with loaded cells (1×106 cells/mL). All degradation experiments were conducted in a humidified environment at 37°C and 5% CO2 with a medium change every other day. Red-fluorescent polystyrene beads were added to all microspheres (including cell-laden ones) for visualization. 5.10. Cell Viability and Multicellular Aggregate Formation in Templated Hydrogels Cell viability was evaluated with live/dead staining. Briefly, cells were first incubated with 0.02 μg/mL of DiOC (green, membrane-permeable dye) for 24 h prior to experiments. 26 ACS Paragon Plus Environment

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Stained cells were then harvested and cell suspension was mixed with 4-arm PEG-Ac/DTBA hydrogel microsphere precursor solution to give a final cell concentration of 3×106 cells/mL. Note that a higher cell density was chosen (compared to degradation experiments) to foster spheroid formation. Cell-laden 4-arm PEG-Ac/DTBA microspheres only and templated hydrogels were prepared as described above. The samples were then placed in 48-well plates, where 50-100 microspheres or 50 μL microsphere-laden gel precursor solutions were dispensed per well and submerged in 500 μL of serum medium. The samples were incubated at 37oC and 5% CO2 with medium change every other day. At pre-selected time points, PI (0.01 mg/mL; red, membrane-impermeable dye) was added and the cells were incubated for additional 30 min. The cells were then imaged under an inverted fluorescent microscope (Ziess, Axiovert 200M) and percent cell viability was calculated as the number of viable cells (total number of cells stained with DiOC minus dead cells stained with PI) normalized by the number of total cells (stained by DiOC). Additionally, for superior visualization of multicellular aggregate formation, cells were cultured in the templated hydrogels for 1, 7 and 14 d, where medium was changed every other day. Cells were then stained with WGA (cell membrane) and DAPI (cell nucleus) as per the manufacturers’ instructions. The cell-laden templated hydrogels were incubated with the dyes for 30 min, washed trice with 1X PBS and imaged with a confocal microscope (Leica TCS SP8 STED 3X, Leica Microsystems, Exton, PA) with an excitation wavelength of 568 nm for WGA (at 1% power of 0.5 mW at the image plane of supercontinuum white light laser detected at 602640 using a HyD detector with 55.7% gain) and 405 nm for DAPI (at 10% power of a 405 diode laser at 1 mW at the image plane and detected between 430-472 nm using a PMT with a voltage gain of 782). 27 ACS Paragon Plus Environment


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5.11. Drug Screening For drug screening, U251 cells were first incubated with 0.02 μg/mL of DiOC for 24 h prior to the experiments. Cells were cultured at 3×106 cell/mL in macroporous hydrogels for 7 d to allow for multicellular aggregates formation. At day 7, bolus drug doses were added to the cell medium at concentrations ten times higher than the reported drug IC50 (50% inhibitory concentration) for monolayer U251 cells seeded on tissue culture polystyrene (TCP).80 Specifically, paclitaxel (PTX), doxorubicin (DOX), carmustine (BCNU), lomustine (CCNU) and temozolomide (TMZ) were added at 4.0010-3, 4.0010-2, 5.0010-2, 3.00, and 1.00 mM, respectively. DMSO (4.5% v/v) was used as vehicle only control. The cell-laden templated hydrogels were incubated with the drugs for 48 h at 37oC and 5% CO2. The medium was then changed with fresh drug-free medium and cells were stained with PI at 0.01 mg/mL, imaged under an inverted fluorescent microscope (Ziess, Axiovert 200M) and cell viability was measured via live/dead staining as described above. Note that >85% cell viability was confirmed on control gels prior to drug screening. 5.12. Statistical Analysis Results are expressed as average ± standard deviation (SD) from 3 independent experiments, where 6-10 samples were used per experiment. Multiple groups were compared using single factor analysis of variance (ANOVA) with a post-hoc test. Two-tailed Student t-test was used to compare between two groups. A value of p < 0.05 was considered significant. Supporting Information Available


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Table 1. Microsphere diameter and size distribution as a function of oil (continuous phase) and PEG (disperse phase) flow rates. Polydispersity is represented as %CV (percent coefficient of variance). Conditions Oil Flow Rate (mL/min)

0.5

1

Results

PEG Flow Rate (μL/min)

Microsphere Diameter (μm)

%CV

5 10 20 40 N/A 5 10 20 40 60

119 ± 16 153 ± 35 266 ± 31 309 ± 40 N/A 118 ± 21 125 ± 18 181 ± 34 327 ± 63 480 ± 108

14 23 12 13 N/A 18 14 19 19 23


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Table 2. Summary on the effect of microsphere confinement, cell loading and incubation medium on microsphere swelling and degradation rate.

Effect on sphere degradation rate

Condition

Effect on sphere swelling

Confinement

Cell loading Serum in incubation medium



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Figure 1. A) Schematic of microsphere fabrication via microfluidics. An example of phase contrast image of PEG microspheres is shown, where microspheres were loaded with green fluorescent microparticles for contrast; scale bar represents 100 μm. B) An image of microsphere fabrication via microfluidics. C) A schematic of the T-junction.


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Figure 2: Templated hydrogel preparation. A) Degradable PEG microspheres were prepared from 4-arm PEG-Ac and DTBA crosslinker. Cells well directly added to the hydrogel precursor solution to result in cell-laden microspheres upon fabrication. B) Phase contrast images of PEG microspheres prepared via microfluidics. Scale bar is 100 μm. C) Templated hydrogels were prepared by adding degradable cell-laden microspheres to a PEGDA precursor solution. The microspheres were then entrapped in the bulk hydrogel upon UV polymerization of the bulk gel. D) A fluorescent image of microspheres encapsulated in a templated hydrogel slab. The microspheres were embedded with fluorescently labeled polystyrene beads for imaging. Scale bar is 1.5 mm.



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Figure 3. A) Effect of buffer pH and buffer type on gelation time as measured by the inverted tube method. *indicates significant differences within the same buffer type of varying pH and #indicates significant differences between buffer types of the same pH (p

Templated Macroporous Polyethylene Glycol Hydrogels for Spheroid

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