Microfluidic fabrication of multi-stimuli responsive tubular hydrogels for

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Microfluidic fabrication of multi-stimuli responsive tubular hydrogels for cellular scaffolds Dongwan Kim, Ara Jo, Kusuma Betha Cahaya Imani, Dowan Kim, Jin Woong Chung, and Jinhwan Yoon Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b00453 • Publication Date (Web): 19 Mar 2018 Downloaded from http://pubs.acs.org on March 20, 2018

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Microfluidic fabrication of multi-stimuli responsive tubular hydrogels for cellular scaffolds

Dongwan Kim1+, Ara Jo2+, Kusuma Betha Cahaya Imani3, Dowan Kim3, Jin-Woong Chung2,*, Jinhwan Yoon3,*

1

Department of Chemistry, Dong-A University, 37 Nakdong-Daero 550 beon-gil, Saha-gu,

Busan 49315, Republic of Korea; 2

Department of Biological Science, Dong-A University, 37 Nakdong-daero 550 beon-gil, Saha-

gu, Busan, 49315, Republic of Korea; 3

Department of Chemistry Education, Graduate Department of Chemical Materials, and Institute

for Plastic Information and Energy Materials, Pusan National University, 2 Busandaehak-ro 63 beon-gil, Geumjeong-gu, Busan, 46241, Republic of Korea

* E-mail: (J.Y.) [email protected], (J-W.C.) [email protected] + D.K and A.J contributed equally to this work.

ACS Paragon Plus Environment

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ABSTRACT Stimuli-responsive hydrogel microfibers and microtubes are in great attention for the biomedical application due to their similarity with the native extracellular matrix. In this study, we prepared pH- and temperature-responsive hydrogel microfibers and microtubes using a microfluidic device through alginate-templated photopolymerization. Hydrogel monomer solutions containing N-isopropylacrylamide (NIPAm) and sodium acrylate (SA) or allyl amine (AA) were irradiated with UV light to invoke in situ photopolymerization. A repulsive force between the ionized SA or AA groups caused by protonation/deprotonation of the acrylate or amine groups, respectively, led to changes in the diameters and wall thicknesses of the fibers and/or tubes depending on the pH of the medium. PNIPAm is a well-known thermally responsive polymer wherein the NIPAm-based copolymer microfibers exhibited a thermal behavior close to the lower critical solution temperature. We have demonstrated that these multistimuli responsive volume changes are fully reversible and repeatable. Furthermore, the positively charged microfibers were shown to exhibit cell adhesion and the number of cells attached to the microfibers could be further increased by supplying nutrients, presenting the possibility of their application in tissue engineering and other biomedical fields.


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1. INTRODUCTION Hydrogels are physically or chemically crosslinked hydrophilic polymer networks with high water content that allows them to maintain their three-dimensional elastic structures and affords biocompatibility. Recently, microfiber-shaped hydrogels have received extensive interest1-15 since their shape is similar to that of the native extracellular matrix, which plays a vital role in the construction of tissues and organs. Accordingly, the biocompatibility of hydrogels combined with the vascular structure of microfibers would indicate that microfibrous hydrogels have enormous potential in tissue engineering1-6. Microfluidic technology based on coaxial flow has been employed to fabricate hydrogel microfibers and microtubes1-15. This technology enables simple, continuous, and large-scale production of various microstructures such as microfibers1-8 and microtubes9-10 with uniform shapes and sizes. To further enhance the advantages of hydrogel microfibers, stimuli-responsive properties would be an interesting development11-15. Volume phase transition can be induced in the hydrogel microfibers by manipulating various stimuli such as pH13, temperature14-15 and light15. Among these stimuli, pH is particularly important for responsive hydrogels for biomedical applications16-17, since different organs and tissues in the human body have different pH values and show pH-dependent functionalities. It should be noted that hydrogel microfibers that exhibit volume change in response to pH have been previously reported13. In that study, the hydrogel microfibers were prepared by the interpenetration of the pH-responsive polymer poly(N-isopropylacrylamide-co-acrylic acid) (p(NIPAm-co-AAc)) with alginate microfibers. However, the volume changes in response to pH and temperature were not fully reversible, limiting its practical application. Previously, it was demonstrated that the reversible volume change of the hydrogels is crucial for controlling of the drug release rate18 and the movement of


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the soft actuators19. Because of the calcium-alginate deformation that occurred upon the shrinking of p(NIPAm-co-AAc), the microfibers did not return to their initial shape, instead returning to only about 70% of their original size. In the current study, we have prepared crosslinked hydrogel microfibers and microtubes that exhibit fully reversible and repeatable volume change in response to temperature and pH changes. Crosslinked networks of poly(N-isopropylacrylamide-co-sodium acrylate; PNS) and poly(N-isopropylacrylamide-co-allyl amine; PNA) in fibrous or tubular form were prepared with a microfluidic device through photopolymerization using an alginate templating method. Through the protonation/deprotonation of pendant ionic groups and the lower critical solution temperature (LCST) behavior, these hydrogel micro-objects showed excellent responsiveness to pH and temperature. Moreover, to investigate their potential application in the biomedical field, we attempted to attach living cells to the microfiber surfaces through the electrostatic attraction between the charged hydrogel and the cell membrane. In general, much research effort has been devoted to the promotion of specific cell interactions with hydrogel microobjects by the introduction of anchorage sites on the scaffold20-21. Incorporation of the arginylglycylaspartic acid (RGD) peptide (Arg-Gly-Asp) on hydrogel surfaces is an effective method for enhancing cell attachment owing to the ability of RGD to bind to integrin receptors. However, specific chemical reactions for binding this peptide to the hydrogel must be performed22-23. Motivated by a previous report regarding a charged hydrogel for cell attachment24, we attempted to attach living Hep G2 (human liver cancer) cells to a positively charged hydrogel microfiber. We herein reveal that the developed microfibers undergo attachment of the cells owing to their positively charged surfaces. Furthermore, after attachment, the living cell density can be increased by supplying nutrients.


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2. MATERIALS AND METHODS 2.1 Materials N-isopropylacrylamide (NIPAm) was purchased from Tokyo Chemical Industry (Nihonbashi-Honcho, Tokyo, Japan). N,N′-methylenebis(acrylamide) (BisAA) was obtained from Bio Basic Inc. (Markham, Canada). Allylamine (AA) and methacryloxyethyl thiocarbonyl rhodamine B (MTRB) were obtained from Samchun Chemical (Seoul, Korea) and Polysciences (Warrington, PA, USA), respectively. Dulbecco’s Modified Eagle Medium (DMEM), fetal bovine serum (FBS), and penicillin/streptomycin were obtained from Capricorn Scientific (Auf der Lette, Ebsdorfergrund, UT, Germany). All other chemicals, including sodium acrylate (SA) and α-ketoglutaric acid (KGA), were obtained from Sigma-Aldrich (St Louis, MO, USA) and used as received.

2.2 Fabrication of the microfluidic device The microfluidic device used to fabricate the hydrogel microfibers and microtubes consisted of two glass capillaries (World Precision Instruments, USA). Different sized capillaries were used to control the dimensions of the microfibers (outer: 300 µm, inner: 210 µm) and microtubes (outer: 320 µm, inner: 180 µm). To make the surfaces of the capillaries hydrophobic, they were immersed in 0.2% (v/v) octadecyltrichlorosilane/toluene solution for 20 min, washed with excess toluene, and dried at 80 °C for 1 h. The capillaries were then arranged through coaxial alignment on a glass slide (76 mm × 52 mm) and fixed using epoxy glue, as shown in Figure 1a.


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2.3 Preparation of hydrogel microfibers and microtubes Monomer solutions (2.5 mL) containing NIPAm and SA or AA at different molar ratios (summarized in Table 1), BisAA (6.8 mg), and sodium alginate (25 mg) were prepared in deionized water; then KGA (2.5 × 10-2 mg), a photoinitiator, was added to the solution. Calcium chloride solution (3 wt%) was prepared with a 0.01 M Tris buffer in a 0.25 M phosphate buffer. All the prepared solutions were degassed to remove dissolved oxygen. The resulting solutions were injected into the outer and inner capillaries using syringe pumps (Legato 100, KD Scientific, USA) at controlled flow rates. For the preparation of microfibers, the monomer and calcium chloride solutions were injected into the inner and outer capillaries, respectively. The injection positions were swapped for the preparation of the microtubes. When the alginate-templated microfibers or microtubes emerged from the capillary, they were exposed to 365 nm UV radiation generated by an Omnicure S1500 lamp (Lumen Dynamics, Canada). The obtained microfibers or microtubes were then immersed in an ethylenediaminetetraacetic acid (EDTA) solution for 15 min to remove the alginate template.

Table 1. Compositions of the polymerization solutions for poly(N-isopropylacrylamide-cosodium acrylate) and poly(N-isopropylacrylamide-co-allyl amine copolymers Sample

NIPAm : SA (mol%)

NIPAm (mg)

SA (mg)

PNIPAm PNS3 PNS5 PNS7 PNS10

100 : 0 97 : 3 95 : 5 93 : 7 90 : 10

243.23 237.16 233.02 228.89 222.56

0.00 6.08 10.22 14.35 20.68

Sample

NIPAm : AA (mol%)

NIPAm (mg)

AA (mg)

PNA3 PNA5 PNA7 PNA10

97 : 3 95 : 5 93 : 7 90 : 10

239.58 236.91 234.23 230.34

3.65 6.32 9.00 12.89

2.4 Preparation of a cell-attached hydrogel microfiber with epithelial cells


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Hep G2 (human liver cancer) cells were treated with the EF1alpha 1.4-GFP virus for 14 to 16 h. GFP-positive cells were identified by puromycin selection. Hep G2 cells were cultured in DMEM supplemented with 10% FBS and 1% penicillin/streptomycin (100 U/mL) at 37 °C in a humidified incubator with 5% CO2. Hydrogel microfibers were irradiated with UV light for 15 min and then soaked in a medium containing 1.0 × 105 GFP-Hep G2 cells/mL in a non-adherent dish for 24 h. To promote the homogeneous attachment of the GFP-Hep G2 cells to the surfaces of the hydrogel fibers, the dishes were agitated slightly for 3 h. The cell-attached microfibers were then transferred to a new dish and cultured. The cell medium was changed every 2–3 days.

2.5 Measurements Fourier-transform infrared (FT-IR) spectroscopy was performed on an FT/IR-4600 spectrometer (JASCO, USA). The diameters of the microfibers and microtubes were determined using an optical microscope (OM; DMI-3000B, Leica, Germany). A heating system (MATS, Leica, Germany) was used to control the temperature of the swelling media. Three-dimensional images of the microtubes were obtained using confocal laser scanning microscopy (CLSM; LSM-700, Carl Zeiss, Germany) in Z-stack mode. The microfiber-attached cells were observed using a fluorescence microscope (AE30, Scientific Instrument Company, USA). The fluorescent area was determined using Image J software. (NIH, USA).


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3. RESULTS AND DISCUSSION

Figure 1. Schematic diagrams of (a) the microfluidic device and the method used to fabricate hydrogel microfibers. (b) Chemical structures of poly(N-isopropylacrylamide-co-sodium acrylate) (PNS) and poly(N-isopropylacrylamide-co-allyl amine) (PNA). (c) Fabrication of microtubes or microfibers depending on the injection positions of the monomer solution and calcium chloride. Figure 1a illustrates the microfluidic device architecture, which comprises coaxially aligned glass capillaries connected to syringe pumps. To fabricate the Ca-alginate template for the microfiber using the microfluidic device, the monomer and calcium chloride solutions are injected at constant flow rates through different capillary inlets. The monomer solution, which is an aqueous mixture of sodium alginate (Na-alginate), hydrogel monomers, crosslinkers, and the photoinitiator, is injected into the inner capillary through inlet (I). The pH-responsive hydrogels are prepared by incorporating ionic pendant groups on the hydrogel backbone, which can be


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achieved by the copolymerization of NIPAm with the acrylate-functionalized monomers SA or AA that provide carboxyl and amino groups, respectively. Various comonomer solutions containing different amounts of PNIPAm and SA or AA (summarized in Table 1) were prepared and injected into the inlet of the microfluidic device. Sodium alginate is a natural anionic polysaccharide composed of α-L-guluronic acid and β-D-mannuronic acid that is generally extracted from brown algae. When aqueous Na-alginate is added to the calcium chloride solution (II), the water-insoluble calcium alginate (Ca-alginate) is immediately formed through a divalent salt bridge between the alginate polymers. The calcium chloride solution (3.0% calcium chloride dissolved in 0.01 M Tris buffer) is injected into the outer capillary and the coaxial flows of the two fluids merge at the junction of the microcapillaries. At this moment, the calcium ions in the outer fluid stream diffuse into the monomer solution of the inner fluid stream, thus forming a crosslinked Ca-alginate template through rapid ionic bridge formation between the alginate backbones and calcium ions. Continuous flow induces the progressive formation of the solidified Ca-alginate, generating a Ca-alginate template in the shape of the microfibers. After fabricating the Ca-alginate templates containing the hydrogel monomer, crosslinker, and photoinitiator, crosslinked hydrogel microfibers can be generated by photopolymerization under UV irradiation (III). The Ca-alginate template can be selectively removed upon incubating in 0.1 M EDTA dissolved in 0.25 M phosphate buffer solution through chelation of the calcium ions by EDTA, yielding pure copolymer hydrogel microfibers15. Through the procedure described above, PNS and PNA microfibers of various chemical compositions can be prepared Figure 1b. A series of PNS and PNA copolymers were synthesized with 3, 5, 7, and 10 mol% SA or AA with respect to the total


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number of monomers, and they are designated as PNSn or PNAn, where n denotes the molar ratio of SA or AA employed, as summarized in Table 1. The fabricated microfluidic device can be used to prepare hydrogel microtubes as shown in Figure 1c by swapping the injection positions of the solutions. To fabricate the microtubes, the monomer solution is injected into the outer capillary, while the CaCl2 solution is injected into the inner capillary. When these solutions come into contact during coaxial flow, Ca2+ ions diffuse into the monomer solution and form tubular alginate templates. After photopolymerization of the monomers and selective removal of the alginate with EDTA, pure hydrogel microtubes are obtained.

Figure 2. Photographs and micrographs of PNS10 (a) microfibers and (b) microtubes after


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EDTA treatment. (c) CLSM images in Z-stack mode and a reconstructed three-dimensional image of a PNS10 microtube. (d) FT-IR spectra of PNS and PNA with different SA or AA content. The formation of hydrogel microfibers or microtubes following the extraction of the alginate was confirmed by observing their shape. As shown in Figure 2a, PNS10 microfibers are long and continuous fibers with uniform diameters. The diameter of the hydrogel fiber was determined from the optical micrograph to be 220 µm, which is comparable to that of the inner glass capillary (210 µm). This means that the dimensions of the hydrogel microfibers can be adjusted by varying the dimensions of the glass capillary for the monomer solution. Figure 2b shows OM and CLSM images of the PNS10 microtubes. As with the microfibers, long and continuous PNS10 microtubes with uniform diameters and no twisting or entanglement can be prepared. The diameter of the microtubes was found to be 320 µm, which corresponds the diameter of the outer glass capillary (320 µm) used in their fabrication. The structure of the microtubes was investigated by CLSM measurements. Hydrogel microtubes can be visualized in CLSM by incorporating fluorescent MTRB in the hydrogel backbone15. Thus, a small amount of MTRB was dissolved in the monomer solution and copolymerized with the other monomers upon photopolymerization. As shown in Figure 2c, the microtubes generated are hollow with a uniform wall thickness. The wall of the PNS10 microtube appears fluorescent red due to MTRB while the empty core appears dark. The wall thickness was found to be 70 µm, which corresponds to half the difference between the sizes of the outer capillary (320 µm) and the inner capillary (180 µm). These results demonstrate that the diameter and wall thickness of the microtubes can be controlled by varying the dimensions of the glass capillaries. The reconstructed three-


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dimensional CLSM image shown on the right side of Figure 2c further confirms the tubular structure of the generated hydrogel microtubes. FT-IR measurements were performed to confirm the incorporation of SA or AA into the PNIPAm25-26. As can be seen in the FT-IR spectra in Figure 2d, for the PNS3 microfiber, a characteristic peak at 1750 cm-1, which corresponds to the carboxylate anion of the SA moiety, is observed, while the spectrum of pure PNIPAm is featureless. Furthermore, the intensity of this SA peak increases as the SA content increases, indicating that the SA is copolymerized with the PNIPAm. For the PNA microfibers, characteristic N-H stretching by the primary amine group is observed as two peaks at 3100 and 3120 cm-1. The pure PNIPAm shows only a broad peak near 3100 cm-1, which is attributed to the secondary amide group, confirming that AA is incorporated into the microfibers. The two peaks become more clearly distinguished and sharper, with the peak at 3100 cm-1 increasing in intensity, as the AA content increases. Thus, we can confirm that the copolymerization of both PNA and PNS proceeded successfully.


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Figure 3. (a) pH-dependent diameter changes for PNS10. (b) DpH 3/DpH 7 for PNS and DpH 11/DpH 7

for PNA vs. comonomer content. (DpH n: diameter of the microfiber at pH n) To assess the pH-responsiveness of the prepared microfibers and microtubes, the pH of

the aqueous medium was controlled by adding small quantities of 0.01 M HCl or 0.01 M NaOH. The diameter changes for PNS10 microfibers as a function of the pH measured at 25 °C are shown in Figure 3a. Similar diameters are observed for the PNS microfibers in the pH range 5.0– 11.0. The diameter of the PNS microfibers dramatically decreases near pH 4.75, which is the pKa value of the carboxyl group of SA. The diameter of the PNS10 microfibers decreases to 147 µm at pH 3.0, which corresponds to a 33.6% decrease, i.e., a 71% volume decrease, compared to the equilibrium diameter of 222 µm at pH 11.0.


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At pH values above the pKa, the carboxyl groups are dissociated into the carboxylate groups, leading to swelling of the microfibers owing to the increased hydrophilicity and electrostatic repulsion between the ionized groups27. When the pH is lowered below the pKa, the carboxylate groups are protonated, inducing shrinkage in the microfibers owing to the decrease in electrostatic repulsion. For the PNA microfibers, we also observed a similar pH-dependent size change near the pKa of the amine group. From the graph, we can see that when the pH increased from 3.0 to 11.0, the diameter of the microfibers remains largely unchanged. However, a significant decrease in diameter occurred when the pH increased to 11.0. The diameter of PNA10 microfibers was 222 µm at pH 3.0 and that decreased to 172 µm which corresponds to 22.5% and 53.4% of diameter and volume decrease respectively. Since the amine groups incorporated into the copolymer have a pKa value of 9.69, they are deprotonated in basic conditions, inducing an increase in diameter owing to the electrostatic repulsion between the ammonium cations28. These pH-dependent diameter changes are linearly proportional to the SA or AA content of the copolymer microfibers, as shown in Figure 3b. To normalize the degree of change, the measured diameters of the microfibers at pH 3.0 or 11.0 were divided by those at pH 7.0. For the PNS3 microfibers, the diameter decreases by 11.7% with a change in the pH of the medium from 7.0 to 3.0, while the PNS7 microfibers show a diameter decrease of 24% under identical conditions. The PNA microfibers show the same trend in diameter change as that observed for the PNS microfibers, i.e., the degree of change is proportional to the AA content. The diameters of PNA3 and PNA7 decrease by 6.8% and 16%, respectively, when immersed in a medium with a pH of 11.0. For PNA10 microfiber, diameter of 222 µm at pH 3.0 decrease to 172 µm at pH 11.0,


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which corresponds to 22.5% and 53.4% of decrease in diameter and volume, respectively. The pH-dependent diameter change of the PNA copolymer microfibers is attributed to the degree of protonation/deprotonation of the AA. Therefore, a higher AA content in the PNA microfibers results in more protonated ammonium groups when the pH is below the pKa, leading to a higher degree of diameter change.

Figure 4. (a) pH-dependent diameter and wall thickness changes for PNS10 microtubes. (b) DpH 3/DpH 7

and WpH 3/WpH 7 for PNS vs. comonomer content (WpH n: wall thickness at pH n). (c)

Diameter and wall thickness of PNA10 microtubes as a function of pH. (d) DpH 11/DpH 7 and WpH 11/WpH 7

for PNA vs. comonomer content.

The pH-dependent structural changes of the hydrogel microtubes were investigated by CLSM in Z-stack mode. From the CLSM images, the diameter and wall thickness of the microtubes can be determined. As shown in Figure 4a, for the PNS10 microtubes, there was no


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significant change in diameter or wall thickness in the pH range 5.0–11.0. Since the carboxyl group of SA maintains its negatively charged state in this pH range, electrostatic repulsion between these groups induced swelling of the hydrogel. A drastic decrease in the diameter and wall thickness was observed in the pH range 3.0–5.0 owing to the protonation of the carboxylate group. The outer diameter of 326 µm and wall thickness 77 µm at pH 5.0 for the PNS10 microtubes decreased to 226 µm and 52 µm, respectively, at pH 3.0. We prepared a series of PNS microtubes with different SA content but similar diameters (~325 µm) and wall thicknesses (~80 µm) at pH 7.0. Upon swelling at pH 3.0, the degree of diameter change varies depending on the SA content of the hydrogel. To gain a better understanding of the influence of SA content on this pH-responsive behavior, fluorescence intensity profiles of the microtubes were obtained at pH 3.0 and 25 °C and plotted. Other PNS microfibers with different SA content show the same pH-dependent dimension changes, albeit to different degrees. As illustrated by the plot in Figure 4b, microtubes with lower SA content show a lower degree of dimension change. WpH n represents the wall thickness in the swollen state at pH n. The degree of dimension change was normalized by dividing the diameter and wall thickness at pH 3.0 (DpH 3 and WpH 3) by DpH 7 and WpH 7, respectively. The PNS3 microtubes undergo an 8.6% diameter decrease and a 9.7% wall thickness decrease upon changing the pH from 7.0 to 3.0, while the same pH change for PNS5 results in a 22% diameter decrease and 23% wall thickness decrease. Thus, the degree of dimension change in the microtubes increases with increasing SA content. Figure 4c shows the pH-dependent diameter and wall thickness changes for the PNA10 microtubes. Within the pH range 3.0–9.0, the diameter and wall thickness of the microtubes remain constant as there is no change in the positively charged ammonium groups. However, as


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the pH increased to 11.0, a dramatic decrease in the dimensions of the PNA10 microtubes is observed, caused by the deprotonation of the ammonium groups. The PNA10 microtubes have a diameter of 325 µm and wall thickness 80 µm at pH 7.0 that decreased to 245 µm and 60 µm, corresponding to a 24% and 26% reduction, respectively. The degree of pH-dependent dimensional decrease of the microtubes was also investigated, and the results are plotted in Figure 4d. Similar to the trend for the PNS microtubes, DpH 11/DpH 7 and WpH 11/WpH 7 are linearly proportional to the AA content of the copolymers. The degrees of diameter and wall thickness change for the PNA10 microtubes are approximately four-times larger than those for PNA3. As discussed above in the section on PNA microfibers, since the deprotonation of ammonium groups leads to a decrease in diameter and wall thickness, a higher degree of deprotonation (i.e., higher AA content) causes a more significant dimensional change in the PNA microtubes.


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Figure 5. (a) Normalized temperature-dependent diameter 〈〉 changes for PNIPAm, PNS10, and PNA10. (b) Shrinkages upon heating from 25 °C to 38 °C vs. comonomer content. (c) A series of CLSM images for PNS10 observed at different temperatures. (d) Diameter and wall thickness changes vs. temperature for PNS10 and PNA10 microtubes. (e) The temperaturedependent changes of diameter and wall thickness for PNS10 and PNA10 microfibers.


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Crosslinked PNIPAm is a typical temperature-responsive hydrogel that exhibits reversible volume phase transition depending on temperature29. Near its LCST of 32 °C, PNIPAm shows phase transition from a swollen hydrated state to a dehydrated state, and thereby its volume decreases. Thus, we expected the PNIPAm-based microfibers and microtubes prepared in this study to exhibit thermoresponsiveness as well as pH response. The temperature-dependent volume changes of the microfibers and microtubes were investigated by observing the changes in their shapes and sizes with increasing temperature from 24 to 38 °C. To measure the equilibrium size at each temperature, the hydrogel microfibers were held for at least 10 min at the new temperature after increasing it by 1 or 2 °C. The PNS and PNA microfibers show very similar thermal behaviors to that of PNIPAm microfibers, except for the degree of volume change Figure 5a. The dimeters of the PNS10 and PNA10 microfibers were found to be 222 and 220 µm, respectively, at 25 °C. The diameters of these copolymer microfibers decrease with increasing temperature, and they exhibit a dramatic volume decrease near to 32 °C. Owing to the volume phase transition, decreases in diameter to 126 and 134 µm are observed for PNS10 and PNA10, respectively, at 38 °C. The diameter of the PNIPAm microfibers was measured to be 220 µm at 25 °C and 76 µm at 38 °C, corresponding to a 65% decrease. The degree of volume change decreases with increasing SA or AA content of the copolymer. Figure 5b shows the shrinkage of the PNS and PNA hydrogel microfibers, which was determined by dividing the measured diameter at 38 °C by that at 25 °C, with the comonomer content. An increase in SA content leads to a decrease in the temperature-dependent diameter shrinkage. Increasing the temperature to 38 °C induces shrinkages in the PNS microfibers ranging from 43% to 59% depending on SA content, while the PNIPAm microfibers show a 63%


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shrinkage for the same temperature change. For the PNA microfibers, temperature-dependent shrinkage also disproportionally decreases with increasing AA content. The PNA3 microfibers show a shrinkage of 58% at 38 °C, while the shrinkages for the PNA5 and PNA7 microfibers are 51% and 43%, respectively. This indicates that the shrinkage of the microfibers is affected by the PNIPAm content, which is the temperature-responsive component of the copolymer hydrogels. Temperature-responsive volume phase transitions for the PNS and PNA microtubes were monitored by CLSM in Z-stack mode during heating of the medium from 25 °C to 38 °C. The representative CLSM images measured at different temperatures shown in Figure 5c reveal that both the diameter and wall thickness of the PNS10 microtubes decrease with increasing temperature. To quantify these changes, we extracted the fluorescence intensity profile for the microtubes from the obtained CLSM images, enabling the numerical determination of the diameter and wall thickness of the microtubes, as shown in Figure 5d. Similar to the microfibers described above, the temperature-dependent dimensions of the microtubes show LCST behavior. To further analyze the thermal behavior of the microtubes, the determined diameters and wall thicknesses were plotted as a function of temperature in Figure 5e. The diameter and wall thickness of the PNS10 microtubes at 25 °C are 327 and 78 µm, and they decrease to 165 and 34 µm, respectively. A dramatic decrease in dimensions is observed near 32–34 °C, which corresponds to the LCST of PNIPAm. As can be seen in Figure 5e, the PNA10 microtubes exhibit very similar thermal behavior to that of PNS10. We also found that other PNS and PNA microtubes with different SA or AA content show the same thermoresponsive dimension changes, but to a degree that is proportional to the NIPAm content of the hydrogel. As the AA or SA content of the hydrogel increases, the degree of dimension change decreases. While the PNS7


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microtubes show a 54% decrease in diameter and a 61% decrease in wall thickness, those for PNS3 are 64% and 70%, respectively. These analyses demonstrate that the hydrogel microfibers and microtubes developed in this study undergo volume phase transitions in response to both temperature and pH changes, indicating that their dimensions can be tailored by controlling both temperature and pH. Of particular interest is the fact that the inner diameter of the microtubes changes in response to multiple stimuli, suggesting their potential application as a means of environment-dependent flow control.

Figure 6. Reversible and repeatable diameter changes over 20 cycles for (a) PNS10 and (b) PNA10 microfibers in response to pH (black) and temperature (red).


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Furthermore, these stimuli-responsive changes in dimensions are reversible and repeatable without variation. The equilibrium diameter of PNS10 microfibers is 220 µm at pH 11.0 and 25 °C, while the diameter at pH 3.0 and 25 °C decreases to 136 µm, as shown in Figure 6a. This pH-dependent volume change at constant temperature is found to be fully reversible, i.e., when the pH value is returned to 11.0, the diameter of the PNS10 microfibers is recovered to its original value. This reversible size change can be repeated over 20 cycles without significant variation. In order to assess the reversibility and repeatability of the temperature response of the microfibers, the temperature of the medium was varied between 25 and 38 °C under fixed-pH conditions (pH = 11.0). The red symbols in Figure 6a represent the temperature-dependent diameter changes for PNS10 microfibers recorded over 20 heating and cooling cycles. The diameters of the microfibers vary uniformly between ~220 µm at 25 °C and ~125 µm at 38 °C, demonstrating that these changes are fully reversible and repeatable. These tests were also repeated for PNA10 microfibers, and the results are plotted in Figure 6b. Thus, we can conclude that the pH- and temperature-induced volume phase transitions for both PNS and PNA are fully reversible and repeatable. Furthermore, the PNS and PNA microtubes exhibit the same reliable multi-responsive volume change. This demonstrates that the developed PNS and PNA hydrogels are chemically stable over several pH and temperature cycles, verifying the potential of the system for application to multi-stimuli responsive materials. In order to explore the potential of the developed hydrogel microfibers for application in tissue engineering, we assessed their cell-attachment properties. Motivated by a previous report that described the efficient interaction between cells and positively charged hydrogel surfaces24, we employed positive ammonium-ion-containing microfibers for cell attachment tests. We


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initially attempted to perform the cell-attachment tests with PNA microfibers. However, the large volume increase upon warming from room temperature to the temperature of the cell culture (37 °C) resulted in the detachment of the cells from the microfiber surfaces. Accordingly, to increase the LCST of the hydrogel above the cell-culture temperature, 15 mol% vinyl-2-pyrrolidinone (VP) was incorporated into the PNA hydrogel network. Hydrophilic VP near the polymer backbone inhibits aggregation of the hydrogel network, increasing the LCST18,30. The resulting poly(N-isopropylacrylamide-co-vinyl-2-pyrrolidinone-co-allyl

amine)

(PNVA)

microfibers

undergo volume phase transition at ~40 °C.

Figure 7. (a) Optical and fluorescence images and fluorescence profiles of PNV, PNVA5, and PNVA10 microfibers following Hep G2 cell-attachment tests. (b) Results of cell-growth tests conducted with PNVA10 microfibers.


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We designed and performed a cell-attachment test using Hep G2 cells on PNVA microfibers. The Hep G2 cells were seeded in non-adherent dishes containing UV-sterilized poly(N-isopropylacrylamide-co-vinyl-2-pyrrolidinone) (PNV), PNVA3, PNVA5, or PNVA10 microfibers. The PNV microfibers were included as a control sample. The microfiber samples were extracted and observed using OM following 24 h in the cell culture medium at pH 7.5. As shown in Figure 7a, Hep G2 cells are attached to the PNVA5 and PNVA10 microfibers, while no cells are detected on the PNV microfiber surface. To quantitatively evaluate the cell attachment to the microfibers, fluorescent images were obtained using cells treated with green fluorescent protein (GFP)31. The GFP vector transfected into Hep G2 cells via a lentivirus recombines with the DNA of the Hep G2 cells. Consequently, the green fluorescence is only expressed by living cells. Since only the live cells on the microfibers appear fluorescent green under fluorescence microscopy, we can quantify the degree of cell-attachment on the microfibers by reference to the fluorescent area in the obtained images. The fluorescent areas for the PNVA5 and PNVA10 microfibers were found to be 2.2 × 103 and 5.9 × 103, respectively, meaning that more cells attached to the PNVA10 microfibers. As expected from the OM image, the fluorescent area for the PNIPAm microfibers was negligible. The electrostatic attraction between the negatively charged cell membranes and positively charged microfiber surface promotes cell attachment to the PNVA microfibers. Therefore, PNVA copolymers with higher AA content show larger fluorescent areas. We cultured these attached cells to determine whether the hydrogel microfibers developed in this study provide suitable microenvironments for cells. The cell-attached PNVA10 microfibers were further cultured with the addition of fresh media every 2–3 days to supply nutrients. As shown in Figure 7b, the number of attached cells increases as the incubation time


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increases, and the microfiber surface is eventually fully covered with Hep G2 cells. The initial fluorescent area of 5.9 × 103 increases to 14.6 × 103 by 7 days and to 19.1 × 103 by 14 days, corresponding to a threefold increase. In this cell culture test, we only supplied nutrients to the cells, and no further Hep G2 cells were added. Thus, the increase in cell number resulted solely from the growth of the cells on the microfibers. These results clearly indicate that the positively charged PNVA microfibers undergo attachment to negatively charged cell membranes via ionic bonding, meaning that these microfibers could be used for three-dimensional cell-culture systems. We propose that the PNVA microfibers developed in this study can be applied as a microfiber platform for cell attachment and growth in tissue engineering.

4. CONCLUSIONS In conclusion, we fabricated pH- and temperature-responsive hydrogel microfibers and microtubes using a microfluidic device by alginate-templated photopolymerization of vinyl monomers. CaCl2 and monomer solutions containing Na-alginate were coaxially flowed through micro-capillaries, forming uniform microfibers or microtubes depending on the injection positions of the solutions. Hydrogel microfibers or microtubes were obtained upon removing the Ca-alginate template by EDTA chelation. To prepare pH- and temperature-responsive micro-objects, pendant carboxyl or amine groups were introduced into the hydrogel networks by the copolymerization of NIPAm with SA or AA. The volume phase transition of the microfibers and microtubes can be triggered by pH as well as temperature. The ionic groups in the hydrogels are protonated or deprotonated in acidic or basic media, respectively, inducing volume changes in the hydrogels due to the repulsive force between the charged groups. The temperature-responsive changes observed originate from the


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PNIPAm, which is the dominant component of the hydrogels. Furthermore, we have demonstrated that these volume changes are fully reversible and repeatable. The enhanced cell attachment to the surfaces of the charged hydrogel microfibers demonstrated their potential for application in tissue engineering. The negative charge of the cell membrane allowed Hep G2 cells to attach to the positively charged surface of the microfibers through ionic attraction. Furthermore, we found that the number of cells attached to the microfibers could be increased by supplying nutrients, suggesting that the multi-stimuliresponsive hydrogel microfibers and microtubes developed in this study can be applied to tissue engineering and regenerative medicine.

AUTHOR INFORMATION Corresponding Author * E-mail: (J.Y.) [email protected], (J-W.C.) [email protected] Author contribution J.Y. and J.C. planed and supervised the project. Dongwan. K. and A.J. carried out bulk of experiments and analyzed biological data. K.B.C.I. and Dowan. K. analyzed the spectroscopic data. All authors contributed to discussion of the results and wrote the paper. Notes The authors declare no competing financial interest.


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ACKNOWLEDGMENTS This research was supported by the Ministry of Science and ICT (MSIT) and the National Research Foundation of Korea (NRF) through the Mid-Career Researcher Fund (Strategy Research)

(2016R1E1A1A01942509)

and

Creative

Materials

Discovery

Program

(2017M3D1A1039287)


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Table of Contents Graphic


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Microfluidic fabrication of multi-stimuli responsive tubular hydrogels for

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