For this work, 20 g of graphite (Synthetic graphite powder <20 μm, Aldrich Chemistry, St. Louis, MO) and 650 mg of BSA (A7906, Sigma-Aldrich, St. Louis, MO), were mixed in a kitchen blender with 200 mL of DI water for 25 min. The graphene solution was allowed to rest for ~24 h to allow for remaining graphite to settle out of the solution. The supernatant was used as the graphene solution.

To prepare the core solutions, 0.1 g of alginate powder (very low viscosity, Product Number A18565 Alfa Aesar, Ward Hill, MA) and a magnetic stirrer were soaked overnight in ethanol under UV light from a biological fume hood. Once dry, 1.8 mL of WFI-Quality Cell Culture grade water (Corning, Corning, NY) and 1.8 mL of freshly mixed UV-sterilized graphene solution were introduce. The resulting solutions were mixed with 0.2 mL of cell suspension (1.1725 × 10⁷ cells mL⁻¹) to form a 5% alginate solution. Various other concentrations of alginate were tried, but a 5% alginate solution was viscous enough to resist shear force within the microfluidic channel, thereby producing smoother fibers.

The sheath solution was created by dissolving 20% poly(ethylene glycol) (PEG) (M_n = 20,000, Aldrich Chemistry, St. Louis, MO) and 0.04% CaCl₂·2H₂O (Fisher Chemical, Waltham, MA) into DI water. The collection bath was 7.5% PEG, 2.5% CaCl₂·2H₂O. Both solutions were sterilized using a 0.22 μm sterile syringe filter.

Cell-laden fibers were introduced into the collection bath, where they sank before they were removed with tweezers and were wrapped around polydimethylsiloxane (PDMS) frames. Frames were fabricated by introducing 0.5 g of mixed PDMS into 12-well plates and thermosetting them at room temperature for 1 day. Once solidified, two sides of the circle were removed and the center was removed with a hole punch.

The microfluidic devices used in this study are fabricated by thermosetting polydimethyl sulfoxide (PDMS) onto photolithographic molds on silicon wafers. The design of the molds has been discussed previously (Sharifi et al., 2016a,b,c; McNamara et al., 2019b; Sharifi et al., 2019). Briefly, the channel had dimensions of 130 μm × 390 μm. Four diagonal grooves on the top and bottom of the channel allow for further shaping of the core fluid; these chevrons are 200 μm apart and have dimensions of 130 μm × 100 μm. The main chamber of this device was 1 mm longer than previously used to ensure sufficient time within the microfluidic device for solidification of the core solution (Sharifi et al., 2016c).

To create the device, PDMS (Dow Corning, Midland, MI) was mixed in a 1:10 ratio of elastomer curing agent to elastomer base. Mixed PDMS was solidified on the molds at 80°C for 25 min. The two halves were bonded using plasma cleaning.

Rat dopaminergic neural cells (N27s) were received as a generous gift from the Department of Biomedical Sciences at Iowa State University. Cells were cultured in maintenance media (MM) containing RPMI Medium 1640 (1X) (Gibco Life Technologies Limited, Paisley, UK), which was supplemented with 10% fetal bovine serum (FBS) (One Shot format, ThermoFisher Scientific, Waltham, MA), 1% penicillin (10,000 U mL⁻¹)-streptomycin (10,000 μg mL⁻¹) (Gibco, Waltham, MA), and 1% L-glutamine 200 mM (100X) (Gibco Life Technologies Corporation, Grand Island, NY).

Cells were cultured in T-25 flasks and were passaged at 70% confluency and were maintained in a 37°C, 5% CO₂ atmosphere. Cells were passaged three times before use. For encapsulation, cells were trypsinized and 0.2 mL of cell suspension (1.1725 × 10⁷ cells mL⁻¹) was added to the alginate and graphene-alginate solutions.

Before use, microfluidic devices were flushed with 70% ethanol and were placed under UV light for a minimum of 5 h. Sterilized solutions were placed into pre-sterilized BD syringes under a biological fume hood. The solutions were introduced into the microchannel via a double syringe pump (Cole-Parmer, Veron Hills, IL) with a FRR of 40:10 μL min⁻¹:μL min⁻¹ (sheath:core), which was optimized to provide sufficient time within the microfluidic device for gelation of the core solution without causing clogging. The microfluidic schematic can be seen in Figure 1.

To study the porosities of the microfibers, a minimum of n = 3 fibers were fabricated directly onto u-shaped copper wire frames. Fibers were allowed to dry overnight, and then were weighed. Afterwards, fibers were soaked in DI water overnight, and their wet weights were measured after excess liquid was removed by blotting the surface of the fibers with a Kimtech wipe. The wet volume was approximated using the diameter on as measured by a Zeiss Axio Observer Z1 Inverted Microscope. The porosities were then calculated using the Equation (1), where M_w and M_d are the wet and dry weights of the fibers, V is the volume of the wet fiber, and ρ is the density of the liquid:

Cell-free fibers were fabricated in a non-sterile environment and were mounted on paper frames to dry. Once dry, an average of three fibers were mounted on plastic using electrically conductive carbon tape to allow for an electronic connection between an electrode and the fibers. Colloidal silver paste was placed over fiber to ensure ohmic contact. Since dry fibers are straight, the resistance of the fibers was measured using a Sinometer MS8269 Digital Multimeter (ShenZhen, China), and the conductivity was calculated with Equation (2), where Ω is the conductivity, L is the length of the fiber, R is the resistance of the fiber, and A is the cross-sectional of area of the fiber. Fibers were modeled as cylinders, with radii calculated from the wide and narrow size of dry fibers (Lu, 2016).

To perform the live-dead cell assays, a 10 μM CellTracker™ CMFDA and 8 μM propidium iodide (PI) (Invitrogen, Carlsbad, CA) was used as dye solution. On the desired time point, MM was carefully removed from the 12 well plate, and the wells were rinsed with FBS-free RPMI media (500 μL). After, dye media (500 μL) was introduced and was incubated for 20 min at 37°C and 5% CO₂. Once incubation was complete, dye media was removed and fibers were suspended into FBS-free RPMI media (500 μL) to keep samples wet during imaging.

Fluorescent images were captured using a Zeiss Axio Observer Z1 Inverted Microscope. Initial processing was carried out with AxioVision Special Edition 64-bit software, but further editing was completed in Adobe Photoshop CC 2018.

SEM images were generated by drying samples overnight and mounting them using electrically conductive carbon and copper tape. Samples were analyzed using a JCM-6000 NeoScope Benchtop SEM with an accelerating voltage of 15 kV.

Statistical analysis was carried out with R Project Statistical Software to conduct an Analysis of Variance (ANOVA) to compare the means across samples.

Both 5% alginate and 9% graphene, 5% alginate microfibers were successfully fabricated utilizing a sheath solution of 0.04% CaCl₂·2H₂O, 20% PEG. Samples were gathered in a collection bath of 7.5% PEG, 2.5% CaCl₂·2H₂O. Since samples are oval in shape, the analysis was carried out on both a long (wide) and short (narrow) axis (Figure 2). Cross-sectional and longitudinal images of both alginate (a₁-a₂) and graphene-alginate (c₁-c₂) can also be observed in Figure 2. After SEM analysis, it was determined that there was no significant difference between the sizes of the alginate and graphene-alginate samples, as is evidenced by Figure 2b. This is to be expected, since graphene flakes generated by the kitchen blender method small enough in size that they do not drastically change the behavior of the alginate as it passes through the microfluidic device. Previous works have shown that the sizes of microfluidically-generated microfibers can be successfully tuned by adjusting the concentrations of the core fluid, the sheath fluid, and the collection bath; the FRR used to fabricate fibers also plays a significant role in determining their size (Sharifi et al., 2016a,b; McNamara et al., 2017, 2019b).

The porosity of hydrogels used for cell encapsulation plays a significant role in long-term cell health and viability, as it affects the diffusion of nutrients into and waste out of the fiber boundaries (McNamara et al., 2017) In order to investigate the porosities of alginate and graphene-alginate microfibers, fibers were fabricated onto copper frames, which allowed for the measurement of wet and dry weights. There were no significant differences between the porosities of the alginate and graphene-alginate samples (Figure 3). This is to be expected, particularly since graphene particles will not interact strongly with water molecules. Fibers of both samples exhibited high porosities; since excess DI water was removed from the surface of the fibers, this indicates that they are capable of absorbing close their own weight in water, as is common in hydrogels (McNamara et al., 2017).

Aqueous solutions of graphene drastically changed the appearance of microfibers, causing them to be black instead of transparent when wet. As observed in Figures 2a₂,c₂, graphene was well-distributed throughout the body of the fibers. Introduction of graphene into alginate microfibers significantly increased the conductivity of the resulting hydrogels by a factor of 2.5, achieving an overall conductivity of 0.0025 S m⁻¹ for dry fibers mounted using carbon tape and colloidal silver paste (Figure 4). This indicates that the graphene flakes within the fibers are consistently dispersed enough to shorten diffusion distance of electrons, thereby aiding in the transportation of electrons through the bulk of the fiber and increasing its electrical conductivity (Wu and Zhong, 2019) Other cell-laden conductive hydrogels, which have thus far taken the form of films, achieved conductivities of 0.2 S m⁻¹ (Dong et al., 2016) and 0.02 S m⁻¹ (Wang et al., 2018) for wet samples. These films are based on synthetic polymers and were biocompatible, but lacked the high degree of control over cell culture geometry that microfibers can provide. In comparison, the conductivity of native human brain tissue ranges from 0.05 to 0.24 S m⁻¹, with an average of 0.12 S m⁻¹ (Akhtari et al., 2016) link Methods to increase the conductivity of the alginate microfibers to better match the conductivity of brain tissues might include incorporating synthetic polymers or increasing the concentration and dispersion of graphene flakes in an aqueous solution by further refining the lab-based process.

Cells were successfully encapsulated within alginate and graphene-alginate hydrogels, and survived for up to 8 days, as can be seen in Figure 5. Initially, alginate fibers held significantly more live cells than the corresponding graphene-alginate microfibers, as can be seen in Figures 5a₁,a₂,d. However, by the fourth day (Figures 5b₁,b₂), the differences between the samples were insignificant, which is a trend that held until the eighth day (Figures 5c₁,c₂). Between days 1 and 4, the number of live cells within both microfiber samples dropped significantly. This may be because the conditions within the microfibers are not conducive to long-term cell survival; however, at this time cells were also observed on the bottom of the well plates. Therefore, live cells were capable of migrating beyond the fiber boundary, thereby decreasing the number of live cells within the fibers. However, with modifications to the microfibers, the rate of cell egress can be controlled to suit the desired application. This can be done by increasing the porosities of the microfibers, or by other chemical modifications, such as the inclusion of arginine-glycine-aspartic acid (RGD) (Anderson et al., 2015; Ho et al., 2017; Santos et al., 2019).

Increasing the amount of live cells within fiber boundaries would be beneficial for long-term experiments. This can be accomplished by increasing the cross-linking of the hydrogel fiber, thereby restricting the ability of cells to migrate outside of the fibers. Additionally, chemically functionalizing the alginate hydrogel during gelation can increase the amount of cell interactions, thereby improving the biocompatibility of the microfibers (Anderson et al., 2015; Wei et al., 2017).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.