Microfluidic-Based Cell-Embedded Microgels Using Nonfluorinated

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Microfluidic-Based Cell-Embedded Microgels Using NonFluorinated Oil as a Model of the Gastrointestinal Niche Seyed Ramin Pajoumshariati, Morteza Azizi, Daniel Wesner, Paula G Miller, Michael Shuler, and Alireza Abbaspourrad ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b16916 • Publication Date (Web): 23 Feb 2018 Downloaded from http://pubs.acs.org on February 26, 2018

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ACS Applied Materials & Interfaces

Microfluidic-Based Cell-Embedded Microgels Using Non-Fluorinated Oil as a Model of the Gastrointestinal Niche Seyed Ramin Pajoumshariati1, Morteza Azizi1, Daniel Wesner2, Paula G. Miller3, Michael L. Shuler3, and Alireza Abbaspourrad1∗

6 7 8 9 1

10 11

Department of Food Science, College of Agricultural and Life Sciences, Cornell University, Ithaca, NY, 14853

12 13

2

14 15

3

Department of Biological and Environmental Engineering, College of Engineering, Cornell University, Ithaca, NY, 14853 Department of Biomedical Engineering, College of Engineering, Cornell University, Ithaca, NY, 14853

16 17

∗

Corresponding author: Dr. Alireza Abbaspourrad, Department of Food Science, Cornell University, Ithaca, NY. Zip code: 14853, Email: [email protected]

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ACS Applied Materials & Interfaces 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1

ABSTRACT

2

Microfluidic-based cell encapsulation has promising potential in therapeutic applications. It

3

also provides a unique approach for studying cellular dynamics and interactions, though this

4

concept has not yet been fully explored. No in vitro model currently exists that allows us to study

5

the interaction between crypt cells and Peyer’s patch immune cells due to the difficulty in

6

recreating, with sufficient control, the two different microenvironments in the intestine in which

7

these cell types belong. However, we demonstrate that a microfluidic technique is able to provide

8

such precise control and that these cells can proliferate inside the microgels. Current

9

microfluidic-based cell micro-encapsulation techniques primarily use fluorinated oils. Herein, we

10

study the feasibility and biocompatibility of different non-fluorinated oils for application in

11

gastrointestinal cell-encapsulation and further introduce a model for studying inter-cellular

12

chemical interactions with this approach. Our results demonstrate that cell viability is more

13

affected by solidification and purification processes that occur after droplet formation rather than

14

the oil type used for the carrier phase. Specifically, shorter polymer crosslinking time, and

15

consequently lower cell exposure to the harsh environment (e.g., acidic pH), results in a high cell

16

viability of over 90% within the protected microgels. Using non-fluorinated oils, we propose a

17

model system demonstrating the interplay between crypt and Peyer’s patch cells using this

18

microfluidic approach to separately encapsulate the cells inside distinct alginate/gelatin

19

microgels, which allow for inter-cellular chemical communication. We observed that co-culture

20

of crypt cells alongside Peyer’s patch immune cells improves the growth of healthy organoids

21

inside these microgels, which contain both differentiated and undifferentiated cells over 21 days

22

of co-culture. These results indicate the possibility of using droplet-based microfluidics for

23

culturing organoids to expand their applicability in clinical research.


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1

Keywords: Droplet-based microfluidics, Non-fluorinated oil, Cell-embedded microgels,

2

Intestinal crypts, and Peyer’s patch.

3



Page 4 of 46

1

INTRODUCTION

2

Using microfluidic techniques to encapsulate cells in monodisperse microgels enables the in

3

vivo delivery of cells within a supportive, tunable microenvironment inside the human body for

4

tissue engineering and regenerative medicine applications

5

potential of single-cell-laden microgels, they can also be used as 3D models to investigate the

6

effects of different toxicological, environmental, and physicomechanical dynamics on specific

7

cell types. Such studies are difficult to achieve by conventional 2D in vitro methods, such as

8

trans-epithelial transport models, which culture a line of gut cells (e.g., Caco-2) on transwell

9

inserts 4. However, these 2D models are overly simplistic due to a lack of cell diversity. To date,

10

a number of microgel cell encapsulation techniques have been developed. These approaches vary

11

widely in terms of the microfluidic setup, the separation method, and the dispersed phase (i.e.,

12

the choice of hydrogel precursor, its associated crosslinking approach, and the encapsulated cell

13

type) and carrier phase (i.e., oil type and the surfactant). This variation makes the comparison of

14

microgel cell encapsulation methods difficult.

1-3

. In addition to the therapeutic

15

In order to expand the application of droplet-based microfluidic cell encapsulation in pre-

16

clinical and clinical settings, as well as to reduce the cost of this process, a comparative

17

examination of different materials used in this technique is needed. Cells embedded in microgels

18

reside in a microenvironment that is defined by nanoscale physicomechanical properties,

19

including heterogeneity, elasticity, and interfacial chemistry. Modulating these properties can

20

have a significant effect on the cellular function, proliferation, and differentiation of the

21

embedded cells 5-6.

22

Microgel composition affects both nano and bulk mechanical properties, as well as the

23

surface topography, all of which may affect the long-term viability of the encapsulated cells.


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1

Hydrogels, such as polyethylene glycol, gelatin methacrylate, and alginate, are frequent choices

2

for polymers used to produce cell-embedded microgels. Alginate—which has fast, ion-based

3

crosslinking kinetics—is a polymer used in encapsulation methods, in which crosslinking ions

4

(e.g., Ca+2) must be present in the carrier phase, complexed with chelating agents (e.g.,

5

ethylenediaminetetraacetic acid, EDTA) or solid nanoparticles (e.g., CaCO3) 2-3, 7-8.

6

The choice of carrier phase (i.e., fluorinated vs. non-fluorinated oils) can also affect the

7

viability of encapsulated cells. Fluorinated oils have been largely used as a carrier phase for three

8

reasons: their high gas solubility improves cell viability during the formation process; they

9

possess a low solubility for non-fluorinated molecules, which facilitates droplet formation; and

10

finally, they are chemically compatible with poly(dimethylsiloxane) (PDMS), which is

11

frequently used for microfluidic device fabrication. Furthermore, past studies using fluorinated

12

oil for encapsulation yielded high cell viability, in the 73–90% range

13

properties, there have been many studies performed using fluorinated oils as the carrier phase,

14

and as a result these oils are well-characterized.

1-3

. Due to these desirable

15

However, non-fluorinated oils are less expensive to use, yet they have been studied to a

16

lesser degree. It is difficult to estimate cell viability in approaches using these oils, as studies

17

employing them are limited and highly variable 7-9. To fill this knowledge gap, we systematically

18

investigated the use of different non-fluorinated oils for microfluidic-based microgel formation

19

using the biopolymer alginate in a Ca-EDTA chelating complex. However, previous studies have

20

shown that there is a lack of cell adhesion binding sites in pure alginate

21

drawback for cell encapsulation, we added gelatin to the alginate microgels to improve cell

22

adhesion and proliferation via gelatin’s cell adhesive amino acid motifs 10. Moreover, it has been

23

shown that biodegradable matrices are essential for the formation and differentiation of


10

. To overcome this


11

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1

organoids

. The biodegradability of gelatin, which occurs due to the presence of

2

metalloprotease sites in its structure, can further support the culture of healthy organoids. We

3

further examined the oil biocompatibility, using different human cell lines, such as Caco-2 cells,

4

as well as isolated murine Peyer’s patch immune and intestinal crypt cells.

5

Using microgels to encapsulate cells in our study affords a number of advantages over

6

traditional co-culture methods, which cannot accurately recreate the unique mechanical,

7

chemical, and biological matrix of the in vivo microenvironments of each cell type. Microfluidic-

8

based methods of producing cell-embedded microgels allow the physicomechanical and

9

physicochemical properties of the cell microenvironment to be controlled. This is particularly

10

advantageous for culturing mammalian cells that occupy a specialized niche. For instance, in the

11

intestine, stem cells reside at the base of the crypt niche, while intestinal immune cells (a unique

12

subset of immune cells, including B cells, T cells, dendritic cells, macrophages, and M cells

13

are localized to Peyer’s patch nodules. The crypt niche is composed of several proteins,

14

including laminin, collagen IV, and fibronectin, which play a role in keeping residing cells

15

healthy 11.

12

)

16

To the best of our knowledge, a microfluidic-based approach of co-culturing crypt cells (as a

17

functional unit of the gastrointestinal tract) and Peyer’s patch cells (a functional unit of the

18

immune system) has not yet been reported. Individual and co-cultures of these cell types inside

19

microgels can be used to study the different cells that exist in the crypt-villous domain as well as

20

immune cells that reside in the Peyer’s patch. However, there are limitations to current culture

21

methods for both cell types. For example, culture methods for crypt cells typically use Matrigel

22

13-15

23

Furthermore, Matrigel cannot be used for transplantation into humans, as it is derived from the

, which is not ideal, as the mechanical properties of this material cannot be controlled.


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1

matrix of cancerous cells. The in vitro culture of immune cells is also challenging, as it depends

2

on the replication of complex factors involved in the development and maturation of the immune

3

cells, as well as mobilization of the immune response

4

synthetic environment, like an encapsulating microgel. As an improved method of culturing both

5

cell types, our in vitro co-culture model incorporates distinct cell-embedded microgels in an

6

interconnected 3D-printed mold, which allows for cell signaling and transport. It is worth

7

mentioning that, in many cases, both soluble mediators and direct cell-to-cell contact play a role

8

in the interaction between two cell types. In this case, however, Peyer’s patch cells are

9

anatomically separated from gastrointestinal epithelial cells, and so the exchange of soluble

10

16

—factors that are easier to control in a

mediators in the gastrointestinal tract plays a more dominant role.

11

This brings us back to one of the primary advantages of microfluidics as an encapsulation

12

technique: control of the physicochemical and mechanical properties of the cell

13

microenvironment. Further characterization of these properties, as a function of the composition

14

and production of microgels, contributes to the potential expansion of microfluidic-based models

15

in applied settings.

16 17

EXPERIMENTAL SECTION Materials

18

Sodium alginate (139 kDa, MW), disodium EDTA, gelatin from bovine skin, hexadecane,

19

lecithin, calcium chloride, mineral oil, sorbitan monooleate (Span 80), and poly-L-lysine (70-150

20

kDa, MW) were purchased from Sigma-Aldrich Chemicals (MO, USA). Fluorinated oil HFE-

21

7500 (3M Novec, MN, USA) containing 1 wt% Pico-Surf surfactant (Dolomite Microfluidics,

22

Royston, UK) was used as a control for the carrier phase. Fibronectin and laminin-111 were

23

purchased from Invitrogen (MD, USA) and collagen-IV was obtained from Corning Inc. (NY,



Page 8 of 46

1

USA). Edible oils, including sunflower, corn, grapeseed, olive, and peanut oil, were purchased

2

from a local supermarket. For fabrication of the microfluidic device, SU-8 2050 negative

3

photoresist

4

Polydimethylsiloxane (PDMS, Sylgard 184) and its curing agent were purchased from Dow

5

Corning (MI, USA). Polyethylene tubing (ID = 0.38 mm, OD = 1.09 mm), 27-gauge syringe

6

needles, and 5 ml Luer-Lok tip disposable syringes were purchased from Becton Dickinson (NJ,

7

USA). RPMI 1640, modified Eagle’s medium (MEM), fetal bovine serum (FBS), Dulbecco's

8

phosphate-buffered saline (DPBS), trypsin, penicillin G, streptomycin, live and dead staining

9

(calcein-AM and ethidium homodimer-1 dyes), and trypan blue were purchased from Gibco

10

(Life Technologies, Carlsbad, CA, USA). A nylon cell-strainer (70 µm) for isolation of crypt and

11

Peyer’s patch cells was obtained from BD Falcon Biosciences.

12

and

developer

were

obtained

from

Microchem

Corp.

(MA,

USA).

Analytical instruments

13

The generation of cell-embedded microgels was monitored by an inverted microscope (Leica

14

DM IL LED, Leica Microsystems, Buffalo Grove, IL, USA) equipped with a high-resolution

15

CCD camera (Phantom V2.7, Vision Research, Ametek, Wayne, NJ, USA). ImageJ software

16

(National Institutes of Health, MD, USA) was used to quantify microgel diameters. For each

17

different oil, 500 distinct microgels were analyzed to calculate their average diameter at that

18

condition. The oil viscosities were determined with a falling-ball viscometer (GV-2100, Gilmont

19

Instruments, Barrington, IL, USA). Interfacial tension between water and oil phases was

20

analyzed using a Ramé–Hart Goniometer (Ramé–Hart, NJ, USA). The mechanical properties of

21

the cell-embedded microgels in liquid medium and bulk hydrogels were analyzed by atomic

22

force microscopy (AFM, MFP-3D system, Asylum Research, CA, USA) using silicon nitride

23

cantilevers at a resonant frequency of 22 kHz (MLCT, Bruker AFM Probes, USA) and a


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1

AR1000-N rheometer (TA Instrument, UK) with 40 mm diameter stainless steel parallel plates,

2

respectively.

3

For the co-culture of Peyer’s patch and crypt cells, we produced two concentric

4

interconnected cylinders with inner and outer diameters of 0.25 mm and 0.45 mm, and a height

5

of 10 mm, designed using AutoCAD software (Autodesk). This cell insert was printed using a

6

desktop 3D ObjetPro printer from Stratasys, which prints UV-curable polymers using PolyJet

7

technology. It was then degassed in a vacuum oven at 40 ºC and coated with Parylene C (Model

8

PDS 2010 LABCTER) in order to make it biocompatible.

9

Fabrication of PDMS microfluidic devices

10

A soft photolithography technique was used to fabricate the mold for the PDMS flow-

11

focusing microfluidic device 3, which we used to generate the spherical and monodisperse

12

droplets. Briefly, SU-8 2050 negative photoresist was spun-coated on a silicon wafer (thickness:

13

80 µm) and patterned by UV exposure through a photolithography mask, and then subjected to

14

the baking and developing processes. The mask with flow focusing channels (80 µm width) was

15

designed using AutoCAD. The mixture of PDMS and its curing agent (10:1) was then poured on

16

the SU-8 deposited wafer and baked for 2 h at 65 ºC. After peeling the PDMS off the patterned

17

wafer, injection holes (1 mm in diameter) were punched and cleaned by cellophane tape (3M

18

Scotch Magic, MN, USA), followed by bonding of the PDMS to a glass slide by applying an

19

oxygen-plasma treatment for 1 min. To further stabilize the bonding strength between PDMS and

20

the glass slide, the device was kept at 65 ºC in an oven for 1 h. Hydrophobic treatment of the

21

PDMS channels was performed with Aquapel (PPG Industries, PA, USA).



1

Microgel generation

2

Sodium alginate was dissolved in deionized (DI) water (or PBS) at a concentration of 2

3

w/v%. Calcium chloride (0.1 M) and EDTA-disodium salt (0.1 M) were separately dissolved in

4

DI water. The two solutions were then mixed and the pH was set to 7.2 by the addition of sodium

5

hydroxide. The dispersed phase was composed of sodium alginate solution and the Ca-EDTA

6

complex (1:1). For those samples that contained gelatin, different amounts of gelatin were added

7

to the dispersed phase solution while the concentration of the total hydrogel was kept constant at

8

1 wt%. For the organoid matrix, fibronectin (0.5 mg.ml−1), laminin-111 (0.1 mg.ml−1), and

9

collagen-IV (0.25 mg.ml−1) were mixed with the gelatin-alginate (1:1) mixture. The carrier phase

10

was composed of the non-fluorinated oil containing 2 w/w% lecithin as a surfactant. For cell

11

encapsulation experiments, we used a Caco-2 cell line from the American Type Culture

12

Collection. To solidify droplets, the collected microgels were re-suspended in the carrier phase

13

oil containing acetic acid at several concentrations (≤ 1 v/v%) for a predetermined time and then

14

diluted with hexadecane containing Span 80 (2 w/w%) to dissolve the carrier phase oil and

15

protect the microgels from any oil that remained stuck to the microgels. The diluted mixture was

16

then centrifuged at 60 g for 3 min and the supernatant oil was aspirated. These steps were

17

repeated twice to remove most of the oil. After this step, the microgels were thoroughly collected

18

and washed with PBS to remove the remaining oils and cultured in MEM medium containing

19

10% FBS in a vented T-25 cell culture flask. This protocol using lecithin and Span 80 is a

20

modification of methods used for the generation of cell-embedded microgels developed by Tsuda

21

et al.17 and Tan et al.18. The mean diameter of the microgels was determined from the images

22

captured by an inverted light microscope (50 spheres for each group) using ImageJ software

23

(National Institutes of Health, MD, USA).


Page 10 of 46

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1


Cell culture

2

Caco-2 cell line was obtained from American Type Culture Collection (ATCC, VA, USA).

3

Caco-2 cells were cultured in MEM supplemented with 10% fetal bovine serum (FBS), 100 U/ml

4

penicillin G, and 100 µg/ml streptomycin at pH 7.2 (Life Technologies, Carlsbad, CA, USA) and

5

maintained in a humidified incubator at 5% CO2 and 37 ºC. After reaching 70% confluency, cells

6

were washed with DPBS and detached enzymatically from the flasks using trypsin (0.1%)-

7

EDTA (0.03%). A predetermined number of cells were re-suspended in the medium, then mixed

8

with the alginate-based solution to generate cell-embedded microgels at a specific cell density.

9

Isolation of Peyer’s patch cells and intestinal crypt cells was performed as stated by 19

and Sato et al.

13

10

Lefrançois et al.

, respectively. All animal experimental procedures were

11

approved by the Ethical Committee in Animal Research from Cornell University College of

12

Veterinary Medicine. Peyer’s patch and crypt cells were isolated from C57BL/6J mice. Briefly,

13

animals were killed by exposure to CO2 followed by cervical dislocation. After removal of the

14

intestine, the ileum was cut out using a surgical scissor and flushed with cold PBS several times

15

by a 10 ml syringe. Following a series of washing steps in cold PBS containing 1X penicillin and

16

streptomycin, Peyer’s patch tissues, which are roughly egg-shaped protruding whitish/greyish

17

lymphatic nodules located on the outer intestinal wall, were then dissected, mechanically

18

disrupted by passage through a 70 µm nylon cell-strainer (BD Biosciences), and collected in cold

19

RPMI 1640 media. Prior to encapsulations, cells were centrifuged at 600 g for 10 min at 4 ºC.

20

Crypt cells were isolated from the intestine. After cleaning steps and removing the mucus and

21

intestinal villi using a glass slide, tissues were cut into small pieces and dissociated in 5 mM

22

EDTA solution in PBS by shaking for 40 min on ice. The crypt cells in the supernatant were then



1

passed through a 70 µm nylon cell-strainer (BD Biosciences) and centrifuged at 600 g for 5 min

2

at 4 ºC prior to encapsulation.

3

Cell viability was assessed by trypan blue exclusion and live/dead fluorescent staining using

4

calcein-AM (green) and ethidium homodimer-1 (red) dyes (Gibco, Life Technologies, Carlsbad,

5

CA, USA).

6

Immunofluorescence staining

7

For immunofluorescence and confocal imaging, the cell-embedded microgels were fixed in

8

an aqueous solution of paraformaldehyde (4%) containing 10 mM CaCl2 for 30 min (to prevent

9

destabilization of alginate), then washed with a PBS buffer containing calcium and magnesium

10

(PBS buffer), and permeabilized with 0.25% Triton X-100 for 15 min at room temperature (22 ±

11

2 °C). After washing with the PBS buffer, microgels were blocked with PBS buffer containing

12

5% bovine serum albumin for 1 h at room temperature, then incubated in primary antibodies

13

against villin and Bmi-1 (SC-58897 and SC-390443, respectively, Santa Cruz Biotechnology

14

Inc., TX, USA, 2µg/ml diluted in blocking buffer) for 1 h at room temperature. After washing

15

five times with the PBS buffer, the cell-embedded microgels were incubated with the secondary

16

antibody (SC-516141, Santa Cruz Biotechnology Inc., TX, USA, 1µg/ml diluted in blocking

17

buffer) for 1 h in the dark at room temperature. Following extensive washing with the PBS

18

buffer, the microgels were mounted with a Fluoroshield mounting medium with 4′ 6-diamidino-

19

2-phenylindole (DAPI) (Sigma-Aldrich Chemicals, MO, USA) to stain the nuclei. Stained

20

organoids were imaged using a water immersion 10X objective on a Zeiss confocal microscope

21

(LSM 710, Carl Zeiss, Göttingen, Germany).


Page 12 of 46

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1


Rheological properties of the hydrogels

2

The Young's modulus of the microgels was determined via AFM (MFP-3D system, Asylum

3

Research). To prevent the movement of microgels during AFM measurement, they were

4

electrostatically adhered to a poly-L-lysine-coated glass slide by incubating for 30 min at room

5

temperature 3. AFM measurements were performed inside liquid MEM medium using silicon

6

nitride cantilevers (MLCT, Bruker AFM Probes). To ensure accurate measurement and

7

calibration, the spring constant was determined by performing thermal tuning (at room

8

temperature and far from the surface after reaching equilibrium), followed by correction of force

9

curves and engaging the tip on a clean and hard surface in contact mode 3. The spring constant

10

ranged between 20 to 50 mN m-1. Topographic images were taken in tapping mode while the

11

force curves and maps were determined under contact mode with a cantilever approaching rate of

12

1 µm s-1. The Hertzian model was used with a pyramid tip indenter to calculate the Young’s

13

modulus.

14

The compressive modulus of the bulk alginate and alginate/gelatin hydrogels was determined

15

using a AR1000-N rheometer (TA Instrument, UK) with 40 mm diameter stainless steel parallel

16

plates. The slope of the stress-strain curve for each disc shape casted hydrogel (40 mm diameter

17

and 2 mm thickness) at 5–10% strain was taken as the bulk compressive modulus (E).

18

Statistical analysis

19

All the presented values are expressed as mean ± standard deviation (SD), and each

20

experiment was performed in triplicate. A two-way ANOVA analysis with replication test

21

followed by Tukey’s post hoc test was used to determine statistical differences. p values less than

22

0.05 were considered statistically significant.



1 2

Page 14 of 46

RESULTS AND DISCUSSION A microfluidic platform for generation of microgels using different oils

3

We combined an alginate-Ca-EDTA complex with gelatin at a ratio of 1:1 to make a 1%

4

hydrogel solution in phosphate-buffered saline (PBS) for single cell encapsulation (see Methods

5

for more details). A flow-focusing microfluidic device was used to encapsulate cells (Figure 1A).

6

Different oils, including olive, peanut, grapeseed, sunflower, corn, and mineral oil containing 2

7

wt% lecithin as an emulsifier (i.e., the carrier phase) were introduced into Inlet 1 of the device,

8

while the alginate-based solutions (i.e., the dispersed phase) entered through Inlet 2. The

9

alginate-based droplets that were generated and stabilized with the 2 wt% lecithin were collected

10

from Outlet 1, as shown in the Figure 1A. Movies S1 and S2 are provided under supporting

11

information to show alginate droplet generation and collection processes, respectively; grapeseed

12

oil containing 2 wt% lecithin was used to generate these droplets within our microfluidic flow-

13

focusing device.

14

The droplet size depends on the nature of the dispersed and carrier phases, the diameter of 20-23

15

the channel, the device geometry, and the ratio of the oil/water phases

. The results of the

16

alginate droplet size generated by different oils are presented in Figure 1B. According to the

17

Hagen-Poiseuille equation in a laminar flow, (∆ =

18

between the flow rate (Q) and pressure drop (∆P) in an incompressible fluid with a defined

19

dynamic viscosity (µ) through a pipe with a specific length (L) and diameter (R) 24-25. Our results

20

indicated that an increase in the pressure drop (in the range of 10-60 kPa, made by applying more

21

vacuum on the collecting outlet) and consequently an increased flow rate led to the generation of

22

smaller alginate droplets in the investigated oils. This trend is consistent with a previous report of

8 ), there is a linear relationship


Page 15 of 46 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


1

Ca-alginate microspheres generated in sunflower seed oil, which were used to encapsulate gold

2

nanoparticles 26. The relationship between droplet size and the flow rate can be explained by the

3

capillary number ( = =

4

and viscosity of the carrier phase, respectively, and is the interfacial tension between the water

5

and oil phases 20, 27. Based on this relationship, the droplet size of the generated microgels can be

6

correlated to the Ca number. To show the stability and monodispersity of the droplets, we took 4

7

µl of the microgels generated, loaded them onto the hemocytometer, then captured images

8

(Figure S1). Results showed that the microgels were stable and mostly monodisperse.

9

), in which is the droplet size, and are the velocity

Moreover, the viscosity and interfacial tension of the oil phase are the two main parameters 28-29

10

that can affect the droplet size and stability at the same flow rates

11

determined the viscosities of several oil phases containing 2 wt% lecithin, with mineral oil

12

(114.6 ± 1.1 cps) >> olive oil (58 ± 0.7 cps) > peanut oil (49.6 ± 0.5 cps) > grapeseed oil (40.6 ±

13

0.5 cps) > corn oil (40.4 ± 0.4 cps) > sunflower oil (37.97 ± 0.4 cps) (three measurements were

14

performed for each oil). Among the tested oils, olive oil could generate stable droplets in a wider

15

range of flow rates. This can be attributed to the lower interfacial tension between the olive oil

16

and water phases compared to other oils in the same aqueous conditions (2 wt% alginate-Ca-

17

EDTA complex; Figure 1C). Moreover, the results showed that the addition of gelatin to the

18

alginate-Ca-EDTA complex led to a significant decrease in droplet sizes in most of the oils due

19

to a decrease in their interfacial tension (Figure 1D). Mineral and corn oils produced the most

20

stable alginate/gelatin droplets, as determined by microscopic observations. A similar trend was

21

observed for the generation of alginate/gelatin microgels, with increased pressure drop resulting

22

in smaller droplet size (Figure 1D). The effect of gelatin addition to alginate (at a ratio of 1:1) on


. In this regard, we


1

droplet size for each oil separately is presented in Figure S2. A two-way analysis of variance

2

showed that the addition of gelatin has a significant effect on the droplet size for peanut, corn,

3

sunflower, and mineral oils.

4

Cell encapsulation within microgels

5

Internal and external gelation methods have been used to crosslink alginate. Each method has

6

its own advantages and disadvantages. In the external crosslinking method, the Ca ions are

7

provided using an external source such as an aqueous calcium solution. This method has been

8

widely used for mostly bulk preparation of hydrogels (e.g., extrusion of hydrogel solution into

9

CaCl2 bath 10). However, the control of the size and shape of droplets (polydispersity) is difficult

10

to achieve in the syringe-extrusion method. But these parameters are highly tunable using drop-

11

based microfluidics. The microfluidic technique can produce monodisperse and uniform

12

microgels by first generating well-defined microdroplets, then by triggering gelation. Direct

13

infusion of CaCl2 into the oil phase, however, could block the flow-focusing junction when

14

making the droplets in a short period due to the local gelation of alginate. Moreover, the infused

15

CaCl2 in the oil phase is not stable for a long period. Internal gelation would be an ideal

16

alternative method to resolve this issue. In this alternative, an external agent (acetic acid) is

17

necessary to trigger the release of calcium ions from the alginate/Ca-EDTA complex solution

18

and consequently, alginate crosslinking. This would delay the crosslinking of alginate till the

19

well-defined microdroplets are formed. This internal gelation method using acetic acid has been

20

used to encapsulate natural compounds 30, proteins 31-33 and cells 2-3, 34.

21

The crosslinking gradient and consequently the heterogeneity inside the microgels—caused

22

by restricted diffusion of ions—are the issues for large alginate microgels or beads (e.g., 400

23

µm-2 mm that challenges researchers

35

) which would directly affect the cell viability inside


Page 16 of 46

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1

these microgels. Here, using the microfluidic technique, we can create much smaller

2

monodisperse microgels (e.g., 50 µm) which can be homogeneously crosslinked quickly. To

3

confirm this, we produced alginate/gelatin (ratio 1:1) droplets and solidified them by 0.25 wt%

4

acetic acid. Then, we analyzed the homogeneity, and the internal and external morphologies of

5

these microgels using cryo-SEM. (Figure 2A,B). As we expected, the exterior and interior parts

6

of the microgels were uniformly solidified. To further confirm the homogeneity and

7

consequently the uniformity of the mechanical properties of the microgels, we also obtained the

8

concentration distribution of a dye inside the microgels over time (Figure S3). For this aim, first,

9

we electrostatically bound the microgels onto a polylysine-coated glass surface, then we injected

10

a FITC-labeled dextran (20 kDa) into the microgel’s surrounding aqueous solution and took

11

images of the inward diffusion of the dye into the microgel core over time (Figure S3A-E). We

12

monitored the dye’s concentration distribution over time, using Matlab R2017a for quantification

13

(Figure S3F). We found that the dynamics of the diffusion of the dye was constant, which

14

indicates that the matrix of the microgel is homogeneous.

15

To further prove this point, we also performed a simulation of the diffusion of the dye into a

16

solid sphere with a constant diffusion coefficient using COMSOL Multiphysics. Eventually, by

17

solving the dimensionless form of the diffusion equation using COMSOL Multiphysics, we

18

obtained the radial distribution of the diffused dye over time. As can be seen in Figure S3F, as

19

time goes on, the concentration inside the sphere becomes higher and finally, the concentration

20

throughout the sphere becomes uniformly distributed. The similarity between the results obtained

21

from simulations and the corresponding experimental data suggests that the diffusion dynamic

22

inside the microsphere follows Fick’s Law and therefore, the diffusivity is constant over the

23

whole microsphere. This fact provides further evidence that the microsphere is indeed



Page 18 of 46

1

homogeneous and suggests that the mechanical properties over the entire volume of the

2

microsphere are uniform.

3

To illustrate the effect of the acetic acid concentration on surface topography and mechanical

4

properties of microgels in liquid medium, we used the AFM (5 microgels were analyzed for each

5

experiment). Results showed that an increase in the acetic acid concentration could change the

6

surface roughness, with smoother surfaces at higher concentrations of acetic acid (Figure 2C,D

7

and Figure S4A-C). It was previously reported that an increase in the concentration of calcium

8

ions in the alginate crosslinking process could reduce the surface roughness of alginate films due

9

to the relatively lower swelling degree during crosslinking

36

. In addition, our results

10

demonstrated that an increase in acetic acid concentration resulted in an increase in mechanical

11

properties of these microgels (Figure 2E).

12

We selected this alginate-Ca-EDTA/gelatin (1:1) solution to make droplets to improve the

13

cell adhesion properties of alginate, and consequently to enhance cell growth and proliferation

14

inside the microgels. We evaluated the encapsulation efficiency of cells inside the microgels over

15

a wide range of cell densities in the aqueous phase (Figure 3A). To calculate the average number

16

of cells in each droplet, 500 distinct microgels were analyzed. At low cell densities, only a small

17

portion of droplets contained at least one cell (other droplets were mostly empty). Increasing the

18

cell density in the aqueous dispersed phase from around 180,000 to 3,700,000 cells/ml resulted

19

in a 14x increase in cell encapsulation efficiency. Based on these results, we set the feed cell

20

density to 2,750,000 cells/ml to create a uniform population of cell-embedded microgels,

21

although we still observed a small portion (< 5%) of empty droplets or those with more than one

22

cell in each droplet under these conditions. A similar trend with no significant difference was

23

observed for the cell encapsulation efficiency using different oils (Figure S5A).


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1

In this study, we used acetic acid to solidify the generated droplets and tested the effect of

2

acetic acid concentration (0.25, 0.5, and 1 wt%) and exposure time on the cell viability (Figure

3

3B). All other parameters, such as washing steps for removal of the acetic acid and EDTA

4

residues, were fixed in these experiments. The results clearly indicated that an increase in acetic

5

acid concentration and exposure time resulted in a significant decrease in cell viability, with 1

6

wt% acetic acid at 2 min exposure time leading to a 50% drop in cell survival. The effect of

7

exposure time was also investigated on cell viability using live and dead cell staining (Figure 3C

8

and Figure S5B-D). At low concentrations of acetic acid, increasing exposure time was needed to

9

solidify the microgels, while at higher concentrations (1 wt%), a very short exposure time was

10

sufficient to generate stable microgels with a high population of viable cells. In a low

11

concentration of acetic acid (0.25 wt%), the majority of embedded cells remained alive (75%),

12

which is consistent with the literature 2-3. An increase in the acetic acid concentration resulted in

13

an increase in the strength and stability of the microgels, as observed by optical microscopy and

14

confirmed by Young’s moduli of the microgels measured by AFM; however, the majority of

15

cells were unable to survive in such high acidic conditions for long. Based on these experiments,

16

we exposed the generated droplets to 1 wt% acetic acid followed by immediate dilution and

17

washing steps with hexadecane containing 2 wt% Span 80. In this fashion, minimizing the

18

exposure time compensated for the adverse effect of the high concentration of acetic acid,

19

allowing for the production of mechanically robust microgels containing viable cells.

20

To determine the residual concentrations of Span 80 and lecithin in the microgel water phase

21

after all washing steps, which can potentially affect cell viability, we extracted these surfactants

22

using isopropanol (IPA). First, we added an equal portion of IPA to the water phase. Then, the

23

mixture was vortexed vigorously and centrifuged at 5000 g for 10 min. Thereafter, the



1

absorbance was measured at 269 and 244 nm for detection of Span 80 and lecithin, respectively,

2

using a UV–vis spectrometer. Residual concentration of Span 80 and lecithin in the water-

3

microgel mixture were then determined using standard curves of Span 80 and lecithin in IPA

4

(Figure S6). Our data showed that 0.00009 and 0.00002 g/ml of the Span 80 and lecithin were

5

detectable in the microgel water phase after all washing steps.

6

Figure 3D represents the effect of the oil type on cell viability during encapsulation, as

7

measured by a trypan blue assay. For these experiments, we applied the optimum approach to

8

solidify the droplets (i.e., using 1 wt% acetic acid followed by immediate dilution). The results

9

show that the viability of the embedded cells was independent of the type of oil used, whether

10

fluorinated (HFE-7500) or non-fluorinated. Mineral oil has a high content of heavy alkanes (C15–

11

C30), while edible oils mostly contain saturated and unsaturated fatty acids, such palmitic, oleic,

12

and linoleic acids

13

among these non-fluorinated oils and that there was a high cell viability (greater than 90%) in

14

both pure alginate and alginate/gelatin microgels. Short exposure time to oil molecules can be a

15

reason for the observed weak effect of oil type on cell viability. We used a method in which the

16

oil was washed out as quickly as the droplets were formed and cross-linked. These findings are

17

consistent with previous publications. For example, Workman et al. showed over 90% cell

18

viability for HEK293 cells in pure alginate microgels encapsulated by mineral oils 7, 41.

37-40

. Results showed that there was no significant difference in cell viability

19

We also investigated cell adhesion, proliferation, and division over a period of one week

20

using optical microscopy. The microgels were adhered to the surface of a cell culture flask

21

coated with polylysine to monitor cell proliferation (Figure 3E-G). The images show that the

22

embedded single cells proliferate and form a clump inside the microgels over a period of 7 days.

23

The incorporation of gelatin into the microgels provides adhesion sites at Arg–Gly–Asp (RGD)


Page 20 of 46

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1

sequences in its structure 2. As demonstrated, the morphology of the microgels did not change

2

significantly with time, maintaining a round shape (monodispersed spheres with a mean diameter

3

of 70 µm).

4

Surface and mechanical properties of microgels

5

We used atomic force microscopy (AFM) analysis to assess the effect of gelatin

6

concentration on the Young’s modulus of the cell-embedded microgels cultured in MEM culture

7

medium. AFM surface topography can be used to measure the wrinkles and dimples on the

8

surface of the microgels, which can be correlated to the average roughness and other quantitative

9

mechanical properties. Roughness can strongly affect the microgel swelling behavior and its

10

erosion rate under mechanical stress during cell culture over time, with greater surface roughness

11

leading to higher swelling and erosion rates 42-43. The topography images of the pure alginate and

12

alginate/gelatin (1:1) microgels are depicted in Figure 4A,B respectively. The surface

13

topography of other alginate/gelatin microgels with ratios of 2:1 and 1:2 are presented in Figure

14

S7A,B. As shown qualitatively in topographic images and quantitatively in terms of the average

15

roughness (Figure 4C), we found that an increase in gelatin concentration also increased the

16

surface roughness of the microgels. For gelatin added to the pure alginate at 0.66 wt% (1:2), the

17

average roughness increased by 12.5% compared to the pure alginate. Based on the topographic

18

images, under stresses applied by the cell culture medium, pure alginate microgels swelled

19

uniformly, while microgels with different gelatin concentrations showed a heterogeneous

20

swelling behavior with less uniform morphology. While surface roughness, indicated by the

21

presence of wrinkles and dimples on the microgels, can increase microgel erosion rates, it can

22

also be beneficial for tissue engineering applications by providing a permissive environment for

23

the fusion of neighboring microgels to form constructs of desired geometries. To investigate the



1

effect of time on the surface topography, we incubated cell-embedded microgels in stagnant cell

2

culture medium for a week at 5% CO2 and 37 ºC, followed by AFM analysis. As shown in

3

Figure 4D, the wrinkles and dimples on the surface of the microgels increased with incubation

4

time. Accordingly, the average roughness and the root mean square of the alginate/gelatin (1:1)

5

microgels over one week increased from 30.49 ± 0.93 nm and 39.72 ± 1.23 nm to 37.97 ± 1.07

6

nm and 49.08 ± 1.34 nm, respectively (Five microgels for each group were analyzed).

7

In this study, gelatin was added to alginate to improve cell attachment and proliferation.

8

However, because of its metalloprotease sites, gelatin has a high biodegradation rate, which will

9

negatively affect the microgel’s mechanical properties

44

. Mechanical properties of microgels

10

have a large impact on the fate of embedded cells in terms of attachment, proliferation, and

11

differentiation. Different approaches have been used to analyze the mechanical properties of

12

microgels, such as AFM, indentation, micropipette aspiration, bulge and compression

13

measurements, and microﬂuidic conﬁnement

14

contains a variety of amino acids, like glycine and proline. A high proportion of gelatin to

15

alginate results in brittleness and can decrease the microgel quality 44. To study this effect, we

16

changed the gelatin content of the cell-embedded alginate/gelatin microgels and monitored the

17

materials’ change in mechanical properties in the culture medium using AFM at two time points,

18

day 1 and day 7 (Figure 4E). Five microgels for each group were analyzed for their force curves

19

and maps and the results showed that an increase in the gelatin concentration resulted in a

20

significant decrease in the nano-Young’s moduli of the microgels, which is related to the brittle

21

properties of the gelatin. Addition of 0.66 wt% gelatin to alginate led to an approximately 85%

22

drop in the Young’s modulus of the microgels (from 21.01 ± 1.1 kPa to 3.78 ± 0.2 kPa).

23

Moreover, the mechanical properties of the cell-embedded microgels decreased over time. This

45

. Gelatin—a denatured form of collagen—


Page 22 of 46

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1

decrease was more pronounced in pure alginate. The decrease in the bulk modulus of

2

alginate/gelatin microgels with time has been previously reported

3

the fast hydrolysis and degradation of gelatin by cell-secreted enzymes and the gradual

4

dissociation of alginate by Ca2+ ion exchange with Na+ in the culture medium. Our results

5

showed that over a 7-day incubation period, the nano-Young’s moduli of cell-laden gelatin-

6

containing microgels did not decrease significantly compared to pure alginate microgels and cell-

7

free microgels (data not shown). This can be explained by the secretion of extracellular matrix

8

(ECM) proteins, such as collagen type I, by viable cells embedded inside the microgels

9

mentioned previously, gelatin is derived from collagen by denaturation of the triple-helix

10

structure. Gelatin has many integrin-binding sites for cell adhesion and can improve cell

11

proliferation and ECM deposition even better than non-denatured collagen

12

moduli of the alginate and alginate/gelatin bulk materials are also presented in Figure 4F, which

13

show the same trend as the corresponding microgels to a more exaggerated degree, with

14

increased gelatin content resulting in a lower Young’s modulus. This finding is upheld by

15

previous results in which Mao et al. reported a similar trend between the bulk and nano behavior

16

of alginate hydrogels3. The relevant stress-strain curves of the bulk hydrogel materials can also

17

be found in the supplementary data (Figure S8).

10

, which can be attributed to

46

10

. As

. The compressive

18

For intestinal organoid culture, mechanically dynamic matrices with specific compositions

19

are required. Temporal changes in matrix stiffness are essential, as matrices with high stiffness

20

(over 1 kPa) at early stages of the cell culture will enhance cell expansion through a yes-

21

associated protein 1 (YAP)-dependent mechanism. A subsequent decrease in matrix stiffness

22

over time is then beneficial for organoid formation and differentiation

23

incorporated three essential proteins of the intestinal crypt niche (fibronectin, collagen IV, and


11

. In this study, we


1

laminin1-1-1) into a biodegradable alginate/gelatin (1:1) matrix. Addition of these proteins had

2

no significant effect on the mechanical properties of the microgels generated, as the amount of

3

these proteins were negligible (0.085 wt%) compared to the alginate/gelatin concentration (1

4

wt%).

5

6

A 3D in vitro microgel-based model for co-culture of crypt and Peyer’s patch cells

7

To test the feasibility of co-culturing crypt and Peyer’s patch cells as a model for in vitro

8

evaluation of immune response on crypt cells, we fabricated a polymeric 3D printed mold

9

composed of two interconnected chambers (Figure 5A–C). The interior chamber was designed to

10

house crypt cell-embedded microgels while the Peyer’s patch-embedded microgels were loaded

11

into the exterior chamber. The soluble mediator channels were designed to allow media

12

exchange between interior and exterior chambers. The encapsulated cells, including crypt

13

(Figure 5D,E) and Peyer’s patch cells (Figure 5F), were separately cultured in each chamber,

14

though they could readily interchange their metabolites via the porous walls of the mold. The

15

results showed that both cell types proliferated during a 2-week co-culture. The observed

16

hydrogel polydispersity in the aqueous medium (as evidenced in Figures 5D and F)—compared

17

with the microgels obtained in oil in Figure 1—could be attributed to the different crosslinking

18

and swelling rates of these microgels. During microgel crosslinking, smaller droplets tend to be

19

more strongly crosslinked (because of fast Ca ions diffusion), thus forming a smaller hydrogel

20

network, in contrast to that formed by bigger droplets. Microgels with loose hydrogel network

21

and bigger sizes tend to swell more, as compared to smaller and more tightly crosslinked

22

microgels. Time evolution of organoid growth is depicted in Figure 5G–J. The two terminal parts

23

of the U-shaped crypts joined at day 1, and they started budding at day 3. Their growth continued


Page 24 of 46

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1

to fill approximately the whole microgel volume. Live and dead cell staining at day 7 is

2

presented in Figure 5K,L. As shown, the number of Peyer’s patch cells increased over time (from

3

day 1 to day 7), and only a few dead cells were visualized, possibly due to nutrient deficiency

4

inside the microgels. Moreover, two controls (Figure S9-A,B) are provided to demonstrate the

5

viability of each cell type alone within the microgels inside the 3D printed mold after a 7-day

6

culture. In contrast to the results obtained in the co-culture method, those crypt cells that were

7

encapsulated inside microgels and cultured alone could not retain viability and mostly died

8

within the 7-day culture due to the lack of exogenous factors. In the co-culture system,

9

exogenously supplemented niche factors produced by Peyer’s patch cells work cooperatively

10

with niche factors generated by crypt cells themselves (e.g., Paneth cells, and factors generated

11

by Paneth cells including Notch, EGF, and Wnt proteins) to support the organoid culture. Our

12

results showed that viable organoids can be cultured within microgels in our co-culture method.

13

On the other hand, the viability of the Peyer’s patch cells did not change significantly with the

14

presence or absence of crypt cells (Figure S9B).

15

To check for the presence of stem cells inside the microgels over the culturing period, we

16

stained cell-embedded microgels with the Bmi-1 antibody. Bmi-1 is part of the Polycomb group

17

gene family, which is expressed in intestinal stem cells (ISCs). Bmi-1+ ISCs are pluripotent stem

18

cells with the capability of self-renewal, which plays an essential role in crypt maintenance 47. As

19

shown in Figure 6A–C, the stemness of crypt stem cells was maintained over 21 days of co-

20

culture. The 3D structures of the organoids after a 7-day culture have also been presented in

21

movies S3 and S4. Green stains in confocal movies show Bmi-1 positive cells and purple stains

22

show the nuclei of the cells (DAPI) cultured inside crypt niche microgels. The presence of

23

differentiated epithelial cells was also evaluated using immunostaining with villin (Figure 6D).



Page 26 of 46

1

Villin, which is highly expressed in gastrointestinal tract epithelial cells, is an actin-modifying

2

protein that regulates actin reorganization, cell motility, and epithelial cell morphology

3

shown in Figure 5D and E, both differentiated and undifferentiated cells were present in the

4

organoids, demonstrating their health in our co-culture model. To make sure the cells were alive,

5

we stained the cultured organoids within microgels using live/dead fluorescent staining dyes

6

(Figure 5F). Results showed that most of the cells were alive after 21-day co-culture. Current gastrointestinal tract models, including ex vivo (e.g., organotypic tissue slices

7 8

precision-cut intestinal slices 50-51) and in vitro models (e.g., gut-on-a-chip

9

55

52-54

48

. As

49

and

and microfluidic

approaches) are indispensable tools for studying the intestine, but do not reflect the living,

10

breathing complexity of the human intestine. Both existing ex vivo and in vitro models have

11

distinct limitations. Although ex vivo models incorporate several components of the

12

gastrointestinal tract, including immune cells (e.g., Peyer's patch cells) and crypt cells, they

13

assume that we can model the pathophysiology of human diseases on animals—an assumption

14

that has led to the costly failure of many clinical drug trials (approximately 9 out of 10

15

Animal models are also unpredictable and fraught with ethical concerns. In vitro models assume

16

that a single type of epithelial cancer cell line (e.g., Caco-2 cells) inside a microfluidic device has

17

the same uptake mechanism and behavior as in the diverse microenvironment of the human

18

gastrointestinal tract. However, this microenvironment is not composed of only one type of cell,

19

but rather a wide array of crypt stem cells, goblet cells, enterocytes, enteroendocrine cells, tuft

20

cells, Paneth cells, immune cells, and microbiota—all of which influence each other through

21

intricate cross-talking mechanisms, such as paracrine and autocrine signaling in order to

22

maintain cell viability. To date, it has been difficult to create in vitro models that reflect this

23

complexity.


56

).

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1

In the present study, by taking advantage of droplet-based microfluidics, we were able to

2

develop an in vitro model that enables practical investigation across Peyer’s patch and crypt

3

niches of the intestine. With Peyer's patch cells’ integrity maintained, the present model can be

4

used for evaluation of intestinal immune responses to infections in a 3D microenvironment in

5

future experiments. Unlike individual normal cell types, which die within days and are

6

continually replaced, organoids are composed of irreplaceable stem cells that do not die over a

7

person’s lifetime, and in fact are responsible for regenerating all other cell types in the crypt-

8

villous domain while exhibiting physiological functions like Na+ absorption and Cl− secretion 13.

9

The present model can be used as a novel approach for the co-culture of organoids to evaluate

10

the effect of different toxicological and environmental factors on the proliferation and

11

differentiation of stem cells. It is worth mentioning that other well-defined matrices for intestinal

12

organoids, such as polyethylene glycol hydrogels with controllable mechanical properties 11, can

13

easily be applied in the present model. In the future, this system could allow us to model

14

inflammation in the gastrointestinal tract to investigate the effect of different environmental

15

factors on both immune and crypt-villous cells in the presence or absence of drug intervention.

87

CONCLUSION

88

In the present study, we compared microgel generation efficiency in several non-fluorinated

89

oils. The effects of oil type on droplet size and the viability of the embedded cells were

90

investigated. We also showed that biological and nano-mechanical properties of microgels can

91

be controlled by hydrogel composition. Based on these findings, we demonstrated a model for

92

the evaluation of the interaction between immune cells and crypt cells using a microfluidic

93

approach to encapsulate crypt cells and Peyer’s patch cells inside distinct alginate/gelatin

94

microgels, and tested the cellular co-cultures inside a static 3D-printed mold. The results of this



1

study put forward the possibility of a novel application of microfluidic-based microgels with

2

tunable physicomechanical and biological properties for organoid cultures, gastrointestinal tissue

3

engineering, and regenerative medicine.

4 5

ASSOCIATED CONTENT

6

Supporting Information

7

The Supporting Information is available free of charge.

8

Movies S1 and S2: alginate droplet generation and collection, respectively, using grapeseed

9

oil containing 2 wt% lecithin in our microfluidic flow-focusing device. Movies S3 and S4: 3D

10

structure of the organoids after a 7-day culture using confocal microscopy. Figure S1: Optical

11

microscopic images of the alginate microgels generated by different oils. Figure S2: the mean

12

droplet size of microgels composed of pure alginate and alginate/gelatin (1:1) using different oils

13

as the carrier phase. Figure S3: The concentration distribution of FITC-labeled dextran inside

14

alginate/gelatin (1:1) microgels as a function of time. Figure S4: Surface topography of the

15

alginate/gelatin (1:1) microgels crosslinked by different concentrations of acetic acid. Figure S5:

16

average cell numbers in the alginate-gelatin (1:1) microgels—based on the cell densities in the

17

feed for droplets generated using different fluorinated and non-fluorinated oils—and

18

representative images of live and dead staining of cell-embedded microgels exposed to different

19

concentrations of acetic acid. Figure S6: The residual concentrations of Span 80 and lecithin in

20

the microgel water phase. Figure S7: AFM surface topography of microgels composed of

21

alginate/gelatin ratios of 2:1 and 1:2. Figure S8: representative stress-strain curves of disc-shaped

22

bulk hydrogels composed of pure alginate and alginate/gelatin at different gelatin ratios (2:1, 1:1,

23

and 1:2). Figure S9: live and dead staining of each cell type (Peyer’s patch and crypt cells)


Page 28 of 46

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1

cultured separately within microgels inside the 3D printed mold after a 7-day culture as controls

2

for the co-culture method.

3 4

AUTHOR INFORMATION

5

Corresponding Author

6

*E-mail: [email protected]

7

Notes

8

The authors declare no competing financial interest.

9

ACKNOWLEDGEMENTS

10

The authors gratefully acknowledge the Cornell Nanoscale Science and Technology Facility

11

(CNF), which is supported through the NSF NNCI program (Grant Number ECCS-1542081) and

12

Cornell Center of Materials Research (CCMR). We thank Dr. Benyamin Davaji for his

13

assistance in microfluidic device fabrication.

14

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1

Figure captions

2

Figure 1. Schematic set-up for single cell encapsulation. (A) Microfluidic flow-focusing device

3

for the generation of single cell-laden microdroplets. The mixture of 2 wt% lecithin with

4

different oil types, including olive, corn, grapeseed, sunflower, mineral, and peanut oils (i.e., the

5

carrier phase) entered through Inlet 1, while the dispersed phase composed of the cells and the

6

precursor hydrogel solution (i.e., alginate and gelatin complexes in PBS) entered through Inlet 2.

7

The resulting embedded cell-laden droplets were then collected from Outlet 1 for further

8

induction of gelation with acetic acid. The mean droplet size of the (B) alginate and (D)

9

alginate/gelatin (ratio of 1:1) microgels generated by different oil types at different driving forces

10

(∆P) (for each data point, 500 droplets were analyzed). Inset images show representative size

11

distribution histograms of the microgels generated by corn oil. (C) Interfacial tension between

12

the aqueous phase, including pure alginate (red) and alginate/gelatin (1:1, blue) solutions, and the

13

oil phase. A statistically significant difference between the test and all other groups is indicated

14

by a “*” sign. Error bars are SD of triplicate samples.

15

Figure 2. Cryo-SEM images of (A) the surface and (B) the focused ion bean sectioned interior

16

phase of the alginate/gelatin (1:1) microgels crosslinked by 0.25 wt% acetic acid infused in oil.

17

The arrow indicates the sectioned part of the microgel (C) Surface topography of the

18

alginate/gelatin (1:1) microgels crosslinked by 0.25 wt% acetic acid infused in oil obtained by

19

AFM in liquid medium. (D) Average roughness and (E) the nano-Young’s moduli of the

20

alginate/gelatin (1:1) microgels crosslinked by different concentrations of infused acetic acid in

21

oil obtained by AFM in liquid medium.

22

Figure 3. (A) Average cell numbers (500 microgels were analyzed) inside the alginate/gelatin

23

(1:1) microgels as a function of the cell densities in the feed. (B) Cell viability determined by the

24

trypan blue exclusion method at three different acetic acid concentrations and various exposure



1

times. (C) We also studied the effect of acetic acid concentration on cell viability using live

2

(green) and dead (red) cell staining of the cell laden alginate/gelatin (1:1) microgels after 1 min

3

exposure to acetic acid. (D) The viability of cell-embedded microgels were assessed by the

4

trypan blue exclusion method using different types of oil for the generation of the

5

alginate/gelatin (1:1) droplets. The generated droplets were solidified in 1% acetic acid in the

6

same oil used for the droplet generation and immediately diluted and washed with hexadecane

7

containing 2 wt% Span 80. (E–G) Optical microscopy images of cell-laden alginate/gelatin (1:1)

8

microgels containing Caco-2 cells at three time points, including (E) day 1, (F) day 3, and (G)

9

day 7. A statistically significant difference between the test and all other groups was indicated by

10

a “*” sign. Error bars are SD of triplicate samples.

11

Figure 4. Surface topography of microgels composed of (A) pure alginate and (B)

12

alginate/gelatin (1:1). (C) Average roughness of the microgels at different alginate/gelatin ratios,

13

including pure alginate, 1:2, 1:1, and 2:1. (D) Cell-embedded alginate/gelatin (1:1) microgels

14

were cultured for one week then subjected to AFM analysis to see the effect of time on surface

15

topography. The mechanical properties of the alginate/gelatin microgels at different

16

alginate/gelatin ratios, including pure alginate, 1:2, 1:1, and 2:1 were determined using (E) AFM

17

for cell-embedded microgels at two time points (day 1 and day 7) and (F) using a rheometer for

18

bulk hydrogels. The average of Young’s modulus of 5 microgels was reported.

19

A statistically significant difference between the test and all other groups was indicated by a “*”

20

sign. Error bars are SD of triplicate samples.

21

Figure 5. (A–C) The 3D printed insert for co-culture of the crypt and Peyer’s patch cell-

22

embedded microgels. (D, E) Encapsulated crypt cells inside alginate/gelatin (1:1) microgels. (F)

23

A Peyer’s patch cell-embedded alginate/gelatin microgel (1:1). (G–J) Crypt cell growth inside


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1

the microgels. Isolated crypt cells from a mouse intestine, cultured in the 3D printed insert at (G)

2

day 1, (H) day 3, (I) day 7, and (J) day 14 along with Peyer’s patch cells. The borders of the

3

microgels have been encircled inside dotted lines. Live (green) and dead (red) staining of (K)

4

Peyer’s patch cells after 7 days co-cultured with (L) crypt cells.

5

Figure 6. Confocal images of intestinal organoids within microgels co-cultured with Peyer’s

6

patch cells in the 3D printed insert. Bmi-1 (red) and DAPI (blue) staining of organoids cultured

7

in crypt niche microgels (alginate/gelatin 1:1, containing fibronectin, collagen IV, and laminin

8

proteins) at (A and E) day 7, (B) day 14, and (C) day 21. (D) Villin (green) and DAPI (blue)

9

staining of the organoids at day 7. (F) live (green) and dead (red) staining of crypt cells within

10

microgels co-cultured with Peyer’s patch cells in the 3D printed insert at day 21. Scale bars, 70

11

µm.

12



1

2 3 4

Figure 1.

5 6 7


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1 2

3 4

Figure 2.

5



1

2 3

Figure 3.

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1

2 3

Figure 4.

4 5 6 7



1 2 3 4 5 6 7

Figure 5.


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11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

Figure 6.



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Microfluidic-Based Cell-Embedded Microgels Using Nonfluorinated

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