communications

1264

Alginate hollow fibers

Synthesis of Cell-Laden Alginate Hollow FibersUsing Microfluidic Chips and MicrovascularizedTissue-Engineering Applications**

Kwang Ho Lee, Su Jung Shin, Yongdoo Park, and Sang-Hoon Lee*

Biomimetic microstructures can be used for tissue assembly,

tissue and organ regeneration, and as drug carriers.[1] To date,

diverse microscale structures that use microengineering

technologies (e.g., micro- or nanoparticles, micro- or nano-

fibers, and 2D patterns) have been used as drug delivery

systems for the controlled release of therapeutic drugs, as

scaffolds for support of cells in defective tissues, and as basic

materials for the study of cell–cell interactions.[2,3] In living

organisms, microscale and porous hollow fibers, such as blood

vessels, are important for the delivery of small amounts of

liquid to specific places. Such structures also have industrial

applications including the filtering and channeling of small

volumes of liquid.[4] In spite of their diverse potential

applications, the study of hollow fibers has been limited to

specific areas, such as hemodialysis, because fabrication is very

difficult.[5] In addition, it is difficult to load cells and proteins

into hollow fibers without loss of biological activity.

Previously, we demonstrated the fabrication of microscale

fibers or hollow fibers based on an ‘‘on-the-fly’’ method that

uses microfluidic chips. However, the material that we

originally used was not biodegradable or biocompatible.[6,7]

Subsequently, we reported the production of alginate solid

fibers and cell encapsulation in these fibers.[8] In addition,

Sugiura et al.[9] have reported the fabrication of an alginate

[�] Prof. S. H. Lee, Dr. K. H. Lee, S. J. Shin

Department of Biomedical Engineering

College of Health Science, Korea University, Seoul (Korea)

E-mail: dbiomed@korea.ac.kr

Dr. K. H. Lee

Research Institute of Health Science

Korea University, Seoul (Korea)

E-mail: biomems.tiger@gmail.com

Prof. Y. Park

Department of Biomedical Engineering

Brain Korea 21 Project for Medical Science

College of Medicine, Korea University, Seoul (Korea)

[��] This study was supported by a grant from the Korea Science andEngineering Foundation (KOSEF) (grant no. ROA-2007-000-20086-0) and the Korea Healthcare Technology R&D Project, Ministry forHealth Welfare and Family Affairs, Republic of Korea (grant no.0405-ER-01-0304-0001).

: Supporting Information is available on the WWW under http://www.small-journal.com or from the author.

DOI: 10.1002/smll.200801667

Figure 1. a) Schematic of the alginate hollow fiber-generating chip. The

chip has two stages: first, generation of core and sample flow; second,

generation of core, sample, and sheath flow. For the generation of cell-laden

fibers, a mixture of HIVE-78 cells and 2% w/w alginate solution was

used as the sample fluid. b) Micrograph of the hollow fiber generation

process (movie clip is in the Supporting Information).

� 2009 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim small 2009, 5, No. 11, 1264–1268

Figure 2. a) SEM image of an alginate hollow fiber. b) Variation of fiber diameter as a function

of sample and sheath flow rate. c) 2D wound alginate hollow fiber matrix on a cover glass.

d) Ultimate strength of alginate hollow fiber synthesized with different materials.

tubular gel and cell encapsulation within

this gel. Despite this progress, it is still

challenging to produce hollow microfibers

whose size can be easily controlled that are

made of non-swelling, biodegradable, and

biocompatible materials that can be loaded

with biomolecules and cells during produc-

tion, and that can be easily handled and

aligned. Here, we propose a new approach

for the synthesis of hollow alginate micro-

fibers that satisfy all these requirements by

use of a microfluidic chip.

Alginate has several advantages over

other biological hydrogels such as hyaluro-

nic acid, fibrin, agar, gelatin, and chito-

san.[10,11] There are many studies of the use

of alginate in therapy, pharmacology, and

medicine, but these studies are limited by

poor treatment methodologies.[12] The use

of microfluidic chips for the construction of

more complex structures may resolve these

problems.[13,14] In particular, changing

the flow rate easily controls the diameter

of the hollow alginate fibers. We also

describe the synthesis of cell-laden and

protein-immobilized alginate hollow fibers

and in situ alignment of fibers. The cells

remained alive within the fibers due to the nontoxic fabrication

process. The properties of synthesized fibers were also

evaluated by measuring mechanical strength. As an applica-

tion of these hollow fibers, we emulated vascular tissue by

embedding endothelial cells within the hollow fibers into

smooth muscle-cell-laden agar–gelatin–fibronectin (AGF)

hydrogels. Cells within the hollow fiber and the hydrogel

were cocultured and remained alive with intact vascular

structure for 7 days, indicating the effectiveness of our method

for the engineering of vascularized tissue.

The microfluidic process used for fabricating hollow fibers

(the ‘‘on-the-fly’’ method) was modified from our previously

reported protocol.[8] The fluidic device consisted of a

pre-perforated poly(dimethylsiloxane) structure (PDMS;

SylgardTM184, Dow Corning, Midland, MI, USA) and two

pulled and one normal glass pipette (outer diameter: 1mm;

inner diameter: 0.75mm). Along the perforated holes in the

PDMS structure, the intermediate, inner, and outer pipettes

were sequentially inserted, self-aligned, and fixed using

oxygen plasma. To produce the alginate hollow fiber, three

fluids were introduced simultaneously via three separate

inlets: a core fluid (100mM CaCl2), a sample fluid (2% w/w

sodium alginate (Sigma, St. Louis, MO, USA)), and sheath

fluid (100mM CaCl2). The hollow microfiber was fabricated in

two stages (Figure 1a). During the first stage, coaxial flow

(consisting of core and sample fluid) is generated and the

solidification process starts at the interface between the core

and the sample flow that forms along the inner wall of the

hollow fiber. During the second stage, sheath flow is

introduced and this meets the preformed coaxial flow from

the first stage. This results in the formation of a three-layered

coaxial flow that consists of core, sample, and sheath fluid.

small 2009, 5, No. 11, 1264–1268 � 2009 Wiley-VCH Verlag Gmb

Solidification occurs simultaneously at both interfacial layers

(between the core and sample, and between the sample and

sheath fluid) (inset of Figure 1a). The three-layered coaxial

flow moves through the outlet pipette and the solidified

alginate hollow fiber is generated continuously and extruded.

To visualize the fabrication of the alginate hollow fiber, the 2%

w/w alginate solution was mixed with a red dye (Papicel Red

IJ-F3B, Eastwell Co., Ltd, Seoul, Korea) (Figure 1b).With this

method, we were able to produce alginate hollow fibers that

were more than 1m in length (see the movie in the Supporting

Information).

Our microfluidic chip method allowed us to successfully

generate hollow fibers using all sorts of alginates. The drying

process used for scanning electron microscopy (SEM)

preparation led to slight distortion of the hollow fiber

(Figure 2a). The diameter of the alginate hollow fibers is

measured in the aqueous solution to prevent distortion using

optical microscopy, and Figure 2b illustrates that the diameter

is determined by the flow rates of the core, sample, and sheath

fluids. This figure indicates that the outer diameter of the fibers

increased in a flow rate-dependent manner of the sample flow

that varied from 0.01 to 0.05mLmin�1 for all sheath flow rates,

showing 103.5% increase on average when the minimum and

maximum sample flows were compared for each sheath flow.

In contrast, as the sheath flow rate increased from 0.38 to

0.83mLmin�1, the fiber diameter decreased 53.2% on average

for minimum and maximum sample flow rates. The diameter

of alginate hollow fiber can be estimated from

D ¼ Rg 1� QsþQsh�Qc

QT

� �12

!12

(1)

H & Co. KGaA, Weinheim www.small-journal.com 1265

communications

Figure 3. a) Red dye liquid movement through an alginate hollow fiber

embedded in agar gel (inset: liquid movement over time). b) Fluor-

escence micrograph of an BSA–FITC-laden hollow fiber (inset: 3D

intensity graph).

1266

where D is the diameter of the alginate hollow fiber, Rg is

the inner radius of the glass pipette, QT is the total flow rate,

Qs is the sample flow rate, Qsh is the sheath flow rate, and

Qc is the core flow rate (see Supporting Information,

Figure S1).

In ourmethod, exact alignment of core, sample, and sheath

flows is very important. We achieved good alignment by use of

a pipette that was pulled through the perforated fiber hole. In

particular, we wound the extruded hollow fibers on cover glass

(Figure 2c) by use of a homemade winding device (see the

movie in the Supporting Information). This result indicates

that it is feasible to align and handle weak alginate hollow

fibers that are extruded from the outlet pipette. This

technology could be used for creating scaffolds needed for

tissue engineering and for other applications.

The alginate hollow fibers with several different alginates

were fabricated and their ultimate strength was measured and

the result is plotted in Figure 2d. As shown in this figure, the

ultimate strength of alginate hollow fiber can be controlled by

the selection of an appropriate material. This mechanical

property may be important for the transportation of

pressurized liquid and for the easy handling and ability for

use as a supportive scaffold.[15]

Fluid transport is one of the major functions of fabricated

hollow fibers. To mimic blood vessels, we embedded hollow

fibers in an agar gel, and tested capillary force-induced fluid

transport by use of red dye solution. The inset of Figure 3a

illustrates the movement of the dye solution through the

embedded hollow fiber over time. In our system, the fluid flows

at about 30mms�1, somewhat faster than the speed through a

capillary vessel in a human body because our flow is driven by

capillary force. This system emulates blood flow through a

capillary that is embedded in tissue and can be used for in vitro

modeling of microcirculation, an important factor in many

cancer studies, and in studies of gas and nutrient delivery in

tissue.

To verify that biomolecules can be immobilized within

these fibers, we fabricated bovine serum albumin–fluorescein

isothiocyanate (BSA–FITC)-laden alginate hollow fibers.

Figure 3b indicates that our method allowed materials to be

loaded stably and uniformly within the hollow fibers. Using

our technique, numerous labile biomolecules (e.g., growth

factors, cytokines) could also be loaded into the layer of

these hollow fibers. Our results also indicate that this fiber

could be used as a ‘‘capillary reactor,’’ in which an enzyme

that is immobilized within the fiber catalyzes the reaction

of substrate(s) that flow through the fiber. Our fibers,

whose porosity and thickness are controlled, also have great

potential for studies of microcapillary effects, such as the

transportation of molecules or neutrophils through vascular

membranes.

Next, we investigated the feasibility of constructing 3D

microvascularized structures using our alginate hollow fibers.

For this purpose, we fabricated HIVE-78 cell-laden fibers and

embedded them into an AGF-hydrogel where smooth muscle

cells (HIVS-125) were cultured (Figure 4a). This structure,

which emulates a tissue with networked microvessels, was

cultured for 7 days. The relative rate of HIVE-78 and HIVS-

125 cell viability was evaluated by prelabeled CellTracker1

www.small-journal.com � 2009 Wiley-VCH Verlag Gm

(CFSE and CMPTX, respectively). There was an approxi-

mately 13.3% (95.6% and 82.3%) difference of viability rate

between the HIVE-78 and HIVS-125 cells as shown in

Figure 4e. The results indicate that both cell types have no

significant decrease during 7 days of cell culture. For better

visualization of embedded cells, we stained HIVE-78 and

HIVS-125 cells with red and green fluorescent dyes and

recorded images after 7 days of culturing (Figure 4b). The

HIVE-78 cells stayed aligned within the fiber structures that

were embedded in the HIVS-125-laden hydrogel. For

verification that our fibers maintained a hollow structure,

we also used 3D scanning confocal laser microscopy (Figure 4c

and d). These images indicate the presence of a perfectly

formed 3D vessel with HIVE-78 and HIVS-125 cells.

Morphogenesis of vascular structures requires cell–cell

interactions following paracrine signals and direct cell

contacts. By adding growth factors and bioactive peptides

bH & Co. KGaA, Weinheim small 2009, 5, No. 11, 1264–1268

Figure 4. a) Schematic of coculturing of HIVE-78-cell-laden alginate hollow fiber and HIVS-

125-cell-laden AGFmatrix to emulate vascularized tissue. b) Confocal image of the interaction

of HIVE-78 cells and HIVS-125 cells within the alginate hollow fiber and the fibronectin-

incorporated agar-gelatin complex hydrogel matrix. c) 3D images of cells that were cocultured

for 7 days. d) Orthogonal profiles of HIVE-78-cell-laden alginate hollow fiber on the complex

matrix. e) Viability of cocultured HIVE-78 and HIVS-125 cells over 7 days.

during the fiber fabrication process, we could stimulate the

cell–cell interaction as well as differentiation for the vascular

morphogenesis. For more qualitative evaluation of vessel

formation, measurement of releasing kinetics of the biomo-

lecules embedded in the hydrogel is required in vitro because

indirect cell–cell interaction is mediated by the cytokines

secreted from the cells. We measured the releasing profile of

growth factors from the alginate fiber for evaluating

transportation of growth factors to the surrounding matrix.

This simple method can be used for the future technologies for

creating biomimetic microenvironments.

We have shown that alginate hollow fibers can be produced

continuously by use of a microfluidic chip. Numerous

therapeutic materials can be easily loaded during the

fabrication process and the resulting fibers have great

potential for use in delivery of drugs and other therapeutic

materials. As an initial application of these hollow fibers, we

small 2009, 5, No. 11, 1264–1268 � 2009 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

emulated vascularized tissue by embedding

endothelial-cell-laden hollow fibers into

smooth muscle cell-laden AGF hydrogels

and successfully cocultured these cells. Our

microfabrication process is a promising

method for engineering vascular tissue

and for studying cell–cell interactions in

3D structures.

Experimental Section

Chip-based fabrication of alginate hollow

fibers: Changing the flow rate of the core,

sample, and sheath fluids controls the dia-

meter of the alginate hollow fiber. By using

several flow rates (sample flow rates: 0.01,

0.03, 0.05mLminS1; sheath flow rates: 0.38,

0.50, 0.66, 0.83mLminS1; fixed core flow

rate: 0.003mLminS1), we fabricated alginate

hollow fibers and measured their diameters

with optical microscopy (Axiovert 200M, Carl

Zeiss, Goettingen, Germany).

Ultimate strength test of alginate fiber: To

enhance themechanical strength, hollow fibers

synthesized with several commercially avail-

able alginate specimens (Protanal 10/60 LF

(FMC Biopolymer, Dramman, Norway), Manugel

DPB, GMB (Kelco International Ltd., London,

UK; used for high strength), Protanal 10/60 LS

(FMC Biopolymer, Dramman, Norway; used for

middle strength), Alginic acid sodium salt

(Sigma no. A2158, St. Louis, MO, USA; used

for low strength)) were fabricated and their

ultimate strength was measured using a digital

force gauge (IMADA, Toyohashi, Japan).

Fluid transport through alginate hollow

fibers: To mimic biological microvessels, we

embedded the hollow fiber into an agar gel

matrix (Sigma, St. Louis, MO, USA) (Figure 3a).

We slowly added the red dye solution, and the

solution moved upward via capillary force. The height of the

solution is determined by the inner diameter of the hollow fiber

and surface tension as follows

g ¼ 1

2rgrh (2)

where g is the surface tension of the liquid, r is the density of the

liquid, g is the acceleration due to gravity, r is the radius of the

tube, and h is the height of capillary rise. The induced fluid

transport was observed by use of the red dye solution and a digital

camcorder (Panasonic MX-500, Tokyo, Japan).

Production of protein-laden alginate hollow fibers: To demon-

strate the ability to load therapeutic materials into the alginate

hollow fibers, we fabricated a BSA–FITC-immobilized hollow fiber

by use of 2% w/w sodium alginate solution (1mL) with 1% w/w

diluted BSA–FITC (50mL). The BSA–FITC-immobilized fiber was

www.small-journal.com 1267

communications

1268

scanned with a confocal laser scanning microscope (LSM-510

META, Carl Zeiss, Goettingen, Germany).

Coculture of HIVE-78-cell-laden alginate hollow fibers with

HIVS-125-cell-laden hydrogel: We prepared an agar solution

(0.8% w/w) by dissolving agar in phosphate buffer saline (PBS)

(Gibco-BRL), autoclaving for 1 h, and adding 3% w/w gelatin

(Sigma, St. Louis, MO, USA) that was dissolved in PBS at 60 -Cwith a magnetic stir bar. Finally, we added 5mgmLS1 fibronectin

that was dissolved in PBS. The three solutions were mixed, and

then the HIVS-125 cells (106 cellsmLS1) were gently suspended to

create a composite hydrogel with uniform cell distribution

(preparation of cells is summarized in the Supporting informa-

tion). We spread 2mL of the pre-warmed AGF–HIVS-125 cell

mixture onto a 60 f cell culture dish (Corning, NY, USA). The HIVE-

78 cell-laden alginate hollow fibers (2.5 �105 cellsmLS1; fiber

length: 40mm, inner diameter: 0.3mm) were placed on the HIVE-

125 AGF matrix (Figure 4a). In addition, we spread 3mL of the

HIVE-125-cell-laden AGF mixture and allowed it to solidify on a

clean bench at room temperature for 1 h. Then, 3mL of the F12K

medium was added to the matrix surface. This resulted in a disk-

sandwich shape, and 15-mm-thick alginate hollow fibers that

were embedded in an AGF-matrix-based biomimetic coculture

system. The specimens were maintained at 37 -C and 5% CO2 for

7 days (Figure 1b). For visualization, HIVS-125 cells were

prelabeled with the CellTracker1 CFSE green fluorescent dye

(Molecular Probes1, Invitrogen, Paisley, UK), and the HIVS-78

cells were prelabeled with a CellTracker1 CMPTX red (Molecular

Probes). Fluorescence micrographs were taken with a fluorescence

microscope (Axiovert 200M, Carl Zeiss, Goettingen, Germany).

www.small-journal.com � 2009 Wiley-VCH Verlag Gm

Keywords:biomaterials . cocultures . fibers . microfluidic chips . tissueengineering

[1] A. Khademhosseini, R. Langer, J. Borenstein, J. P. Vacanti, Proc.

Natl. Acad. Sci. USA 2006, 103, 2480.[2] S. L. Tao, T. A. Desai, Adv. Drug Delivery Rev. 2003, 55, 315.[3] K. Y. Suh, A. Khademhosseini, P. J. Yoo, R. Langer, Biomed.

Microdevices 2004, 6, 223.[4] W. Nijdam, J. de Jong, C. J. M. van Rijn, T. Visser, L. Versteeg, G.

Kapantaidakis, G. H. Koops, M.Wessling, J. Membr. Sci. 2005, 256,209.

[5] A. Saito, Nephrology (Carlton) 2003, 8, S10.[6] M. J. Ellis, J. B. Chaudhuri, Biotechnol. Bioeng. 2007, 96, 177.[7] W. Jeong, J. Kim, S. Kim, S. Lee, G. Mensing, D. J. Beebe, Lab Chip

2004, 4, 576.[8] S. J. Shin, J. Y. Park, J. Y. Lee, H. Park, Y. D. Park, K. B. Lee, C. M.

Whang, S. H. Lee, Langmuir 2007, 23, 9104.[9] S. Sugiura, T. Oda, Y. Aoyagi, M. Satake, N. Ohkohchi, M. Naka-

jima, Lab Chip 2008, 8, 1255.[10] N. A. Peppas, J. Z. Hilt, A. Khademhosseini, R. Langer, Adv. Mater.

2006, 18, 1345.[11] A. D. Augst, H. J. Kong, D. J. Mooney,Macromol. Biosci. 2006, 6, 623.[12] J. L. Drury, D. J. Mooney, Biomaterials 2003, 24, 4337.[13] V. L. Tsang, S. N. Bhatia, Adv. Drug Delivery Rev. 2004, 56, 1635.[14] E. Amici, G. Tetradis-Meris, P. de Torres, F. Jousse, Food Hydro-

colloids 2008, 22, 97.[15] R. H. Schmedlen, K. S.Masters, J. L.West,Biomaterials2002,23, 4325.

bH & Co. KGaA, Weinheim

Received: October 30, 2008Revised: December 26, 2008Published online: March 19, 2009

small 2009, 5, No. 11, 1264–1268