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Biomaterials 20 (1999) 1133 } 1142

Porous chitosan sca ! olds for tissue engineeringSundararajan V. Madihally, Howard W.T. Matthew *

Department of Chemical Engineering & Materials Science, Wayne State Uni versity, 5050 Anthony Wayne Dri ve, Detroit, MI 48202, USA

Received 9 November 1998; accepted 13 January 1999

Abstract

The wide array of tissue engineering applications exacerbates the need for biodegradable materials with broad potential. Chitosan,the partially deacetylated derivative of chitin, may be one such material. In this study, we examined the use of chitosan for formation

of porous sca ! olds of controlled microstructure in several tissue-relevant geometries. Porous chitosan materials were prepared bycontrolled freezing and lyophilization of chitosan solutions and gels. The materials were characterized via light and scanning electronmicroscopy as well as tensile testing. The sca ! olds formed included porous membranes, blocks, tubes and beads. Mean pore diameterscould be controlled within the range 1 }250 m, by varying the freezing conditions. Freshly lyophilized chitosan sca ! olds could betreated with glycosaminoglycans to form ionic complex materials which retained the original pore structure. Chitosan sca ! olds couldbe rehydrated via an ethanol series to avoid the sti ! ening caused by rehydration in basic solutions. Hydrated porous chitosanmembranes were at least twice as extensible as non-porous chitosan membranes, but their elastic moduli and tensile strengths wereabout tenfold lower than non-porous controls. The methods and structures described here provide a starting point for the design andfabrication of a family of polysaccharide based sca ! old materials with potentially broad applicability. 1999 Elsevier Science Ltd.All rights reserved

Keywords: Chitosan; Lyophilization; Porous microstructure; Sca ! old; Elastic modulus

1. Introduction

The tissue engineering approach to repair and regen-eration is founded upon the use of polymer sca ! oldswhich serve to support, reinforce and in some casesorganize the regenerating tissue [1 } 6]. The sca ! old maybe required to release bioactive substances at a control-led rate or to directly in #uence the behavior of incorpo-rated or ingrowing cells. Furthermore, many application

scenarios call for the use of biodegradable polymericmatrices or matrices which are at least amenable tointegration with growing tissue. Performance of thesevaried functions usually demands a porous sca ! oldmicrostructure, with the porosity characteristics beingapplication speci " c. Desirable aspects of sca ! old chem-istry may include speci " c interaction with, or mimicry of,extracellular matrix components, growth factors, or cellsurface receptors. Likewise, surface display of reactive oreasily derivatized groups may be useful for some applica-tions.

* Corresponding author. Tel.: 001 313 577 5238; fax: 001 313 5773810; e-mail: h.matthew @wayne.edu

A number of natural and synthetic polymers are cur-rently being employed as tissue sca ! olds [7 }10]. Themicrostructures of these systems span the range fromhydrogels, to open-pore structures, to " brous matrices.Since the range of potential tissue engineered systems isbroad, there is a continuous ongoing search for materialswhich either possess particularly desirable tissue-speci " cproperties, or which may have broad applicability andcan be tailored to several tissue systems. The amino

polysaccharide chitosan (poly 1,4D

-glucosamine) may beone such broadly applicable material. Chitosan [11] isa partially deacetylated derivative of chitin [12, 13], theprimary structural polymer in arthropod exoskeletons.Depending on the source and preparation procedure,molecular weight may range from 300 kD to over 1000 kD.Commercially available preparations have degrees of deacetylation ranging from 50 to 90%. Chitosan is a crys-talline polysaccharide and is normally insoluble in aque-ous solutions above pH 7. However, in dilute acids(pH ( 6), the free amino groups are protonated and themolecule becomes soluble (Fig. 1). This pH-dependent

solubility provides a convenient mechanism for process-ing under mild conditions. The high charge densityin solution allows chitosan to form insoluble ionic

0142-9612/99/$- see front matter 1999 Elsevier Science Ltd. All rights reserved.PII: S 0 1 4 2 - 9 6 1 2 ( 9 8 ) 0 0 0 1 1 - 3


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Fig. 1. Chitosan molecular structure. For this study the fraction of acetylated amino groups was 10%.

complexes with a wide variety of water soluble polyan-ionic species. Complex formation has been documentedwith anionic polysaccharides such as heparin and alginicacid, as well as synthetic polyanions like poly(acrylicacid) [14 }19]. While these interactions have previouslybeen used to form microcapsules and membranes for cellculture and transplantation, they may also providea simple mechanism for modifying the surface and bulkproperties of chitosan structures.

Chitosan and some of its complexes have been studiedfor use in a number of biomedical applications. Theseinclude wound dressings [19], drug delivery systems[20 }22] and space " lling implants [23, 24]. However,little has been done to explore use of chitosan within thetissue engineering paradigm. In this study, we describethe preparation and characterization of porous chitosansca ! olds in a number of con " gurations. The structuresdescribed provide a starting point for use of this versatilematerial in engineered tissues.

2. Materials and methods

Chitosan (85 }90% deacetylated) was obtained fromCarbomer Inc. Heparin USP solution (100 U/ml in 0.9%NaCl) was purchased from Elkins-Sinn, NJ. Unlessotherwise stated all other reagents were obtained fromFischer Scienti " c.

2.1. Chitosan sca w old formation

Chitosan solutions with concentrations of 1, 2 or

3 wt% were prepared by dissolution in 0.2M

acetic acid.Bulk chitosan sca ! old samples were prepared by freezingand lyophilizing chitosan solutions in pre-cooled, #atbottomed glass tubes. The sample tubes employed had aninner diameter of 1.5 cm and a 1.2 mm wall thickness.Freezing was accomplished by immersing the tubes, con-taining 3 }5 ml of solution, in freezing baths maintainedat ! 203C, ! 783C or ! 1963C. The samples were thenlyophilized until dry. Planar sca ! olds were formed byfreezing 25 }50 ml of chitosan solution in 10 cm diameterpolystyrene petri dishes. The dishes were held with theirbottom surfaces in contact with either liquid nitrogen or

a dry ice slab to form pores perpendicular to the sampleplane. Alternatively, pores approximately parallel to thesample plane were formed by " lling a 5 ; 50 ; 100 mm

( ; = ; H ) polystyrene mold with chitosan solution fol-lowed by slow immersion into liquid nitrogen. Afterfreezing, molds were opened to expose one frozen surfaceand then lyophilized.

2.2. Microstructural characterization

Lyophilized sca ! olds were sectioned at various planes,attached to sample stubs with conductive paint and sput-ter-coated with palladium prior to examination undera Hitachi S-2400 Scanning Electron Microscope (SEM).Imaging was conducted at an accelerating voltage of 12}15 kV. Mean pore diameters were determined by im-age analysis (Sigma Scan Pro, Jandel Scienti " c) of lightmicrographs of thin sections from para $ n embeddedsca ! olds. In some cases, pore diameter estimates wereobtained by analysis of digital SEM images from sec-tioned samples.

2.3. Porous microcarriers

Directly frozen chitosan microcarriers were preparedby extruding droplets of a 1 wt% chitosan solution froma 23 gauge needle and collecting the droplets in a stirredbeaker of either liquid nitrogen or dry ice cooled methyl-ene chloride. Temperatures above ! 783C were obtainedby varying the thickness of insulation between themethylene chloride container and the dry ice.

For preparing gelled microcarriers, droplets were " rstcollected in stirred 0.1 M NaOH to induce gellation. Afterrinsing brie #y with water, the gelled chitosan beads werethen transferred to the freezing bath. Both gelled anddirectly frozen microcarriers were cooled to ! 783C,lyophilized and subsequently examined under the SEM.

2.4. Tubular sca w olds

Porous tubular sca ! olds were formed by freezinga chitosan solution in the annular space between concen-tric silicone or PTFE tubes. Chitosan solution of 1 or2 wt% concentration was injected into the annular space

and the whole assembly was frozen by direct contact withdry ice (! 783C). The outer tube was then removed andthe assembly was lyophilized.

To form porous tubes bearing a non-porous luminalmembrane, an inner silicone tube was " rst coated witha chitosan " lm by dipping into a 2% chitosan solution.The " lm was gelled by brief immersion in aqueous 30%ammonia solution and then air dried. Finally, the annu-lar mold was assembled using the coated inner tube, " lledwith chitosan solution and processed as described above.

2.5. Sca w old rehydration

Lyophilized sca ! olds were rehydrated and stabilized ineither dilute NaOH or an ethanol series. For the NaOH

1134 S.V. Madihally, H.W.T. Matthew / Biomaterials 20 (1999) 1133 } 1142


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process, samples were slowly immersed in 0.05 M NaOH,equilibrated for & 10 min and then washed twice withwater and twice with phosphate bu ! ered saline (pH 7.4).In the ethanol process, sca ! olds were immersed in abso-lute ethanol for & 1 h and then sequentially in 70% (v/v)and 50% ethanol for & 30 min each. Sca ! olds were" nally equilibrated with PBS prior to mechanical testing.

2.6. Mechanical testing of planar sca w olds and control membranes

Rectangular test samples (1 cm ; 5 cm) were cut fromethanol rehydrated planar sca ! olds and strained to failureon an MTS Bionix 100 materials testing system. A con-stant strain rate of 0.4 min \ was used. Results werecompared with similarly tested non-porous chitosan mem-branes. The non-porous membranes were formed by " rstgelling 25 ml of 2% chitosan solution in a polystyrene

petri dish using vapor phase ammonia. The gel was thenair dried under room temperature air #ow and rehydratedusing the ethanol process described above.

2.7. Heparin }chitosan complexes

Chitosan sca ! olds were complexed with heparin viaionic interaction by rehydrating the freshly lyophilizedsca ! olds in a heparin USP solution (100 U/ml +0.7 mg/ml) for 2 h. The hydrated complex was then rinsedwith water and either air or vacuum dried. The " nal driedsca ! olds were sectioned and examined by SEM.

3. Results

Our studies were conducted with the goal of develop-ing procedures by which porous sca ! old structures couldbe generated from chitosan. The speci " c sca! old geo-metry required is of course application-speci " c. Below,we describe a number of sca ! old systems which may beapplicable to several types of engineered tissues.

3.1. Bulk sca w oldsBulk sca ! olds were formed by the simple procedure of

freezing a chitosan solution in a suitable vessel and sub-sequently lyophilizing the frozen structure. Figure 2shows SEM micrographs of the fractured central regionof one such sca ! old prepared in a cylindrical glass tube.The freezing and lyophilization process generated anopen pore microstructure with a high degree of intercon-nectivity. Because of surface cooling in the cylindricalgeometry, the pores were radially oriented. The micro-graphs also show that two levels of porosity were induced

in the liquid nitrogen frozen sample. A 20 m thickregion adjacent to the vessel wall was composed of short,highly connected pores, 5 }10 m in diameter. In contrast,

sample regions further from the wall consisted of elon-gated, larger diameter pores ( ' 50 m). This two-phasepore structure is most likely the result of the di ! erence inice nucleation conditions at the solution }glass interface,versus secondary nucleation conditions at the solution }ice interface. Bulk sca ! olds were also prepared ina planar geometry by freezing chitosan solutions in shal-low dishes. Figure 3 shows a sectional view of a sca ! oldprepared by freezing chitosan in a shallow dish, using dryice as the freezing agent. This produced a planar sca ! old& 5 mm thick with perpendicularly oriented, thin walledpores. The pores were fairly uniform and parallel witha polygonal cross section. In this geometry, lateral poreconnectivity appeared to be much lower than for cylin-drical sca ! olds. This result was likely due to the highlyparallel ice crystal growth, which in turn was caused bythe strongly one-dimensional nature of the thermalgradients established during freezing. Under these condi-

tions, the probability of branching and abuttment of growing crystals (a major contributor to lateral poreconnectivity) would be low. In contrast, the radial ther-mal gradients established during freezing of cylindricalsca ! olds produced numerous crystal impacts duringgrowth, thus increasing lateral connectivity. Pore con-nectivity was also in #uenced by the concentration of thechitosan solution. Under " xed freezing conditions, highersolution concentrations produced lower connectivity.

3.2. E w ect of freezing temperature on pore microstructure

Cylindrical sca ! olds formed by freezing at varioustemperatures were embedded in para $ n and sectionedlongitudinally at planes approximately midway betweenthe sample surface and center. Microscopic images of areas perpendicular to the pore direction were capturedand analyzed to evaluate the mean pore diameter. Asshown in Fig. 4, the mean pore diameter could be con-trolled within the range 40 }250 m, by varying thefreezing temperature and hence the cooling rate. A lesser

e! ect of chitosan solution concentration was alsoobserved, with smaller pores being formed at higherconcentrations. Since ice crystal growth and hence porediameter are functions of the temperature gradient, porediameters can be expected to vary with radial position incylindrical sca ! olds. In fact, pore diameter increasedsigni " cantly moving from the edge towards the center of the sample. In general, this variation with position couldbe minimized by limiting sca ! old thickness.

3.3. Porous microcarriers: e w ects of solution versus gel freezing

Porous chitosan beads or microcarriers could beformed by either directly freezing droplets of chitosan

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Fig. 2. SEM images of a cylindrical bulk sca ! old. Chitosan solution (2 wt%) was frozen in a pre-cooled, 1.5 cm ID, cylindrical glass tube by immersionin liquid nitrogen. After lyophilization, the cylindrical sample was fractured transversely and processed for SEM imaging. (a) Low-magni " cation viewof fracture zone showing surface and interior pore structure. (b) Higher-magni " cation view of fracture zone. (c) High-magni " cation view of porestructure at glass-contacting surface.

Fig. 3. SEM image of a bulk sca ! old frozen in a planar geometry.A 2% chitosan solution was frozen in a shallow polystyrene dish by

contact with dry ice. The lyophilized sample was sectioned parallel tothe sample plane.

Fig. 4. E ! ects of freezing temperature and chitosan concentration onmean pore diameter of cylindrical chitosan sca ! olds. Sca ! olds werepara $ n embedded and thin longitudinal sections were obtained.Microscopic images of the sections were analyzed to determine the areamean diameter. Error bars are standard deviations calculated froma minimum of 30 pores.



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Fig. 5. SEM micrographs of porous chitosan microcarriers. Micrographs show low- and high-magni " cation views of microcarriers formed by variousfreezing protocols: (a, b) chitosan droplets directly frozen in liquid nitrogen; (c, d) interior views of directly frozen microcarriers; (e, f) chitosan gel beadsdirectly frozen in liquid nitrogen; (g, h) chitosan gel beads directly frozen in methylene chloride at ! 53C.

solution or by freezing gelled chitosan beads. Both tech-niques produced porous beads, but with drastically dif-ferent pore microstructures. Figure 5a } c illustrate thesurface and interior pore structures of beads formed by

directly freezing chitosan droplets in liquid nitrogen. Thesurfaces of the spherical beads exhibited pores rangingfrom 20 to 50 m in diameter (Fig. 5a and b). In manycases, pores were arranged in parallel arrays along ridges



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Fig 5. (continued ).

or grooves in the bead surface. Fractured beads (Fig. 5c)revealed interior pores arranged in bundles with a gen-eral radial orientation. Within a given bundle, pore dia-meter was fairly uniform, though signi " cant di ! erencesbetween bundles were seen. Figure 5d shows one suchbundle at higher magni " cation.

As an alternative formation method, chitosan dropletswere gelled in NaOH before freezing. Figure 5e showsa low-magni " cation SEM image of a liquid-nitrogen-frozen bead. The bead surface appeared macroscopicallysmooth except for a few expansion induced cracks. How-ever high magni " cation (Fig. 5f) revealed a highly por-ous, " brous microstructure with e ! ective pore diametersin the 1 }5 m range. The " brous structure suggests thatthe rapid liquid nitrogen freezing produced very small icecrystals which e ! ectively immobilized and maintainedthe chitosan gel network. In contrast, Gel beads frozen ina methylene chloride bath at ! 53C exhibited surfacepores which could be more accurately described as sur-face depressions (Fig. 5g and h). This together with thelack of any semblance of a " brous structure suggestedthat nucleation occurred external to the gel, and that

subsequent ice crystal growth served to collapse and fusethe gel " ber structure. In keeping with this hypothesis,SEM imaging of sectioned beads con " rmed that theinterior was mainly dense chitosan with few pores.

Figure 6 summarizes the e ! ects of freezing temperatureon the mean surface pore diameter of chitosan microcar-riers. Taken as a whole, it is clear that the e ! ect of temperature on pore size is much greater between ! 53Cto ! 153C than between ! 153C and ! 1963C. Sincecrystal growth and hence pore size are functions of bothheat and mass transfer rates, this result suggests than atthe lower temperatures, the system is dominated by mass

transfer rates and pore size becomes essentially indepen-dent of the heat transfer rate and hence the freezingtemperature.

Fig. 6. E ! ects of freezing protocol on the surface pore diameters of chitosan microcarriers. Mean diameter values were determined byimage analysis of SEM micrographs. Error bars are standard devia-tions determined from a minimum of 30 pores.

3.4. Tubular sca w olds

A number of tissue engineering applications require

use of tubular constructs to mimic a particular tissuegeometry. We explored the feasibility of using chitosanfor such applications by generating porous tubular scaf-folds both with and without a luminal membrane. Micro-graphs of each type are shown in Fig. 7. Figure 7a showsa section of a thick walled conduit 3.5 mm in diameterwith a 1 mm wall thickness. The open pore structure of this sample, while essentially radial in orientation, showsa high level of connectivity (Fig. 7b). In contrast Fig. 7cshows a sectional view of the wall of a second generationconduit with a thin luminal membrane. The sample wallis & 100 m thick with a clear radial pore orientation.

The luminal membrane thickness in this case was lessthan 10 m, but could be controlled by forming multi-layer membranes. Luminal membranes up to at least



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Fig. 7. SEM micrographs of tubular sca ! olds. Sca ! olds were formed by freezing and lyophilizing a 2% chitosan solution in annular molds. Freezingwas accomplished by direct contact with dry ice. (a, b) Segment of thick-walled, completely porous tube. (c) Section of thin- walled tube with

non-porous luminal membrane.

50 m thick can be routinely formed by the chitosangellation }dehydration process.

3.5. Sca w old rehydration and mechanical properties

The sca ! olds described above were imaged in the de-hydrated state. Newly lyophilized sca ! olds were sti ! andinelastic. If such samples were rehydrated in a neutralaqueous medium, the chitosan exhibited rapid swelling

and ultimately dissolved. This indicated that thelyophilized structure was composed of soluble chitosanacetate. Sca ! old dissolution could be prevented by re-

hydrating samples in either dilute NaOH (0.1 M ), or in anethanol series such as 100, 70, 50, 0%. At each stage thesamples were equilibrated with the solution for at least30 min. Sca ! olds hydrated in NaOH exhibited someshrinkage and distortion, probably caused by base-in-duced changes in crystallinity and associated structuralstresses [25]. These changes were only partially reversedupon transfer to a pH 7 aqueous solution. On the otherhand, samples hydrated through an ethanol series exhib-

ited no signi " cant volume or shape changes. The base-induced shrinkage also resulted in numerous entrappedair bubbles. However, the low surface tension of the



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absolute alcohol allowed complete hydration of thesample with few entrapped bubbles. All entrapped bub-bles could be removed from ethanol hydrated samples bysubjecting the samples to brief vacuum treatment whilesubmerged in absolute ethanol. The alcohol hydra-tion protocol has the added advantage of allowingdirect sterilization of samples in 70% ethanol. Suchsterilized sca ! olds could be equilibrated with culturemedium or PBS for several hours prior to any cell seed-ing or in vivo implantation activity.

Hydrated chitosan sca ! olds were soft, spongy and very#exible. But overall strength was generally low. Figure 8illustrates typical stress }strain plots for a non-porouschitosan membrane and a porous planar sca ! old withperpendicular pores. Both materials were tested in thehydrated state. While non-porous chitosan membranesexhibited a maximum strain of 30 } 40%, the poroussca ! olds were more extensible, with maximum strain

ranging from&

30 to&

110% as a function of poreorientation and pore diameter (data not shown).

3.6. Polyanionic modi xcation of chitosan sca w olds

The choice of chitosan as a sca ! old material waspartially governed by the multiple modes by which thematerial 's surface or bulk structure could be derivatizedor chemically modi " ed. One of the simplest modi " cationapproaches for chitosan involves forming ionic com-plexes with polyanionic species. Our interest inglycosaminoglycans (GAGs) led to an examination of methods for complexing chitosan sca ! olds with thesepolyanions. Rehydration of planar sca ! olds in 0.7%heparin e ! ectively converted the chitosan acetate sca ! oldto a chitosan }GAG complex structure. Signi " cant swell-

Fig. 8. Stress }strain plots of typical non-porous and porous chitosanmembranes. The inset shows an expanded view of the low stress portionof the graph. Samples were strained to failure at a constant strain rate of 0.4 min \ .

ing of the material was observed during rehydration, butSEM examination of subsequently dehydrated samplescon " rmed that the internal porous microstructure wasretained. Interestingly though, the outer surfaces of thesca ! old appeared partially sealed in some regions by anionic complex membrane (Fig. 9a). Removal of the mem-brane revealed the porous structure beneath (Fig. 9b).We also noted that previously smooth pore walls nowappeared signi " cantly more textured after the GAGtreatment. Preliminary work with dense chitosan mem-branes cast in dishes showed that GAG complexation

Fig. 9. SEM micrographs of heparin }chitosan complex sca ! old.(a) Surface view of sca ! old showing continuous membrane. (b) Viewof sca ! old with surface layer removed. Note increased roughness of pore surfaces compared to Fig. 3.



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caused dimensional changes and extensive wrinkling of the complex membrane. A similar swelling and wrinklingmechanism was probably responsible for the increasedtexture of the pore walls.

4. Discussion

The explosion in tissue engineering research hasaccentuated the need for new classes of biodegradablepolymers with the potential for speci " c or controllablebioactivity. In this paper we have described proceduresfor fabricating tissue sca ! olds from chitosan, an enzym-atically degradable polysaccharide with broad potential.The polymer 's hydroxyl and amino groups provide sev-eral possibilities for derivatization or grafting of desirablebioactive groups, and chitosan 's pH-dependent solubilityallows use of relatively mild processing methods. This

feature is particularly important if incorporation of bio-active species is desired prior to forming a " nal threedimensional microstructure. Furthermore, the nitrousacid-based procedure for precise stoichiometric chaincleavage [26] provides a mechanism by which molecularweight, crystallinity and ultimately mechanical proper-ties can be additionally controlled. Formation of densechitosan membranes and extruded " bers has been exten-sively described and characterized elsewhere [25, 35],and provide alternate approaches for biomedical implantfabrication.

Chitosan 's precursor, chitin, is the major structuralmolecule in arthropod cuticles. In that environment, in-teractions with matrix proteins and connective tissuecomponents are numerous and undoubtedly intimate.Partial deacetylation of the molecule to form chitosancreates the potential for a variety of interactions in themammalian implant environment. However, with theexception of collagen [27], interactions with mammalianproteins have not been extensively characterized. How-ever, many interesting interactions with mammaliantissues have been reported. The cells involved rangedfrom osteoblasts and " broblasts to macrophages and

keratinocytes [23, 24, 28 }32]. In most cases the cellularinteractions have been positive from the tissue repair andregeneration standpoint.

Chitosan 's cationic nature also allows for pH-depen-dent electrostatic interactions with anionic glycosamino-glycans, proteoglycans and other negatively chargedspecies. These ionic interactions may serve as a mecha-nism for retaining or accumulating these moleculeswithin a tissue sca ! old during colonization or after im-plantation. Since a large family of growth factors andcytokines are known to be bound and modulated byGAGs, (in particular heparin and heparan sulfate) a scaf-

fold incorporating a chitosan }GAG complex may pro-vide a means of retaining and concentrating desirablefactors secreted by colonizing cells. Such a system may

even be capable of recruiting desirable growth factorsfrom surrounding tissue #uids.

A number of researchers have examined the tissueresponse to chitosan-based implants [16, 23, 24, 28, 29].In general, the material has been found to evoke a mini-mal foreign body reaction and is thus considered to be&biocompatible '. Implants have been shown to degrademainly through lysozyme-mediated hydrolysis, with thedegradation rate being inversely related to the degree of crystallinity. The crystallinity of the chitin }chitosan fam-ily of materials exhibits a minimum at intermediate levelsof deacetylation with chitin and fully deacetylatedchitosan showing the highest levels of crystallinity. Infact, the low degradation rate of highly deacetylatedchitosan implants is believed to be due to the inability of hydrolytic enzymes to penetrate the crystalline micro-structure. This issue has been addressed by derivatizingthe molecule with side chains of various types [33, 34].

Such treatments alter chain packing and increase theamorphous fraction, thus allowing more rapid degrada-tion. They also inherently a ! ect both the mechanical andsolubility properties.

The porous materials described in this paper providestarting points for the development and optimization of a variety of tissue sca ! olds and regeneration aids. Bulksca ! olds can be formed in speci " c shapes by using suit-able molds. Alternatively, a cylindrical or cuboidal scaf-fold could be cut and trimmed to a speci " c size andshape. This approach may be used to form colonizableconstructs for space " lling implants. In contrast, planarsca ! olds as described here may be applicable to mainlytwo-dimensional tissues such as skin or articular carti-lage. Porous beads or microcarriers may " nd applicationin bioreactors for cell expansion or metabolic function.Microcarrier aggregates or slurries may also be used asconformal space " lling implants for soft tissue appli-cations, or as sca ! olds for assembling organ-like neo-tissues in atypical locations. Finally, tubular sca ! oldsmay be useful for design of tissue systems requiringa tubular geometry. Included in this group are bloodvessels, the gastrointestinal tract and tissues with ducts or

multi-tube microstructures.Control of sca ! old pore morphology is critical forcontrolling cellular colonization rates and organizationwithin an engineered tissue. Furthermore, angiogenesis isa requirement for some sca ! old application scenariosand can be grossly a ! ected by material porosity. Poremorphology can also be expected to signi " cantly a ! ectsca ! old degradation kinetics and the mechanical proper-ties of the developing tissue. While controlled ratefreezing is limited by the magnitude and directionality of thermal gradients, it does provide a simple, straightfor-ward and reproducible way of introducing directional

pores into a polymer structure. The methods detailedhere allow optimization of pore morphology over aphysiologically relevant range and serve as an additional



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method of tailoring sca ! old properties for particulartissue requirements.

In conclusion, the studies described in this paper andelsewhere indicate that chitosan has excellent potential asa structural base material for a variety of engineeredtissue systems. Porous structures could be easily fab-ricated with control over pore morphology. The chemicalnature of the molecule also provides many possibilitiesfor covalent and ionic modi " cations which in turn allowextensive adjustment of mechanical and biological prop-erties.

Acknowledgements

The authors wish to thank Ms. Sonia Garg andMr. Goutam Reddy for their assistance in mechanicalproperty measurements. This work was supported by

grants from the National Science Foundation (CAREER:BES-9624151) and the Whitaker Foundation for Bio-medical Engineering (97-0361).

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