tissue engineering'. in: encyclopedia of polymer science...

Vol. 12 TISSUE ENGINEERING 261

TISSUE ENGINEERING

Introduction

Accidents and diseases lead to devastating tissue losses and organ failures, whichresult in more than 8 million surgical operations each year in the United Statesalone (1). These problems convert to a national annual healthcare cost of approx-imately half trillion U.S. dollars. The state-of-the-art clinical therapies to tissuelosses and organ failures can be categorized into three approaches, ie, transplanta-tion, surgical reconstruction, and the use of prostheses. Each of these approacheshas contributed to solving or alleviating the severity of these clinical problems,while all of them have serious limitations.

Organ transplantation became successful in the early 1960s because of thesuccess of immunologic suppression in the clinical setting (2). Transplantationhas saved, and is continuing to save, countless lives. This approach, however, isseverely limited by the dearth of donor organs. For example, fewer than 3000available donors are way shorter than the needs of approximately 30,000 Amer-icans for liver transplants each year (3). Surgical reconstruction utilizes tissuesharvested from the patient to rebuild a critically needed body part. The use of thepatient’s own tissue is advantageous in that it has a higher success rate resultingfrom the avoidance of immune rejection than using tissues from other sources.However, the need for second site of surgery, limited supply, inadequate size andshape, and the morbidity associated with donor site are all major concerns (4,5).In response to the shortages of needed tissues and organs, prostheses are devel-oped to replace certain body parts for their structural and mechanical functions.There are approximately 100,000 people in the United States with transplants,while there are more than 10,000,000 with biomedical implants (6). However, theprostheses are made from artificial materials, are not biologically functional, andare therefore subject to long-term complications and rejections.

Tissue engineering is a new approach to resolve the missing tissue and organproblems. Tissue engineering has been defined as an interdisciplinary field thatapplies the principles of engineering and the life sciences toward the developmentof biological substitutes that restore, maintain, or improve tissue function (3).There are three strategies in tissue engineering (3,7):

(1) The use of isolated cells or cell substitutes to replace those cells that supplythe needed function, including genetic or other manipulations before thecell infusion (8);

(2) The delivery of tissue-inducing substances, such as growth and differentia-tion factors, to targeted locations (9,10);

(3) Growing cells in three-dimensional (3-D) matrices (scaffolds) or devices,where cells can be either recruited from the host tissues in vivo or seeded(encapsulated) in vitro (3).

The advantage of the use of isolated cells is the simplicity. Cells are oftendirectly injected into the targeted locations to avoid complex procedures and as-sociated complications. The disadvantages include cell death and loss of function

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262 TISSUE ENGINEERING Vol. 12

because cells need the right nutritional and substrate environment to grow andfunction. The success of using tissue-inducing substances is dependent on the eco-nomical large-scale production and purification of the bioactive molecules, and onthe development of delivery systems that can deliver these molecules with thedesired profiles. Certain gene therapy techniques can also be used for this ap-proach. The above two approaches, ie, the use of isolated cells or tissue-inducingsubstances, are considered when the defects are very small and well contained.For engineering tissues of practical size scale and with predetermined 3-D struc-tures, these two approaches are seriously limited. Therefore, the third approach,ie, growing cells in 3-D matrices (scaffolds) or devices, has become increasinglyactive.

Some of these matrices or devices are used to develop immobilized cell sys-tems, either as implants or extracorporeal devices (11,12). The core of this technol-ogy is semipermeable membranes or matrices that have well-defined molecularweight or size cutoff (13,14). They serve as immunoprotective barriers to supportcell growth and function, which allow nutrients, metabolic products, and wastesto diffuse through, but not immune cells or antibodies. These devices offer certainbiological functions but are not living tissue/organ replacements (see MEMBRANE

TECHNOLOGY). Some other 3-D matrices work as templates (called scaffolds) toguide cells to grow, synthesize biological molecules and extracellular matrix com-ponents, and facilitate the organization and formation of functional tissues andorgans (15–17). After fulfilling the templating function, the scaffolds degrade anddisappear, leaving nothing foreign to the biological system (Fig. 1). There aremany additional advantages in this approach. Patient-derived cells (stem cells ordifferentiated cells) or future universal cell sources (nonimmunogenic) can be usedso that there will be minimum complication associated with immune rejections.These cells can be expanded in vitro to solve the donor shortage limitations. Anytissue/organ structure can be potentially mimicked by the scaffolding design. Theengineered tissues will have the capacity of growing, modeling, and remodeling inconcert with dynamic changes in physiological environment of the body. In this ap-proach, biodegradable polymers (natural or synthetic) are the materials of choice.Polymers (or macromolecules) are currently used as scaffolds for nearly every tis-sue type including bone and other mineralized tissues. Besides polymers, onlylimited inorganic materials are used for certain mineralized tissue engineeringresearch.

Although growing cells on two-dimensional (2-D) substrates (such as petridishes and culture plates) dates back centuries, designing 3-D scaffolds for tissueengineering is a new field, where polymer science and engineering play pivotalroles.

There are a few basic requirements that have been widely accepted for de-signing polymer scaffolds. First, the scaffold has to have high porosity and properpore size. These pores allow cell seeding and migration to achieve the neededrelative uniform distribution. The pores also provide the space for cell prolifera-tion and neo tissue deposition. The pores also satisfy the needed mass transportrequirements for nutrients, signaling molecules, metabolic products, and wastes.The porous structure should also allow for vascularization and innervation forsustained function of the regenerated tissues when implanted in vivo. High poros-ity is also beneficial because of the reduced polymer amount and its degradation


Fig. 1. Schematic diagram showing the tissue engineering concept. Scaffolding materials(temporary synthetic extracellular matrices) are designed, on which mammalian cells cangrow to regenerate the needed tissue or organ in three dimensions (3-D). Because thescaffolds are biodegradable, they will resorb after fulfilling the template function and leavenothing foreign in the body.

products in the neo tissues, which can provoke unwanted inflammatory responsefrom the host. Second, a high surface area (a.k.a. surface-to-volume ratio) isneeded, which is the 3-D surface area of the porous scaffolds (not just the out-side surface area). Many cell types are anchorage-dependent, ie, they can survive,grow, and function only when they are attached to an appropriate substrate. Thehigh surface area provides cells with the sufficient area to attach, grow, and de-posit neo tissue components. Third, biodegradability is generally required, and aproper degradation rate is needed to match the neo tissue formation. If the scaf-fold degrades too fast, it can collapse before the new tissue is formed so that itfails to serve as a 3-D guidance for the neo tissue organization. If the scaffolddegrades too slowly, it remains for a prolonged time period after the neo tissue isformed and stabilized. It may hinder the new tissue replacement of the scaffoldspace, and may cause complications associated with long-term foreign body reac-tions. Fourth, the scaffold must have the needed mechanical integrity to maintainthe predesigned tissue structure to serve the 3-D guidance. However, scaffoldsare often not as mechanically strong as the tissues to be replaced because of therequired high porosity for scaffolds. They usually serve the scaffolding purposewell as long as they can maintain the structural integrity under the cultivationor implantation conditions, while the goals are that the engineered tissues fromthe scaffolds are mechanically and biologically functional as their natural coun-terparts. Fifth, the scaffold should not be toxic to the cells (biocompatible). Inaddition to the polymer, the degradation products of the polymer should not betoxic to the cells, which is usually a more restricting requirement than for thepolymer. Sixth, ideally the scaffold should positively interact with cells, includingenhanced cell adhesion, growth, migration, and differentiated function. To achievethese positive cell–scaffold interactions, surface or bulk modifications of the


polymer structure are often employed (18–21). Even drug, protein, or gene de-livery techniques are often considered in the scaffold design (22–25).

Polymers in Tissue Engineering

As discussed above, polymers play a pivotal role in tissue engineering. To fulfillthe diverse needs in tissue engineering, various polymers have been exploited intissue engineering research, including natural polymers (macromolecules), natu-ral polymer-derived materials, synthetic polymers, and synthetic polymers madeof natural monomers or modified with natural moieties. Various copolymers, poly-mer blends, or polymeric composite materials are also used. This section is notintended to be a complete and exhaustive review of all the polymers used in tissueengineering. Instead, some of the most frequently used polymers (macromolecules)in tissue engineering are briefly reviewed.

Polymers for Porous Scaffolds. The polymers to be discussed in this cat-egory can form stable porous structures in the solid form to serve as pre-designed3-D scaffolds. They generally do not dissolve or melt under in vitro tissue cultureconditions (in an aqueous tissue culture medium) or when implanted in vivo.

Linear Aliphatic Polyesters. Linear aliphatic polyesters are the most fre-quently used synthetic biodegradable polymers in tissue engineering and manyother biomedical applications (26–28). These polymers degrade through hydroly-sis of the ester bonds in the polymer backbone. The degradation rates and profilesdiffer between these polymers owing to their compositional, structural, and molec-ular weight differences.

Polyglycolide, also called poly(glycolic acid) (PGA), polylactide, also calledpoly(lactic acid) (PLA), and their copolymers, poly(lactide-co-glycolide), also calledpoly(lactic acid-co-glycolic acid) (PLGA), are a family of linear aliphatic polyesterscalled poly(α-hydroxy acids) or poly(α-hydroxy esters) (Fig. 2). These polymers canbe synthesized by direct condensation of the hydroxy acid monomers (resulting inlow molecular weight polymers, such as lower than 10,000) or more commonly bya ring-opening polymerization of the cyclic dimers (to achieve a higher molecularweight), from where the names of polyglycolide and polylactide stem.

Among the family of glycolic acid and lactic acid homopolymers and copoly-mers, PGA is the simplest in chemical structure, has many advantageous prop-erties, and is therefore one of the most widely used scaffolding polymers (17).Because of the chain structural regularity, PGA is highly crystalline and has ahigh melting point of around 220◦C (17). It does not dissolve in most common or-ganic solvents. Because of its hydrophilic nature, PGA degrades rapidly in aqueoussolutions or in vivo, and loses mechanical integrity between 2 and 4 weeks depend-ing on the molecular weight and physical structure of the scaffolds or implants,and in vitro or in vivo conditions (17,29). It was the material used to develop thefirst synthetic absorbable suture, and has been processed into nonwoven fibrousfabrics as one of the most widely used scaffolds in tissue engineering today.

PLA is also widely used for scaffold fabrication because of its biodegrad-ability. Because of the extra methyl group in PLA repeating unit in comparisonto PGA, PLA is more hydrophobic. The hydrophobic methyl group reducesthe molecular affinity to water and leads to a slower hydrolysis rate of PLA.

Fig. 2. Polymers frequently used as scaffolds for tissue engineering.

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It takes months to years for a PLA scaffold or implant to lose mechanical in-tegrity in vitro or in vivo (30,31). Lactic acid is a chiral molecule and exists intwo stereoisomeric forms, ie, the D-lactic acid and L-lactic acid. The two stereoiso-mers lead to four different types of PLA: poly(D-lactic acid) (PDLA), poly(L-lacticacid) (PLLA), racemic poly(D,L-lactic acid) (PDLLA) and meso poly(D,L-lactic acid)(meso-PLA). Because of the stereoregularity, PDLA and PLLA are semicrystalline,and are less susceptible to water attack (hydrolysis), resulting in slower degra-dation rates. The racemic copolymer PDLLA is amorphous, and therefore moresusceptible to hydrolysis, resulting in a faster degradation rate. Since L-lactic acidis the natural stereoisomer of the lactic acid in the body, PLLA is the frequentlyused PLA. To achieve intermediate degradation rates between PGA and PLLA,various lactic acid and glycolic acid ratios are used to synthesize PLGAs. However,there is not a linear relationship between degradation rate and lactic acid con-tent. When lactide is copolymerized with glycolide, two distinctly different effectswith regard to degradation are introduced. The first is that the lactic acid unit ismore hydrophobic, contributing to a slower degradation rate. The second effect isthat stereoregularity is disrupted by the different repeating units, resulting in alower crystallinity or amorphous structure, contributing to a faster degradationrate. The resulting degradation rate is a compromise of these two factors. Theo-retically, when the LA/GA ratio is higher than 50:50, these two factors are in thesame direction when the amount of lactide is increased. When the LA/GA ratiois lower than 50:50, these two factors are against each other when the amountof lactide is increased. However, in a large middle range of the composition, thecopolymer is amorphous and the crystallinity does not play a role. In both highand low ends of the LA/GA ratio, the crystallinity of the specific scaffold or deviceplays an important role on the degradation rate. In addition, high crystallinityincreases modulus, yield strength, and ultimate strength of the polymers.

In addition to the biodegradability and biocompatibility, these polymers(PLA, PGA, and PLGAs) are among the few synthetic polymers approved by theFood and Drug Administration (FDA) for certain human clinical applications suchas surgical sutures and some implantable devices. There is, however, some con-troversy around the use of these polymers for orthopedic applications. Based on astudy of more than 2500 patients operated on using pins, rods, bolts, and screwsmade of PGA or PLA, 4.3% were affected by a clinically significant local inflam-matory tissue reaction (32). Similarly, other biodegradable polyesters suited formanufacturing absorbable fixation implants appear to also elicit adverse tissueresponses. These adverse tissue reactions are generally considered to result fromthe released acid during degradation. For polymer scaffolds with extremely highporosity used in tissue engineering applications (usually ≥90%) as opposed tosolid implant devices used in orthopedics, the acid release (pH variation) is a lesssevere problem (17,30).

Polymer degradation is often categorized into bulk degradation (bulk ero-sion) and surface degradation (surface erosion) in the fields of biomedical engi-neering and biotechnology (33,34). There are actually differences between erosionand degradation. Degradation is the bond cleavage of the polymer, resulting in re-duction in molecular weight of the polymer. Erosion is the mass loss process. Forideal bulk degradation, hydrolysis (or other forms of degradation) occurs in theentire polymer volume simultaneously. The length of time that polymer persists


can be altered by changing chemical composition but not by the material’s size andshape. For ideal surface degradation, hydrolysis (or other forms of degradation)only occurs on the external surface of the material, ie, at the interface betweenthe material and its immediate surrounding environments, such as medium orbody fluids. Erosion rate is directly proportional to the external surface area. Thelength of time that polymer persists can be altered by varying the sample volume(thickness). Surface erosion is often advantageous for controlled drug deliveryapplications. For example, a thin slab erodes by reducing its thickness and main-taining a nearly constant surface area to achieve constant release rate.

PLA, PGA, and PLGA polymers degrade via bulk hydrolysis. However, thelength of time that a polymer persists is affected by the shape and size of thematerial because the degradation does not occur homogeneously. Common sensewould lead people to think that the surface layer degrades faster since this area isexposed to the most abundant medium or body fluids, which hydrolyze ester bondsin the polymer chains. The reality is just the opposite (35). The cleavage of esterbonds yields carboxyl and hydroxyl end groups. The produced carboxyl end groupsare capable of catalyzing the hydrolysis of remaining ester bonds in the polymerchains. In the surface layer, the chains containing carboxyl end groups or thegenerated small acid molecules diffuse out of the device and the acidity is reduced.This autocatalysis phenomenon is enhanced by the diffusion limitations in thethick devices. In the interior, the acidity increases as ester bonds are hydrolyzedwhile the acidic by-products are trapped. The increased acidity further catalyzeshydrolysis, leading to heterogeneous degradation, ie, a faster degradation in theinterior than on the surface.

For semicrystalline polymers such as PGA and PLLA, the degradation be-havior is also related to the crystallinity and domain structures of the polymers(17). The amorphous domains are easily attacked by water molecules and degradefirst. Initial mass loss occurs in the amorphous domains and causes an apparentcrystallinity increase (17). As the polymer further degrades, the crystalline do-mains also degrade, which leads to drastic decrease in melting point and mechan-ical properties, and finally leads to the disintegration of the entire material (seeBIODEGRADABLE POLYMERS, MEDICAL APPLICATIONS).

There are other linear aliphatic polyesters, such as poly(ε-caprolactone)(PCL) and poly(hydroxy butyrate) (PHB) (Fig. 2), which are also used in tissueengineering research and other biomedical applications. PCL has been investi-gated for a variety of biomedical applications. It can degrade by microorganisms,hydrolytic, enzymatic, or intracellular mechanisms under physiological condi-tions (36,37). PCL is a semicrystalline polymer. It has a very low glass-transitiontemperature of around −62◦C, and thus is always in the rubbery state and hashigh material permeability under physiological conditions. For highly porousscaffolds, material permeability however does not contribute significantly to thescaffold permeability. Compared with PLA, PGA, and PLGA, PCL degrades ata significantly slower rate (38), which makes PCL less attractive for generaltissue engineering applications but more attractive for long-term implants andcontrolled release applications.

Poly(hydroxy butyrate) (PHB), also called poly(β-hydroxy butyrate) orpoly(hydroxy butyric acid), is the simplest member of the polyhydroxyalka-noate (PHA) polyesters (see POLY(3-HYDROXYALKANOATES)). PHB is made by


microorganisms via fermentation. It is biocompatible and biodegradable, but ishighly crystalline and therefore very brittle. PHB degrades very slowly (yearsin vivo) because of the high crystallinity and hydrophobicity (39,40). PHB-basedcopolymers are less crystalline and mechanically more flexible, but degrade alsovery slowly because of the hydrophobic nature, and therefore are less popularcompared to PGA, PLA, and PLGA for tissue engineering applications.

Other Important Synthetic Biodegradable Polymers. Poly(phosphoesters)(PPE) are a class of biodegradable polymers synthesized in the Leong group(Fig. 2) (41). PPE contains P(O) O C linkage in the polymer backbone and thepentavalent phosphorus atom allows for attachment of a pendant component.PPE degrades through hydrolysis of phosphoester bond in the PPE backbone. Thevariation in backbone structure and pendant side group allows for variation instructure and properties of these polymers. They have been used for controlleddrug delivery and gene delivery applications (42), and are also explored for cer-tain tissue engineering applications (43) (see CONTROLLED RELEASE TECHNOLOGY;GENE-DELIVERY POLYMERS).

Poly(propylene fumarate) (PPF) (Fig. 2) has been synthesized as a biodegrad-able polymer that degrades through hydrolysis of the ester bonds similar to gly-colide and lactide polymers. PPF is an amorphous unsaturated polymer and hasbeen extensively studied in the Mikos group as either a prefabricated (44) oran injectable biomaterial (45) for bone tissue engineering. Its copolymers withpoly(ethylene glycol) (PEG), poly(propylene fumarate-co-ethylene glycol) poly-mers, can form hydrogels (46).

Polyphosphazenes (qv) contain a long-chain backbone of alternating phos-phorus and nitrogen, with two side groups attached to each phosphorus (Fig. 2)(47). Various synthesis routes are used to prepare these polymers, allowing differ-ent side groups (R) to be attached to the backbone. These polymers can have verydifferent properties, from degradable to nondegradable, by changes in the sidegroups. Polyphosphazenes are finding more and more applications in controlleddrug delivery applications (48,49), and are also finding ways into the tissue engi-neering research (50,51).

Although polyanhydrides (qv) are attempted for use as tissue engineeringscaffold (52), they are designed primarily for controlled drug delivery purposes inthe Langer group (53). These polymers are very hydrophobic, and degrade throughsurface erosion. The surface erosion characteristic allows easy accomplishment ofconstant release rate for sustained drug delivery purpose. Drugs are also well pro-tected when embedded in such polymers owing to the nearly no water penetrationbefore the polymer erodes. Similarly, poly(ortho esters) are primarily designedfor controlled drug delivery applications because of their surface erosion proper-ties (54). Nevertheless, they have been explored for tissue engineering scaffoldingapplications as well (55).

Amino acids are natural nutrients. Polymers derived from amino acids mayhave advantageous biocompatibility, as demonstrated by PLA. However, aminoacid based polymers often do not have good material properties. They are oftendifficult to process into 3-D structure, and possess poor mechanical properties.Tyrosine-derived polymers have been developed in the Kohn group as biodegrad-able polymers, which have shown certain promising properties (56), and have beenused for bone tissue engineering research (57).


Nondegradable polyurethanes (qv) have been widely used for biomedical ap-plications because of their elastic properties (58). The segmented polyurethanes al-low the structural variations to achieve a range of mechanical properties. A majorlimitation of polyurethanes for biomedical applications is the involvement of toxicprecursors (such as toluene diisocyanates) in the synthesis. Recent efforts havebeen on development of biodegradable polyurethanes or urethane-based polymersusing less toxic diisocyanates (59). These polymers have good mechanical proper-ties and have been explored for vascular and other tissue engineering applications(60,61). Considering the general audience in the field of polymer science and engi-neering, synthetic polymers and 3-D scaffold fabrications are the main foci of thisarticle. However, it should be noted that natural polymers, such as proteins andpolysaccharides, have also been used for tissue engineering applications. Collagen(qv) is a major natural extracellular matrix component (62). Collagen proteinspossess triple-helix structure over a large portion of the molecules. Among thevarious collagen proteins the most abundant is type I collagen. Collagen has beenused for various tissue regeneration applications especially for soft tissue repair(63,64). The use of collagen as a scaffolding material, however, remains challeng-ing because of the potential pathogen transmission and immune reactions, poorhandling and mechanical properties, and less controlled biodegradability (enzy-matic) of the natural material from biological sources. On the other hand, collagenas a natural extracellular component has useful biological properties desired fortissue engineering applications. Yannas and colleagues have developed collagen-glycosaminoglycan (GAG) copolymers and fabricated them into scaffolds for tis-sue engineering (65,66). Denatured collagen (gelatin) is also processed into porousmaterials for tissue repair (67). Polysaccharides (qv) are another class of naturalpolymers, eg, alginate (68) and chitosan (69), which have been explored as tissueengineering scaffolds.

Polymers for Hydrogel Scaffolds. Hydrogels (qv) are cross-linked hy-drophilic polymers that contain large amounts of water without dissolution. Hy-drogels are attractive candidates for certain tissue engineering applications be-cause of the ability to fill irregularly shaped tissue defects, allowance of minimallyinvasive procedures such as arthroscopic surgeries, and the ease of incorporationof cells or bioactive agents (70–75).

Among the synthetic hydrogels, PEG (Fig. 2) has been more frequently stud-ied for tissue engineering research than other synthetic hydrogels. PEG has beenextensively used in biomedical applications often to prevent protein and cell adhe-sion (76). This property also made PEG useful for prevention of tissue adhesionsafter surgery (77,78). In the tissue engineering field, PEG has been used as a modelsystem for studies related to adhesion peptides. In such a case, the inertness ofthis polymer to cells and proteins is ideally utilized. PEG–diacrylamide was syn-thesized by the Hubbell group (79). A photopolymerization step can be performedin contact with cells, providing a means to produce scaffolds for tissue engineering.A PEG-based interpenetration network has been used for cartilage tissue engi-neering by Elisseeff and colleagues (80). Peptide-modified PEG materials havebeen used by Griffith and West groups to study relationships between cell behav-ior (such as adhesion, spreading, and migration) and peptide density and spacing(81–83). A major limitation of PEG hydrogel for tissue engineering scaffolds is itslack of degradability. Efforts are made to impart degradability to PEG by either


formulating copolymers with PLA or PGA, or by introducing enzyme-degradablelinkages into the PEG backbone (84–86).

Alginate is the salt form of alginic acid. Alginic acid, a polysaccharide fromseaweed, is a family of natural copolymers of β-D-mannuronic acid (M) and α-L-guluronic acid (G) (Fig. 2) (87,88). They have been processed into gel beadsencapsulating living cells as a means of immunoprotection (89). Alginate hy-drogels cross-linked using calcium sulfate (CaSO4) have recently been used ascell delivery vehicles for in vivo tissue engineering research (74,90,91). Peptide-modified alginate has been studied to improve cell attachment (74). However,one of the major disadvantages of the use of CaSO4 is that gelation kinetics isdifficult to control, and the resulting structure is not uniform. Structural unifor-mity in tissue engineering scaffolds is necessary not only for uniform cell dis-tribution, but also for well-controlled material properties. Mechanical propertiesare more consistent throughout the gel and between batches if structural unifor-mity can be achieved. We have developed methods to control the gelation rate sothat cross-linking is allowed to take place uniformly throughout the gel to forma structurally uniform and mechanically strong alginate gel with predesigned3-D shapes (Fig. 3a) (70). Cells can be uniformly incorporated into such gels(Fig. 3b).

Collagen, in addition to porous foam, has also been used as hydrogels fora variety of tissue repair and regeneration studies (92,93). Collagen gels havebeen studied as a therapeutic option for the treatment of burn patients or chronicwounds (94). Cartilage defect repair has been studied using chondrocyte–collagengel constructs (95). Studies have also been conducted with collagen gels to delivermesenchymal stem cells and biological agents to improve healing of ligaments andtendons (96).

Polymer Processing and 3-D Scaffolds for Tissue Engineering

In the body, tissues are organized into 3-D structures as functional organs andorgan systems. Each tissue or organ has its own characteristic architecture de-pending on its physiological function. This architecture is also believed to providethe necessary channels for mass transport and spatial cell organization (15). Masstransport includes signaling molecules, nutritional supplies, and metabolic wasteremoval. Spatial cellular organization allows for cell–cell and cell–extracellularmatrix interactions to occur, which are critical to physiological functions of a tis-sue or organ. To successfully engineer functional tissues and organs, the scaffoldshave to be designed to facilitate the desired cell distribution and to guide tissueregeneration in 3-D. This section is, therefore, devoted to the 3-D pore structureand pore wall architecture of scaffolds for tissue engineering.

Textile Technologies and Traditional Fibrous Scaffolds. The textiletechnologies have developed over thousands of years to produce various fibrousfabrics for clothing, filtration, packaging, and many other industrial and householdapplications. Fibrous fabrics have excellent mechanical properties. It was natu-ral that fibrous materials found many biomedical applications in recent yearsincluding sutures from biodegradable polymers. It was also not surprising thatearlier tissue engineering scaffolds were fibrous fabrics of biodegradable polymers


Fig. 3. Alginate hydrogels used as scaffolds for tissue engineering: (a) alginate gels ofvarious molded shapes made from a slow gelation process; (b) MC3T3-E1 osteoblasts in-corporated into slow-gelling alginate gels and cultured in vitro for 3 weeks, demonstratinguniform cell distribution (H&E staining, original magnification: 100×). Reprinted fromRef. 70. c© 2001, by permission of Elsevier.

fabricated using textile technologies. PGA, PLA, and many other semicrystallinepolymers can be processed into fibers using an extruder and these fibers can be fur-ther processed into woven, knit, or nonwoven fabrics using the textile technologies(see NONWOVENS, STAPLE FIBER).

One of such scaffolds widely used in tissue engineering research is PGA non-woven scaffolds (Fig. 4). PGA is typically melt-spun into fibers with a diameter ofaround 15 µm. These fibers are then processed into nonwoven fabrics using textiletechnologies such as carding, needling, heat pressing, and so forth. Porosity higherthan 90% can be easily achieved. These PGA nonwoven scaffolds degrade via hy-drolysis both in vitro and in vivo. The mass of PGA scaffolds decrease exponentiallyin vitro, fitting well to a first-order degradation kinetics (17). The amorphous re-gions in the fibers degrade first presumably because of the easy access by water


Fig. 4. SEM micrograph of a PGA nonwoven scaffold with a porosity of approximately95% and a fiber diameter of approximately 15 µm.

molecules. The crystalline regions start to degrade soon after, leading to loss ofmolecular orientation in the fibers and decrease in mechanical properties. ThesePGA scaffolds have been used either alone or combined with other biodegrad-able polymers for the engineering of cartilage (97–99), tendon (100), ureter (101),intestine (102), blood vessels (103,104), heart valves (105), and other tissues.

Although PGA nonwoven scaffolds have stimulated a large wave of tissue en-gineering research and generated considerable excitement in the field, there areseveral limitations of PGA nonwoven scaffolds, such as low mechanical strength,fast degradation rate (losing mechanical properties within 2 weeks), difficulty inpore shape control, and limited fiber diameter variations. Therefore, many otherprocessing techniques have been developed to fabricate scaffolds for desired prop-erties in various tissue engineering applications.

Particulate-Leaching Techniques. Particulate-leaching is anothertechnique that has been widely used to fabricate porous materials as scaffoldsfor tissue engineering applications. This technique was first developed by Mikosand colleagues (16), and was improved and standardized for large batch fabrica-tions (7). This process is schematically illustrated in Figure 5, which is often calledsalt-leaching technique because NaCl crystals are most often used as the particlesfor pore generation (porogen). Briefly, salt is first ground into small particles, andthe particles are then separated into different size ranges using standard sieves.The particles of desired size are added into a mold of a needed shape. A polymersolution is prepared and cast into the salt-filled mold. After the evaporation ofsolvent, the salt crystals are leached away using water to form the pores of thepolymer foam (Fig. 6).

There is no complicated equipment needed for this technique. The processis easy to carry out and the materials used (water and salt) are economical. Thetechnique is also versatile and can be used to fabricate foams from a variety ofpolymers as long as the polymer can be dissolved in a solvent that does not dissolve


Fig. 5. Schematic illustration of the salt-leaching technique for porous foam fabrication.

the salt. The pore size can be controlled by the size of the salt crystals, and theporosity can be controlled by the salt/polymer ratio. For example, larger particlesize results in larger pore size, and higher salt/polymer ratio results in higherporosity. Therefore, this technique is widely used in the tissue engineering field.However, certain critical variables such as pore shape and interpore openingsare not controlled. For example, salt crystals have the characteristic cubic shape(Fig. 6). A spherical pore shape is not achievable using salt crystals. The resultingpores are often not well-connected because the polymer solution tends to pene-trate between salt particles. To overcome the shortcomings of the salt-leaching

Fig. 6. SEM micrograph of a PLLA foam fabricated using the salt-leaching technique.


technique as well as the textile technologies, various other techniques have beendeveloped to have better control over the pore structure of the scaffolds.

Phase Separation and Novel Architectures of Scaffolds. For a ho-mogeneous multicomponent system, under certain conditions the system becomesthermodynamically unstable and tends to separate into a multiphase system inorder to lower the system free energy. Controlled phase separation processes aredesired for the generation of porous structures for tissue engineering scaffold-ing fabrication. For a polymer solution, phase separation can be induced in sev-eral different ways such as non-solvent-induced phase separation, chemically in-duced phase separation, and thermally induced phase separation. When phaseseparation occurs, a polymer solution separates into two phases, a polymer-richphase (with a high polymer concentration) and a polymer-lean phase (with alow polymer concentration). After the solvent is removed by extraction, evap-oration, or sublimation, the polymer-rich phase solidifies. Depending upon thesystem and phase separation conditions the resulting materials are different inphysical form: powder, closed-pore foam, or open-pore foam (Fig. 7). Porous ma-terials formed through phase separation have been utilized as membranes forfiltration and separation (106). However, the pores formed through phase sepa-ration usually have diameters on the order of a few to tens of micrometers andare often not uniformly distributed, which are not suitable for tissue engineeringapplications.

Fig. 7. Schematic illustration of phase separation processes that lead to different materialforms: A, powder; B, open porous structure suitable as a scaffold for tissue engineering;and C, foam with closed pores.


As discussed earlier, a scaffold must have a pore size big enough for accom-modating cells, allowing proliferation, and having room for new tissue synthesis.They also need to have high surface area to enhance cell adhesion. The pore struc-tures should also allow for vascularization and innervation, and should facilitatediffusion of nutrients and waste materials. Thermally induced phase separationtechniques have been explored in our laboratory for fabricating scaffolds to satisfythe requirements of tissue engineering. This type of phase separation is based onthe thermodynamic and kinetic behavior of polymer solutions, which is a compli-cated process (26,107,108). The important variables are the polymer, solvent, con-centration of the polymer solution, and the phase separation temperature. Thesevariables have been manipulated to generate a variety of porous architectures forscaffold fabrication as briefly discussed below.

Solid–Liquid Phase Separation. Phase separation can be achieved by in-ducing solvent crystallization by decreasing the temperature of a polymer solution.This process is defined as a solid–liquid phase separation (solid-phase formationin a liquid phase). To achieve this, the crystallization temperature (freezing point)of the solvent in the polymer solution needs to be higher than the liquid–liquidphase separation temperature, or the liquid–liquid phase separation is kineti-cally too slow to take place before the temperature reaches the solid–liquid phaseseparation. When the temperature of the solution decreases and reaches the crys-tallization temperature, the solvent crystallizes and the polymer is expelled fromthe solvent crystallization front (separated from the solvent crystal lattice). Afterthe removal of the solvent crystals (sublimation or solvent exchange), the spaceoriginally taken by the solvent crystals becomes pores. By manipulating the phaseseparation variables, foams with a variety of pore morphologies can be obtained.For example, PLLA and PLGA solutions have been used to fabricate intercon-nected pore structures as scaffolds using solid–liquid phase separation techniques(Fig. 8) (5,109). This technique can be used to fabricate scaffolds of more than onetype of materials, including composite scaffold fabrication (Fig. 8c) (109). Theseinterconnected pore structures also allow further modification of the pore wallsurfaces after the scaffold fabrication. Bone-like apatite coating on such polymerscaffolds via a biomimetic process is such an example (20). Proteins and otherbioactive molecules can also be used for such internal pore surface modifications(110).

By manipulating the phase separation conditions, various pore structurescan be achieved. For example, many tissues (such as nerve, muscle, tendon,ligament, dentin, and so on) have oriented tubular or fibrous bundle architectures.To facilitate the organization and regeneration of such tissue types, a scaffoldwith a high porosity and an oriented array of open microtubules may be desirable.To achieve this goal, a novel phase separation technique has been developed forthe creation of a parallel array of microtubules (Fig. 9) (111). In such a process,nucleation of the solvent is induced at one side of the polymer solution. The nucleiwill grow in all directions until they contact with each other on this side. Thenthey cannot grow further in this plane, and are forced to grow along the directionperpendicular to this plane, leading to the formation of an array of parallel rod-like solvent crystals. The polymers are expelled from the solution and squeezedinto thin walls surrounding these parallel solvent rods. After the removal of theserods, a parallel array of microtubules is formed as demonstrated. This oriented

Fig. 8. SEM micrographs of polymer scaffolds fabricated using solid–liquid phase sep-aration: (a) PLLA scaffold fabricated from 5% PLLA/dioxane solution (with local regularpore structure), (b) PLLA scaffold fabricated from 2.5% PLLA/dioxane solution (with lessregular structure), (c) PLLA/HAP (hydroxyapatite) composite scaffold (PLLA/HAP: 50/50)fabricated from a 2.5% PLLA/dioxane solution. Reprinted from Refs. 5 and 109. c© 2001,by permission of John Wiley & Sons.

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Fig. 9. SEM micrographs of a PLLA scaffold with oriented microtubular architectureprepared from 5% PLLA solution: (a) longitudinal section, and (b) cross section. From Maand Reprinted from Ref. 111. c© 2001, by permission of John Wiley & Sons.

tubular scaffold has anisotropic mechanical properties similar to fibrillar andtubular tissues, and has been shown to facilitate cell organization into orientedfibrillar or tubular tissues (Fig. 10) (111).

Liquid–Liquid Phase Separation. In contrast to solid–liquid phase separa-tion, lowering temperature can induce liquid–liquid phase separation of a polymersolution with an upper critical solution temperature and when the crystalliza-tion temperature of the solvent is sufficiently lower than the phase separationtemperature. In an equilibrium phase diagram of a polymer solution, the spin-odal curve divides the liquid–liquid phase separation region into two regions: athermodynamically metastable region (between the binodal and spinodal) and athermodynamically unstable region (enclosed by the spinodal) (Fig. 11). Above the


Fig. 10. MC3T3-E1 cells cultured on a PLLA scaffold with parallel open microtubu-lar architecture for 4 weeks in vitro, von Kossa’s silver nitrate staining. Reprinted fromRef. 111. c© 2001, by permission of John Wiley & Sons.

binodal curve, the solution is homogeneous. When the temperature of the polymersolution is lowered to a point below the binodal curve, the solution is not thermo-dynamically stable and tends to separate into two phases, a polymer-rich phaseand a polymer-lean phase, to lower the system free energy. In a solution of verylow polymer concentration, when the temperature is lowered to a point in themetastable region (between binodal and spinodal), the phase separation occursvia a nucleation and growth mechanism. This process can lead to the formationof small polymer-rich domains (droplets) in a polymer-lean matrix. After the re-moval of the solvent, polymer powder is formed (Fig. 7a). When the temperatureis lowered into an unstable region (spinodal region), the phase separation occursvia a spinodal decomposition mechanism. This process leads to the formation of a

Fig. 11. Schematic equilibrium phase diagram for a polymer solution with an upper crit-ical solution temperature.


bicontinuous structure, in which the polymer-rich and polymer-lean phases areboth continuous phases. After the removal of the solvent, foam with open-porestructure is formed (Fig. 7b). In a solution of very high polymer concentration,when the temperature is lowered to a point in the metastable region, the phaseseparation leads to the formation of small solvent (polymer-lean) domains in apolymer-rich matrix. After the removal of the solvent, polymer foam of closed-porestructure is formed (Fig. 7c). If the polymer is crystalline, the phase separationprocess is more complicated (26). There is a competition between the liquid–liquidphase separation and polymer crystallization under certain conditions, makingthe final pore structure more complex (26,112).

Thermally induced liquid–liquid phase separation can be utilized to fabricatescaffolds for tissue engineering. For example, a solvent can be selected, where thecrystallization temperature is sufficiently lower than the liquid–liquid phase sep-aration temperature of an amorphous polymer solution. A liquid–liquid phaseseparation can be induced by lowering the temperature into the unstable regionon the phase diagram but above the solvent crystallization temperature. For thePLA and PLGA family, a mixture of dioxane and water has been used for liquid–liquid phase separation to fabricate polymer scaffolds with interconnected porestructure (Fig. 12) (112,113).

Nanofibrous Matrix. Collagen (qv) is a major natural extracellular matrixcomponent and possesses a fibrous structure with fiber bundles varying in di-ameter from 50 to 500 nm (62,114). To mimic the nanofibrous architecture ofcollagen and to overcome the concerns over materials from a natural source suchas pathogen transmission and immune rejection, a novel phase separation tech-nique has been developed in our laboratory to fabricate nanofibrous matrices fromsynthetic biodegradable polymers (112). For example, PLLA solutions are cooledto induce phase separation and gelation. The solvent is directly sublimated or firstexchanged with a different solvent and then sublimated. Several solvents and sol-vent mixtures have been utilized to fabricate the desired nanofibrous matrices inour laboratory (Fig. 13) (112).

Three-Dimensional Pore Architecture Design. As discussed abovevarious novel scaffolds have been developed for tissue engineering to address thecritical shortage of donor tissues/organs. These scaffolds have facilitated the tissueengineering research in demonstrating the feasibility and tremendous potentialof tissue engineering. However, these scaffolds are not perfect. One of the com-mon shortcomings of these fabrication technologies is the lack of precise control of3-D pore architecture of the scaffolds. To tackle this problem, computer-assisted-design and computer-assisted-manufacture (CAD and CAM) techniques that havebeen widely used in modern manufacture industry are finding ways into the fieldof tissue engineering (115).

These techniques were early explored by a group of materials scientists andchemical engineers at MIT (116,117). They used one of the solid free-form fabri-cation (a.k.a. rapid prototyping) techniques, called 3-D printing (3DP). With sucha technique, complex-shaped objects can be designed using CAD softwares anddirectly fabricated from the generated CAD models. For example, they fabricatedvarious structures from biodegradable polymers by ink-jet printing a binder ontosequentially laid polymer powder layers. The advantages of such a technique in-clude the precise control of geometry and the feasibility for repeated fabrication


Fig. 12. SEM micrographs of a porous scaffold prepared from a 10% solution of PLGA(85/15) in a mixture of dioxane/H2O (80/20) at two different magnifications: (a) low mag-nification and (b) high magnification. Reprinted from Ref. 112. c© 1999, by permission ofJohn Wiley & Sons.

of the same structure (118). However, the smallness of the powder particles andthe binder drops (pixels) are limited (a few hundred micrometers). The accuracyof positioning the printing nozzle is also limited. The size of a feature (shape of apore or architectural component) is dependent on the resolution of the pixel sizeand positioning control. Therefore the preciseness of the technology is seriouslylimited.

The anticipation of the future improvement of the resolution of such tech-nologies has, nevertheless, maintained the research momentum in this direction.Similarly, another rapid prototyping technique, fused deposition modeling,has been used to fabricate 3-D scaffolds (119). This technique is suitable forprocessing thermoplastic polymers. A heated nozzle is utilized to extrude polymer


Fig. 13. SEM micrographs of a PLLA nanofibrous matrix prepared from 2.5% (w/v)PLLA/THF solution at a phase separation temperature of 8◦C: (a) 500×; (b) 20,000×.Reprinted from Ref. 112. c© 1999, by permission of John Wiley & Sons.

filament on to a platform. A layer is patterned by depositing the continuouspolymer filament and raster in the x–y directions. The motion along z directionallows layer-by-layer integration of the 2-D patterns into 3-D structures. Inour laboratory, a similar process is used to fabricate the negative replica of thescaffold. A polymer solution is cast into such a mold and solidified after theremoval of the solvent. The mold is then dissolved away to form the polymerscaffold with the designed 3-D pore network (Fig. 14).

Lithography is another processing technique that is actively explored for 3-Dscaffold fabrication. Stereolithographic models derived from X-ray computed to-mography and CAD software were used to recreate complex anatomic structure ofa human pulmonary and aortic graft. These stereolithographic models were usedto generate biodegradable heart valve scaffolds by a thermal processing technique(120). Hydroxyapatite scaffolds with various pore shapes have also been fabricated


Fig. 14. Scaffold fabrication using rapid prototyped negative replica: (a) A computer-generated 3-D negative replica of a scaffold with a cubical pore shape; (b) the negativereplica of the scaffold fabricated using a rapid prototyping machine; (c) the polymer scaffoldfabricated using the negative replica; (d) SEM micrograph of the internal pore structureof the generated scaffold (V. J. Chen and P. X. Ma, unpublished data, 2003).

by casting ceramic slurry into molds generated using stereolithography and a sub-sequent sintering process (Fig. 15) (121). Lithography techniques have also beenused to fabricate devices to localize cell populations in patterned configurationson rigid substrates as a potential artificial liver (122). A complex silicon struc-ture of branched vascular channels as a model for liver fabrication has also beendeveloped using stereolithography (123).

However, in addition to the special equipment requirements all these fab-rication techniques have their inherent shortcomings such as limited materialselections and inadequate resolution like the 3DP techniques discussed earlier.Furthermore, the resulting constructs have structural heterogeneity due to the“pixel assembly” nature of these fabrication processes.

In our laboratory, reversed fabrication processes have been developed toovercome these shortcomings for tissue engineering scaffold fabrications (Fig. 16)


Fig. 15. Hydroxyapatite scaffolds fabricated by casting ceramic slurry into molds gener-ated using stereolithography and a subsequent sintering process. Reprinted from Ref. 121.c© 2001, by permission of Kluwer Academic Publishers.

(124). To fabricate 3-D biodegradable polymer scaffolds with well-controlledinterconnected spherical pores, paraffin spheres are fabricated with a dispersionmethod. These paraffin spheres are then transferred into a 3-D mold of a designedshape or anatomical shape of a body part. The spheres are bonded together in themold through a heat treatment process. A polymer solution is cast into the paraffinassembly in the mold (Note: the solvent used should not be a solvent of the paraffinspheres). After removal of the solvent through evaporation or other means, thepolymer–paraffin sphere assembly is immersed in a solvent of the paraffin but nota solvent for the polymer (such as hexane for PLA or PLGA scaffolds) to dissolvethe paraffin sphere assembly. In this way, an interconnected spherical pore struc-ture is created in a predetermined shape by the mold (Fig. 17). Importantly, thegenerated scaffolds have homogeneous foam skeleton (platelet-like, continuous,

Fig. 16. Schematic fabrication processes for polymer scaffolds with interconnected spher-ical pores.


Fig. 17. SEM micrographs of polymer scaffolds [(a)–(d) PLLA; (e) and (f) PLGA85/15]with an interconnected spherical pore structure prepared using paraffin spheres (porogen):(a)100×, paraffin spheres: 250–420 µm; (b) 250×, paraffin spheres: 250–420 µm; (c)1000×,paraffin spheres: 250–420 µm; (d) 3000×, paraffin spheres: 250–420 µm; (e) 50×, paraffinspheres: 420–500 µm; (f) 100×, paraffin spheres: 420–500 µm. Reprinted from Ref. 124. c©2001, by permission of Mary Ann Liebert.

or other complex features, depending on the polymer and phase separation con-ditions) with homogeneous material properties, which are not easily achievablewith free-forming, 3DP or lithography because of the limitation of pixel-by-pixelconstruction. Equally importantly, the spherical pore size is determined by the

Fig. 18. PLLA nanofibrous scaffold with helicoidal tubular macropore network preparedfrom PLLA/THF solution and a helicoidal sugar fiber of the scaffold assembly: (a) schematicillustration of helicoidal sugar fiber assembly; (b) SEM micrograph of the scaffold at anoriginal magnification of 35×, and (c) original magnification of 250×. Reprinted fromRef. 15. c© 2000, by permission of John Wiley & Sons.

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paraffin sphere size used, which can be controlled through paraffin sphere prepa-ration. The features generated are significatly better than those achievable withthe resolutions of the current 3-D fabrication machines. Furthermore, the inter-pore openings are circular and the size of these openings is adjustable throughmanipulating the heat treatment conditions. For example, longer heat treatmenttime leads to larger openings between the pores (124). Finally there is no expensiveequipment investment compared to the textile technologies or rapid prototypingtechniques, which allows this technology to be easily adapted in a research as wellas an industrial setting.

To improve the 3-D structure of nanofibrous scaffolds for cell seeding anddistribution, mass transport, vascular invasion, and tissue organization, tech-niques have also been developed in our laboratory to build predesigned macroporenetworks in nanofibrous matrices (15). For example, larger water-soluble fibers(diameters from 100 µm to 1 mm) are prepared from sugar as a geometrical poro-gen element, and are assembled into various 3-D structures (such as helicoidal)(Fig. 18a). PLLA solution is then cast into this 3-D assembly and is thermallyinduced to phase-separate for nanofibrous matrix formation. After the solvent re-moval, the sugar fiber assembly is dissolved away using water to achieve nanofi-brous scaffolds with predesigned helicoidal tubular pore network (Figs. 18b and18c). Similarly, we have combined the interconnected spherical pore network de-scribed earlier with nanofiber techniques to generate nanofibrous scaffolds withinterconnected spherical macropores (Fig. 19) (125).

As discussed above, tissue engineering scaffold fabrication is a fast evolvingarea. Many other polymer processing techniques such as gas-foaming, emulsionfreeze-drying, and so forth have also been explored (126–128). Current efforts forthese techniques include overcoming the disadvantages of closed-pore structuresand undesired pore size ranges.

Again this article is intended to introduce the general concepts of tissueengineering in relation to polymer science and engineering, but is not intended tobe a complete and exhaustive review of the field of tissue engineering. There are

Fig. 19. SEM micrographs of nanofibrous scaffolds with interconnected spherical porenetwork: (a) original magnification 30×, (b) original magnification 4000×. Reprinted fromRef. 125. c© 2003, by permission of John Wiley & Sons.


recently developed comprehensive books (129–131) available for further readingon various topics in tissue engineering.

BIBLIOGRAPHY

1. J. P. Vacanti and R. Langer, Lancet 354 (Suppl 1), SI32–SI34 (1999).2. N. Couch, R. Wilson, E. Hager, and J. Murray, Surgery 59(2), 183–188 (1966).3. R. Langer and J. P. Vacanti, Science 260, 920–926 (1993).4. V. M. Goldberg and S. Stevenson, Orthopedics 17, 809–810, 821 (1994).5. P. X. Ma, R. Zhang, G. Xiao, and R. Franceschi, J. Biomed. Mater. Res. 54, 284–293

(2001).6. R. M. Nerem and A. Sambanis, Tissue Eng. 1(1), 3–13 (1995).7. P. X. Ma and R. Langer, in J. Morgan and M. Yarmush, eds., Tissue Engineering

Methods and Protocols, Humana Press, Inc., Totowa, N.J., 1999, pp. 47–56.8. L. Peterson, M. Brittberg, I. Kiviranta, E. Akerlund, and A. Lindahl, Am. J. Sports

Med. 30(1), 2–12 (2002).9. J. Bloch, E. Fine, N. Bouche, A. Zurn, and P. Aebischer, Exp. Neurol. 172, 425–432

(2001).10. G. N. King, Curr. Pharm. Biotechnol. 2(2), 131–142 (2001).11. C. K. Colton, Nat. Biotechnol. 17, 421–422 (1999).12. E. S. Tzanakakis, D. J. Hess, T. D. Sielaff, and W. S. Hu, Annu. Rev. Biomed. Eng. 2,

607–632 (2000).13. H. D. Humes, W. H. Fissell, W. F. Weitzel, D. A. Buffington, A. J. Westover, S. M.

MacKay, and J. M. Gutierrez,Am. J. Kidney Dis. 39, 1078–1087 (2002).14. S. Del Guerra, C. Bracci, K. Nilsson, A. Belcourt, L. Kessler, R. Lupi, L. Marselli,

P. De Vos, and P. Marchetti, Biotechnol. Bioeng. 75, 741–744 (2001).15. R. Zhang and P. X. Ma, J. Biomed. Mater. Res. 52, 430–438 (2000).16. A. G. Mikos, A. J. Thorsen, L. A. Czerwonka, Y. Bao, R. Langer, D. N. Winslow, and

J. P. Vacanti, Polymer 35, 1068–1077 (1994).17. P. X. Ma and R. Langer, in A. G. Mikos, K. W. Leong, M. L. Radomsky, J. A. Tamada,

and M. J. Yaszemski, eds., Polymers in Medicine and Pharmacy, MRS, Pittsburgh,Pa., 1995 pp. 99–104.

18. J. A. Hubbell, Curr. Opin. Biotechnol. 10(2), 123–129 (1999).19. D. Barrera, E. Zylstra, P. Lansbury, and R. Langer, J. Am. Chem. Soc. 115,

11010–11011 (1993).20. R. Zhang and P. X. Ma, J. Biomed. Mater. Res. 45, 285–293 (1999).21. H. Shi and B. D. Ratner, J. Biomed. Mater. Res. 49(1), 1–11 (2000).22. H. Lee, R. A. Cusick, F. Browne, T. Ho Kim, P. X. Ma, H. Utsunomiya, R. Langer, and

J. P. Vacanti, Transplantation 73, 1589–1593 (2002).23. J. Bonadio, E. Smiley, P. Patil, and S. Goldstein, Nat. Med. 5, 753–759 (1999).24. H. Lo, S. Kadiyala, S. E. Guggino, and K. W. Leong, J. Biomed. Mater. Res. 30,

475–484 (1996).25. M. H. Sheridan, L. D. Shea, M. C. Peters, and D. J. Mooney, J. Controlled Release

64(1–3), 91–102 (2000).26. R. Zhang and P. X. Ma, in A. Atala and R. Lanza, eds., Methods of Tissue Engineering,

Academic Press, San Diego, Calif., 2001, pp. 715–724.27. J. H. Elisseeff and Y. Yamada, in K.-U. Lewandrowski, D. L. Wise, D. J. Trantolo,

J. D. Gresser, M. J. Yaszemski, and D. E. Altobelli, eds., Tissue Engineering andBiodegradable Equivalents: Scientific and Clinical Applications, Marcel Dekker, Inc.,New York, 2002, pp. 1–23.

28. J. M. Pachence and J. Kohn, in R. P. Lanza, R. Langer, and J. Vacanti, eds., Principlesof Tissue Engineering, Academic Press, San Diego, Calif., 2000, pp. 263–277.


29. A. Reed and D. Gilding, Polymer 22, 494–498 (1981).30. R. Zhang and P. X. Ma, Polym. Prepr. 41, 1618–1619 (2000).31. B. Eling, S. Gogolewski, and A. J. Pennings, Polymer 23, 1587–1593 (1982).32. O. M. Bostman and H. K. Pihlajamaki, Clin. Orthop. 371, 216–227 (2000).33. J. A. Tamada and R. Langer, Proc. Nat. Acad. Sci. U.S.A. 90, 552–556 (1993).34. A. Gopferich, Biomaterials 17(2), 103–114 (1996).35. S. Li, J. Biomed. Mater. Res. 48, 342–353 (1999).36. C. G. Pitt, M. M. Gratzl, G. L. Kimmel, J. Surles, and A. Schindler, Biomaterials 2,

215–220 (1981).37. S. C. Woodward, P. S. Brewer, F. Moatamed, A. Schindler, and C. G. Pitt, J. Biomed.

Mater. Res. 19, 437–444 (1985).38. C. G. Pitt, in M. Chasin and R. Langer, eds., Biodegradable Polymers as Drug Delivery

Systems, Marcel Dekker, Inc., New York, 1990, pp. 71–119.39. S. J. Holland, A. M. Jolly, M. Yasin, and B. J. Tighe, Biomaterials 8, 289–295 (1987).40. N. D. Miller and D. F. Williams, Biomaterials 8(2), 129–137 (1987).41. M. Richards, B. I. Dahiyat, D. M. Arm, P. R. Brown, and K. W. Leong, J. Biomed. Mater.

Res. 25, 1151–1167 (1991).42. J. Wang, H. Q. Mao, and K. W. Leong, J. Am. Chem. Soc. 123, 9480–9481 (2001).43. S. Wang, A. C. Wan, X. Xu, S. Gao, H. Q. Mao, K. W. Leong, and H. Yu, Biomaterials.

22, 1157–1169 (2001).44. M. S. Wolfe, D. Dean, J. E. Chen, J. P. Fisher, S. Han, C. M. Rimnac, and A. G. Mikos,

J. Biomed. Mater. Res. 61(1), 159–164 (2002).45. S. J. Peter, S. T. Miller, G. Zhu, A. W. Yasko, and A. G. Mikos, J. Biomed. Mater. Res.

41(1), 1–7 (1998).46. E. Behravesh, S. Jo, K. Zygourakis, and A. G. Mikos, Biomacromolecules 3, 374–381

(2002).47. H. R. Allcock, in M. Chasin and R. Langer, eds., Biodegradable Polymers as Drug

Delivery Systems, Marcel Dekker, Inc., New York, 1990, pp. 163–193.48. P. Caliceti, F. M. Veronese, and S. Lora, Int. J. Pharm. 211(1/2), 57–65 (2000).49. N. N. Aldini, P. Caliceti, S. Lora, M. Fini, G. Giavaresi, M. Rocca, P. Torricelli,

R. Giardino, and F. M. Veronese, J. Orthop. Res. 19, 955–961 (2001).50. A. M. Ambrosio, H. R. Allcock, D. S. Katti, and C. T. Laurencin, Biomaterials 23,

1667–1672 (2002).51. S. Cohen, M. C. Bano, L. G. Cima, H. R. Allcock, J. P. Vacanti, C. A. Vacanti, and

R. Langer, Clin. Mater. 13(1–4), 3–10 (1993).52. A. K. Burkoth, J. Burdick, and K. S. Anseth, J. Biomed. Mater. Res. 51, 352–359 (2000).53. K. W. Leong, J. Kost, E. Mathiowitz, and R. Langer, Biomaterials 7, 364–371 (1986).54. U.S. Pat. 04093709 (1978), N. S. Choi and J. Heller.55. K. P. Andriano, Y. Tabata, Y. Ikada, and J. Heller, J. Biomed. Mater. Res. 48, 602–612

(1999).56. K. James, H. Levene, J. R. Parsons, and J. Kohn, Biomaterials 20, 2203–2212

(1999).57. K. A. Hooper, N. D. Macon, and J. Kohn, J. Biomed. Mater. Res. 41, 443–454 (1998).58. H. B. Lin, W. Sun, D. F. Mosher, C. Garcia-Echeverria, K. Schaufelberger, P. I. Lelkes,

and S. L. Cooper, J. Biomed. Mater. Res. 28, 329–342 (1994).59. G. A. Skarja and K. A. Woodhouse, J. Biomater. Sci., Polym. Ed. 12, 851–873 (2001).60. J. J. Guan, M. S. Sacks, E. J. Beckman, and W. R. Wagner, J. Biomed. Mater. Res. 61,

493–503 (2002).61. J. Y. Zhang, E. J. Beckman, N. P. Piesco, and S. Agarwal, Biomaterials 21, 1247–1258

(2000).62. E. D. Hay, Cell Biology of Extracellular Matrix, 2nd ed., Plenum Press, New York,

1991, pp. 7–44.


63. E. Bell, M. Rosenberg, P. Kemp, R. Gay, G. Green, N. Muthukumaran, and C. Nolte,J. Biomech. Eng. 113(2), 113–119 (1991).

64. J. M. Pachence, J. Biomed. Mater. Res. 33(1), 35–40 (1996).65. I. V. Yannas, J. Cell. Biochem. 56(2), 188–191 (1994).66. S. M. Mueller, S. Shortkroff, T. O. Schneider, H. A. Breinan, I. V. Yannas, and

M. Spector, Biomaterials 20, 701–709 (1999).67. Y. S. Choi, S. R. Hong, Y. M. Lee, K. W. Song, M. H. Park, and Y. S. Nam, J. Biomed.

Mater. Res. 48, 631–639 (1999).68. L. Shapiro and S. Cohen, Biomaterials 18, 583–590 (1997).69. S. V. Madihally and H. W. Matthew, Biomaterials 20, 1133–1142 (1999).70. C. K. Kuo and P. X. Ma, Biomaterials 22, 511–521 (2001).71. J. P. Fisher, T. A. Holland, D. Dean, P. S. Engel, and A. G. Mikos, J. Biomater. Sci.,

Polym. Ed. 12, 673–687 (2001).72. J. H. Elisseeff, A. Lee, H. K. Kleinman, and Y. Yamada, Ann. N.Y. Acad. Sci. 961,

118–122 (2002).73. J. A. Burdick, A. J. Peterson, and K. S. Anseth, Biomaterials 22, 1779–1786 (2001).74. E. Alsberg, K. W. Anderson, A. Albeiruti, R. T. Franceschi, and D. J. Mooney, J. Dent.

Res. 80, 2025–2029 (2001).75. B. K. Mann, A. S. Gobin, A. T. Tsai, R. H. Schmedlen, and J. L. West, Biomaterials 22,

3045–3051 (2001).76. N. A. Alcantar, E. S. Aydil, and J. N. Israelachvili, J. Biomed. Mater. Res. 51, 343–351

(2000).77. N. D. Winblade, I. D. Nikolic, A. S. Hoffman, and J. A. Hubbell, Biomacromolecules 1,

523–533 (2000).78. N. K. Mongia, K. S. Anseth, and N. A. Peppas, J. Biomater. Sci., Polym. Ed. 1055, 7,

1055–1064.79. D. L. Elbert and J. A. Hubbell, Biomacromolecules 2, 430–441 (2001).80. J. Elisseeff, K. Anseth, D. Sims, W. McIntosh, M. Randolph, M. Yaremchuk, and

R. Langer, Plast. Reconstr. Surg. 104, 1014–1022 (1999).81. L. G. Griffith and S. Lopina, Biomaterials 19, 979–986 (1998).82. D. J. Irvine, A. M. Mayes, and L. G. Griffith, Biomacromolecules 2(1), 85–94 (2001).83. B. K. Mann and J. L. West, J. Biomed. Mater. Res. 60(1), 86–93 (2002).84. D. K. Han and J. A. Hubbell, Macromolecules 30, 6077–6083 (1997).85. A. S. Sawhney, C. P. Pathak, and J. A. Hubbell, Macromolecules 26, 581–587 (1993).86. J. L. West and J. A. Hubbell, Macromolecules 32, 241–244 (1999).87. A. Martinsen, G. Skjak-Bræk, and O. Smidsrød, Biotechnol. and Bioeng. 33, 79–89

(1989).88. K. I. Draget, K. Ostgaard, and O. Smidsrod, Carbohydr. Polym. 14, 159–178 (1991).89. P. Aebischer, E. Buchser, J. Joseph, J. Favre, T. N. de, M. Lysaght, S. Rudnick, and

M. Goddard, Transplantation 58, 1275–1277 (1994).90. A. Atala, W. Kim, K. Paige, C. Vacanti, and A. Retik, J. Urol. 152(2, Ptart 2), 641–643

(1994) (discussion 644).91. K. Paige, L. Cima, M. Yaremchuk, B. Schloo, J. Vacanti, and C. Vacanti, Plast. Reconstr.

Surg. 97(1), 168–178 (1996) (discussion 179–180).92. S. Wakitani, T. Goto, S. J. Pineda, R. G. Young, J. M. Mansour, A. I. Caplan, and

V. M. Goldberg, J. Bone Joint Surg. 76, 579–592 (1994).93. D. Ceballos, X. Navarro, N. Dubey, G. Wendelschafer-Crabb, W. R. Kennedy, and

R. T. Tranquillo, Exp. Neurol. 158, 290–300 (1999).94. F. A. Auger, M. Rouabhia, F. Goulet, F. Berthod, V. Moulin, and L. Germain, Med.

Biol. Eng. Comput. 36(6), 801–812 (1998).95. S. Kawamura, S. Wakitani, T. Kimura, A. Maeda, A. I. Caplan, K. Shino, and T. Ochi,

Acta Orthop. Scand. 69(1), 56–62 (1998).


96. K. A. Hildebrand, F. Jia, and S. L. Woo, Microsc. Res. Tech. 58(1), 34–38 (2002).97. C. Vacanti, R. Langer, B. Schloo, and J. Vacanti, Plast. Reconstr. Surg 88, 753–759

(1991).98. P. X. Ma, B. Schloo, D. Mooney, and R. Langer, J. Biomed. Mater. Res. 29, 1587–1595

(1995).99. L. E. Freed, R. Langer, I. Martin, N. R. Pellis, and G. Vunjak-Novakovic, Proc. Natl.

Acad. Sci. U.S.A. 94, 13885–13890 (1997).100. Y. Cao, J. P. Vacanti, X. Ma, K. T. Paige, J. Upton, Z. Chowanski, B. Schloo, R. Langer,

and C. A. Vacanti, Transplant. Proc. 26, 3390–3392 (1994).101. A. Atala, J. Vacanti, C. Peters, J. Mandell, A. Retik, and M. Freeman, J. Urol. 148(2,

Part 2), 658–662 (1992).102. G. Organ, D. Mooney, L. Hansen, B. Schloo, and J. Vacanti, Transplant. Proc. 25(1,

Part 2), 998–1001 (1993).103. T. Shinoka, D. Shum-Tim, P. X. Ma, R. E. Tanel, N. Isogai, R. Langer, J. P. Vacanti,

and J. E. Mayer Jr., J. Thorac. Cardiovasc. Surg. 115, 536–545 (1998) (discussion545–536).

104. L. E. Niklason, J. Gao, W. M. Abbott, K. K. Hirschi, S. Houser, R. Marini, andR. Langer, Science 284, 489–493 (1999).

105. T. Shinoka, P. X. Ma, D. Shum-Tim, C. K. Breuer, R. A. Cusick, G. Zund, R. Langer,J. P. Vacanti, and J. E. Mayer Jr., Circulation 94(9, Suppl.), II164–II168 (1996).

106. R. W. Baker, Membrane Technology and Applications, McGraw-Hill, New York, 2000.107. P. vandeWitte, P. J. Dijkstra, J. W. A. vandenBerg, and J. Feijen, J. Membr. Sci.

117(1/2), 1–31 (1996).108. R. Bansil, G. D. Liao, and P. Falus, Physica A 231, 346–358 (1996).109. R. Zhang and P. X. Ma, J. Biomed. Mater. Res. 44, 446–455 (1999).110. K. M. Woo, R. Zhang, H. Deng, and P. X. Ma, Transaction of the 28th Annual Meeting

of the Society for Biomaterial, 2002, p. 605.111. P. X. Ma and R. Zhang, J. Biomed. Mater. Res. 56, 469–477 (2001).112. P. X. Ma and R. Zhang, J. Biomed. Mater. Res. 46(1), 60–72 (1999).113. Y. S. Nam and T. G. Park, J. Biomed. Mater. Res. 47(1), 8–17 (1999).114. T. Elsdale and J. Bard, J. Cell Biol. 54, 626–637 (1972).115. S. Yang, K. F. Leong, Z. Du, and C. K. Chua, Tissue Eng. 8(1), 1–11 (2002).116. R. A. Giordano, B. M. Wu, S. W. Borland, L. G. Cima, E. M. Sachs, and M. J. Cima,

J. Biomater. Sci., Polym. Ed. 8(1), 63–75 (1996).117. A. Park, B. Wu, and L. G. Griffith, J. Biomater. Sci., Polym. Ed. 9(2), 89–110 (1998).118. J. Zeltinger, J. K. Sherwood, D. A. Graham, R. Mueller, and L. G. Griffith, Tissue Eng.

7, 557–572 (2001).119. D. W. Hutmacher, T. Schantz, I. Zein, K. W. Ng, S. H. Teoh, and K. C. Tan, J. Biomed.

Mater. Res. 55, 203–216 (2001).120. R. Sodian, M. Loebe, A. Hein, D. P. Martin, S. P. Hoerstrup, E. V. Potapov,

H. Hausmann, T. Lueth, and R. Hetzer, ASAIO J. 48(1), 12–16 (2002).121. T.-M. G. Chu, J. W. Halloran, S. J. Hollister, and S. E. Feinberg, J. Mater. Sci. Mater.

Med. 12, 471–478 (2001).122. S. N. Bhatia, U. J. Balis, M. L. Yarmush, and M. Toner, Biotechnol. Prog. 14, 378–387

(1998).123. S. Kaihara, J. Borenstein, R. Koka, S. Lalan, E. R. Ochoa, M. Ravens, H. Pien,

B. Cunningham, and J. P. Vacanti, Tissue Eng. 6(2), 105–117 (2000).124. P. X. Ma and J. Choi, Tissue Eng. 7(1), 23–33 (2001).125. K. M. Woo, V. J. Chen, and P. X. Ma, J. Biomed. Mater. Res. (in press).126. D. J. Mooney, D. F. Baldwin, N. P. Suh, J. P. Vacanti, and R. Langer, Biomaterials 17,

1417–1422 (1996).127. K. Whang, C. H. Thomas, K. E. Healy, and G. Nuber, Polymer 36, 837–842 (1995).

Vol. 12 TRANSPORT PROPERTIES 291

128. C. M. Agrawal and R. B. Ray, J. Biomed. Mater. Res. 55(2), 141–150 (2001).129. R. P. Lanza, R. Langer, and J. Vacanti, Principles of Tissue Engineering, 2nd ed.,

Academic Press, San Diego, Calif., 2000.130. A. Atala and R. P. Lanza, Methods of Tissue Engineering, Academic Press, San Diego,

Calif., 2002.131. J. R. Morgan and M. L. Yarmush, in J. M. Walker, ed., Tissue Engineering Methods

and Protocols (Methods in Molecular Medicine), Vol. 18. Humana Press, Inc., Totowa,N.J., 1999.

PETER X. MA

University of Michigan

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