Abraham-2008-Guide to collagen ch

ReviewGuide to Collagen Characterization for Biomaterial Studies

Leah C. Abraham,1,2 Erin Zuena,1,2 Bernardo Perez-Ramirez,3 David L. Kaplan1,2

1 Departments of Chemical and Biological Engineering, Tufts University, Medford, Massachusetts 02155

2 Department of Biomedical Engineering, Bioengineering and Biotechnology Center, Tufts University,Medford, Massachusetts 02155

3 Genzyme Corporation, BioFormulations Development, Framingham, Massachusetts 01701-9322

Received 3 January 2007; revised 19 November 2007; accepted 18 December 2007Published online 3 April 2008 in Wiley InterScience (www.interscience.wiley.com). DOI: 10.1002/jbm.b.31078

Abstract: The structure and remodeling of collagen in vivo is critical to the pathology and

healing of many human diseases, as well as to normal tissue development and regeneration. In

addition, collagen matrices in the form of fibers, coatings, and films are used extensively

in biomaterial and biomedical applications. The specific properties of these matrices, both in

terms of physical and chemical characteristics, have a direct impact on cellular adhesion,

spreading, and proliferation rates, and ultimately on the rate and extent of new extracellular

matrix formation in vitro or in vivo. In recent studies, it has also been shown that collagen

matrix structure has a major impact on cell and tissue outcomes related to cellular aging and

differentiation potential. Collagen structure is complex because of both diversity of source

materials, chemistry, and structural hierarchy. With such significant impact of collagen

features on biological outcomes, it becomes essential to consider an appropriate set of

analytical tools, or guide, so that collagens attained from commercial vendors are

characterized in a comparative manner as an integral part of studies focused on biological

parameters. The analysis should include as a starting point: (a) structural detail—mainly

focused on molecular mass, purity, helical content, and bulk thermal properties, (b) chemical

features—mainly focused on surface elemental analysis and hydrophobicity, and (c)

morphological features at different length scales. The application of these analytical

techniques to the characterization of collagen biomaterial matrices is critical in order to

appropriately correlate biological responses from different studies with experimental

outcomes in vitro or in vivo. As a case study, the analytical tools employed for collagen

biomaterial studies are reviewed in the context of collagen remodeling by fibroblasts. The goal

is to highlight the necessity of understanding collagen biophysical and chemical features as a

prerequisite to (a) studies with cells and tissue formation, and (b) suggest modes to establish

comparative outcomes for studies conducted in different laboratories. ' 2008 Wiley Periodicals,

Inc. J Biomed Mater Res Part B: Appl Biomater 87B: 264–285, 2008

Keywords: collagen; characterization; denatured; structure; biomaterials

INTRODUCTION

Collagen Biomaterials

The prevalence of collagen in human tissues makes it a

natural choice as a polymer for biomedical materials and

tissue-engineering matrices. Collagen is the most abun-

dant protein present in mammals, composing 30% by

weight of body protein tissue.1,2 Collagen is also biode-

gradable, biocompatible, and enhances cellular penetra-

tion and wound repair.3 The potential value of collagen

as a biomaterial has led to research on use in scaffolds

for ligament repair, collagen grafts for scar and burn

repair, and the engineering of osteochondral tissue.4–7

Collagen is also a target for study in diseases involving

extensive collagen remodeling, including aortic heart

valve repair and bone repair.8,9 A better understanding of

the interactions between cells and collagen should allow

for the more rational design and use of these substrates

depending on the cells, tissues, and environments

involved in vitro and in vivo. For the purpose of this

review, the focus is mainly on commercially available

collagens for use in studies of biomaterial structure and

function. While many other sources (e.g., collagen

derived from various tissues, recombinant DNA derived

Correspondence to: D. L. Kaplan (e-mail: [email protected])

' 2008 Wiley Periodicals, Inc.

264

collagens, collagen-like peptides) and variations of colla-

gen (e.g., crosslinked, with telopeptides, and related var-

iations) are available to researchers who isolate their own

materials, many of the core analytical tools would remain

similar to those described here in the context of commer-

cially available prepared sources.

There are over 20 known types of collagen (Table I).11

The fibril-forming (fibrillar) collagens include collagen type

I [a1(I)]2a2 (I) comprising fibril bone, skin, tendons, liga-

ments, cornea, and internal organs, accounting for 90% of

body collagen; collagen type II [a1(II)]3 comprising fibril

cartilage, intervertebral disc, notochord, and vitreous humor

of the eye; collagen type III [a1(III)]3 comprising fibril

skin, blood vessels, and internal organs; collagen type V

[a1(V)]2a2(V) and a1(V) a2(V) a3(V) fibril (with type I)

comprising tissue similar to those for type I collagen; colla-

gen type XI a1(XI) a2(IX) a3(XI) fibril (with type II) com-

prising tissue similar to collagen type II.11 For all collagen

types, each collagen chain of �1000 amino acids is com-

posed of three left-handed a helix chains that twist together

to form the right-handed helix of the collagen molecule.2,13

The collagen molecule is about 300 kDa, composed of

�10% each of proline and hydroxyproline, and has glycine

present at every third amino acid position.13

A variety of commercial collagen sources are used for

tissue-engineering applications as well as for cell studies

that require collagen matrices. Table II lists some of the

more common commercial collagen sources used in these

types of studies. These materials, often from poorly speci-

fied preparations, make comparisons between studies of

various collagen materials difficult. An additional issue is

that collagen source materials are often prepared differently

in each laboratory, complicating further attempts at com-

parisons of biological outcomes. One of the aims of this

review is to summarize the characteristics of collagen

source materials to draw common themes in terms of how

the source material and the physical features of the source

relate to biological outcomes. To accomplish this goal, we

have focused on the characterization of commercially avail-

able collagen preparations in order to highlight some of the

complications and strategies that can be employed toward a

working ‘‘guide’’ for assessment of these materials.

Vendors, such as Sigma Aldrich, continue to refer to

collagens both by the widely used ‘‘modern’’ type classifi-

cation system defined by the chain types (type I 5(a1[I])2a2[I])1, type II 5 (a1[II])3)

10 and by types as defined

in the early 1970s by the researchers who first separated

collagen chains (Miller type II cartilage).28 The earlier col-

lagen classifications tended to be defined more by the tissue

type from which the collagen had been extracted than the

collagen chain content.

TABLE I. List of Some of the Collagen Types and Information on Chain Composition, Structure, Tissue Location and RelatedInformation2,10–12

Types Chain Composition Structural Details Localization Notes

I [a1(I)]2[a(I)] 300 nm, 67-nm banded fibrils Skin, tendon, bone, etc. 90% of all collagen of the

human body. Scar tissue-

the end product when

tissue heals by repair.

II [a1(II)]3 300 nm, small 67-nm fibrils Cartilage, vitreous humor Articular cartilage

III [a1(III)]3 300 nm, small 67-nm fibrils Skin, muscle, frequently

with type I

Collagen of granulation tissue,

and is produced quickly by

young fibroblasts before the

tougher type I collagen is

synthesized.

IV [a1(IV)2[a2(IV)] 390 nm C-term globular domain,

nonfibrillar

All basal lamina Basal lamina

V [a1(V)][a2(V)][a3(V)] 390 nm N-term globular domain,

small fibers

Most interstitial tissue,

assoc. with type I

Most interstitial tissue, assoc.

with type I

VI [a1(VI)][a2(VI)][a3(VI)] 150 nm, N1C term. globular

domains, microfibrils,

100-nm banded fibrils

Most interstitial tissue,

assoc. with type I

Most interstitial tissue, assoc.

with type I

VII [a1(VII)]3 450 nm, dimer Epithelia Epithelia

VIII [a1(VIII)]3 130 nm, N1C term. Globular

domains

Some endothelial cells Some endothelial cells

IX [a1(IX)][a2(IX)][a3(IX)] 200 nm, N-term. Globular

domain, bound proteoglycan

Cartilage, assoc. with type II Cartilage, assoc. with type II

X [a1(X)]3 150 nm, C-term. Globular

domain

Hypertrophic and mineralizing

cartilage

Hypertrophic and mineralizing

cartilage

XI [a1(XI)][a2(XI)][a3(XI)] 300 nm, small fibers Cartilage Cartilage

XII a1(XII) 75-nm triple helical tail, central

globule, three 60-nm globule

arms

Interacts with types I and III Interacts with types I and III

Mainly types I–IV have been utilized to varying degrees in tissue-engineering related biomaterials studies, with some efforts on the other types shown.

265GUIDE TO COLLAGEN CHARACTERIZATION FOR BIOMATERIAL STUDIES

Journal of Biomedical Materials Research Part B: Applied Biomaterials

TABLE

II.CollagensandAssociatedData

Available

From

Vendors,UnlessOtherw

iseIndicated

Collagen

Material

Source

Preparation

Characteristics

TissueEngineeringApplication

Vitrogen

Collagen

Corp,PaloA

lto,CA

Bovinedermal

collagen

dissolved

in

0.012NHCl

99.9%

pure

collagen

bySDS

polyacrylamidegel

electrophoresis

inconjunctionwithbacterial

collagenasesensitivityandsilver

staining

Collagen

scaffold

fortendonrepair14

BD

matrigelTM

matrix

BD

Biosciences

Extractedfrom

EHSmouse

sarcoma,

atumorrich

in

ECM

proteins

Solubilized

basem

entmem

brane

preparation.Majorcomponentis

laminin

(56%),followed

by

collagen

IV(31%),heparan

sulfate

proteoglycans,andentactin

(8%)

Cellattachmentanddifferentiationin

3T3-F442A

preadipocytes1

5,16

RochetypeIrattail1

179179

Roche

Purified

from

rattailtendonbya

modificationofthemethodof

Bornstein17;Michalopoulosand

Pitot18

Collagen

from

rattail,mainly

oftype

Icollagen

Collagen

remodelingby

fibroblasts1

9,20

RocheCollagen

Sfrom

calf

skin

1098292

Roche

Collagen

ispurified

from

calfskin

by

extractionwith0.5M

acetic

acid,

pH

2.5

Collagen:[98%,collagen

type1:

[95%,collagen

typeIII:\5%

Comparisonto

collagen

extracted

from

tissues

21

SigmabovinetypeIcalfskin

sterilesolution

SigmaAldrich

Prepared

from

calfskin.Further

details

arenotprovided

bythe

vendor22

0.1%

(1mg21mL)solutionofcalf

skin

collagen

in0.1M

acetic

acid

Recommended

foruse

asacell

culture

substratum

at6–10lg

cm22.

Notsuitable

for3D

gel

form

ation

SigmabovinetypeIcalfskin

BioChem

ikasoluble

SigmaAldrich

Prepared

byamodificationofthe

procedure

ofGallopandSeifter

23

Solubilitynotedas

5mg21mLin

water

byvendor,hazy,colorless

andviscous

Synthesis

andcharacterizationofa

model

extracellularmatrixthat

inducespartial

regenerationofadult

mam

malianskin

Sigmabovinenasal

septum

SigmaAldrich

Prepared


pepsinextractionprocedure

of

Niyibiziet

al.24

TypeIIcollagen

(BornsteinandTraub

classification22)

Activityofstachyrase

Aagainst

collagen

25

SigmabovinetypeIAchilles

tendon

SigmaAldrich

Prepared

bythemethodofEinbinder

J

andSchubert26

TypeIcollagen

(BornsteinandTraub

classification22)

Suitable

foruse

asasubstrate

for

collagenase

Sigmachicken

sternal

SigmaAldrich

Prepared


methodofTrentham

D.E.,et

al.27

Thiscollagen

has

beentested

in

culture

withmam

maliancellsto

verifyitislow

inendotoxin

content.

MillerclassificationtypeII28

Recommended

foruse

asacell

culture

substratum

at6–10lg

cm22

Sigmahuman

typeI

Sigma-Aldrich

Prepared

from

human

skin

by

modificationofGallopandSeifter

23

�95%

(SDS-PAGE)typeI,acid-

soluble

Preparationofsoluble

collagen

23

Characterizationofcollagen

matricespertinentto

collagen

traffickingandremodeling.Asmentioned

earlier,theanalyticalassessments

described

areconfined

primarilyto

commercially

available

sources

ofcollagen.This

focus

allowsthose

interested

incollagen

isolationfrom

tissuesources

usingvariousextractionandmodificationprotocolsto

employmethodsfrom

theliterature

orfrom

theirownlabs,butto

then

touse

theanalytical‘‘guide’’provided

herein

toassess

theircollagen

preparations.

266 ABRAHAM ET AL.


The rationale behind this review is the remarkable impact

collagen matrix structure has on cell and tissue outcomes

from recent studies of cellular aging as well as on retention of

stem cell differentiation potential.19,20,29,30 Collagen bioma-

terials have been used in many tissue-engineering applica-

tions. For example, osteoblast-like cells (Saos-2) adhere

more effectively and proliferate on xenogenic bone biomate-

rial containing collagen fibers compared to deproteinated

bone.31 Layering of collagen sheets and scaffolds seeded

with cardiomyocytes has enhanced cell survival with macro-

scopic pulsation similar to that of native heart tissue.32 Colla-

gen-based scaffolds are also prominent in the field of dermal

repair.33–35 In each of these reports, the characteristic of the

collagen biomaterial used in the experiments was important

to the success of the biological system under study. Further-

more, the widespread use of collagen biomaterials in many

biomedical applications with different rates and extents of

degradation, and where different release profiles of therapeu-

tics are sought,3,36,37 suggest that there exist important rela-

tionships between collagen structure and function in the

biomedical context. With such a significant impact of colla-

gen features on biological outcomes, it becomes essential to

consider an appropriate set of characterization tools so that

collagens are characterized in a comparative manner as an in-

tegral part of any study focusing on biological outcomes. Of

particular interest to the biopharmaceutical industry is to

have standard analytical tools and procedures that could be

used in characterization, scale-up, and comparability analysis

of delivery systems based on collagen.

The interaction of cells with a biomaterial, particularly at

the surface, determines adhesion and spreading and conse-

quently plays an important role in determining the pathways of

cellular differentiation, growth, and survival.38 In addition, a

biomaterial matrix for tissue engineering must have porosity

and mechanical stability suitable for the target cells and tissue

functions.34 For example, the goal of engineering blood vessels

using tissue engineering approaches has led to the utilization

of synthetic biopolymers to approximate the three-layered

structures present in native arteries.39 The tailoring of biomate-

rial properties to mimic those of the target tissue is desired to

help increase the chances of success of tissue-engineering

implants.36 In order to successfully mimic these chemical and

physical features of native tissues with collagen, both the tis-

sues and biomaterials must be extensively characterized. Char-

acterization tools for collagen must consider structural,

morphological, and chemical features because of the impor-

tance of physical and chemical factors on cell responses. In

addition, these features directly influence rates and extent of

collagen remodeling in vitro and in vivo, thus playing a major

role on functional outcomes. Bulk and surface properties need

to be considered because of its impact on stability, mechanical

performance, and cell interactions.

Objective

Despite the large number of studies with designed biomate-

rial surfaces, there remains the need to engineer biomateri-

als that can provide both surface and bulk requirements for

tissue-engineering matrices and also promote desired cell

responses in vivo for tissue repair. Although collagen is fre-

quently used as a biomaterial, the understanding of colla-

gen biomaterial characteristics as a function of cellular

responses is far from complete, particularly when consid-

ered in the light of the prominent role this family of fibrous

protein plays in vivo. The objective of this review is to es-

tablish a more consistent basis of collagen characterization,

so that biological responses can be more accurately related

to differences in structure, morphology, and chemistry.

These types of relationships are critical in order to optimize

and control cell and tissue outcomes on collagen-based bio-

materials, as well as to better predict and control rates and

extent of integration of in vitro prepared and/or grown tis-

sues in vivo. This insight should lead to the better design

and control of matrix structural and morphological features,

resulting in more predictable and relevant cell and tissue

outcomes in vitro and in vivo. We present information on

collagen biomaterial use and characterization along with

our results related to characterization of collagen matrices

pertinent to collagen trafficking and remodeling. As men-

tioned earlier, the analytical assessments described are con-

fined primarily to commercially available sources of

collagen. This focus allows those interested in collagen

isolation from tissue sources using various extraction and

modification protocols to employ methods from the literature

or from their own labs, but then to use the analytical ‘‘guide’’

provided herein to assess their collagen preparations.

ANALYTICAL METHODS

Overview

The characterization of collagen is divided into three major

areas in this review: (a) structural detail—mainly focused

on molecular mass, purity, helical content, and bulk ther-

mal properties, (b) chemical features—mainly focused on

surface elemental analysis and hydrophobicity, and (c) mor-

phological features at different length scales. In total, this

suite of analytical assessments of collagens can provide a

consistent basis for comparison of materials and thus bio-

logical outcomes with these materials. While this list of

characterization methods suggested for collagen biomateri-

als is not exhaustive, it provides a starting point for consis-

tency in analysis. Table III includes a technique summary

listing for each method, the collagen format needed, solu-

tion concentrations, information gathered, and major limita-

tions of the method.

Structural Information

Mass Spectroscopy. Collagens proteins have been

reported with molecular masses from 28340 to 300 kDa.41

Mass spectroscopy can be employed to provide molecular

mass data on collagen as well as the identification of

telopeptides and other potential contaminants in collagen



preparations that can directly impact cell functions.42 Ma-

trix-assisted laser desorption ionization time-of-flight

(MALDI-TOF) mass spectroscopy is commonly used. Pro-

tein fragments are dissolved in an organic acid, then dried

onto matrices—most often metals or ceramics.10 After exci-

tation with a laser, the protein fragments are accelerated in

an electric field.4 The detector identifies the proteins and

fragments by their mass and charge.11 Protein samples are

typically applied at �10 lM. The sample is spotted onto to

a metal MALDI plate after dissolution in a solution of

water and organic with a crystallized molecules such as

3,5-dimethoxy-4-hydroxycinnamic acid (sinapinic acid), a-cyano-4-hydroxycinnamic acid (alpha-cyano or alpha-ma-

trix), or 2,5-dihydroxybenzoic acid (DHB).42–49

Collagen chains (�94 kDa40 to 98 kDa42 exceed the

working range of many mass spectrometers. Therefore,

digestion of the �300 kDa collagen to �94- to 98-kDa

fragments is required. The lack of data reported for

digested collagen and the complication due to its high mo-

lecular mass provide significant challenges in mass spec-

troscopy assessments of collagens; thus, gel electrophoresis

is more commonly utilized (see later). Complications also

arise from low-molecular-mass collagen telopeptides (�6–

14 kDa42) that usually require a separate MALDI-TOF ma-

trix for appropriate analysis. Mass spectroscopy is also use-

ful to track bone remodeling and the formation of new

bone collagen, where the presence of telopeptides is com-

mon.45 The purity of the collagen source material can also

be assessed using mass spectrocscopy.46

Sodium Dodecyl Sulfate Polyacrylamide Gel Electro-

phoresis. Sodium dodecyl sulfate polyacrylamide gel elec-

trophoresis (SDS-PAGE) is most commonly used to assess

collagen source material purity and breakdown. SDS-PAGE

allows visualization of protein fragments by loading of pro-

tein samples in the wells of a thin gel, then using electric

voltage to drive the protein fragments through the gel. The

smallest protein fragments are least impeded by the gel ma-

trix and travel furthest through the gel. Coomassie blue or

silver stain are commonly used to visualize the protein

bands. Small sample amounts from nanogram to microgram

are needed, and molecular mass banding patterns are

obtained. Subsequent Western blots can be used to assess the

specificity of collagen type using monoclonal antibodies.47 A

summary of collagen materials used in tissue engineering,

including the characterization of molecular mass, is shown in

Table IV. SDS-PAGE gels commonly used for collagens are

4–20% polyacrylamide. Collagen samples can be loaded

directly in dilute acid solutions [0.1% glacial acetic acid

(GAA)].

The materials described earlier were purified by vendors

and provided as purified single type collagens. For samples

that are purified in the laboratory directly from tissues,

modified methods such as interrupted electrophoresis can

provide an additional tool to separate different collagen

types from a single tissue. Interrupted electrophoresis

begins with a collagen sample in the well. After the bandsTABLE

III.

CharacterizationTechniquesforCollagen

Collagen

Description

Mass

Spectroscopy

SDS-PAGE

CD

DSC

XPS

Contact

Angle

AFM

SEM

ESEM

Materialform

atSingle

strand

(ureatreated)

solution

Solution

Solutionin

cuvette

or

film

onplate

Dehydratedfilm

inDSCpan

Films,fibers,

gels

Filmsorfibers

Filmsorfibers

Films,fibers,

gels,sponges

Films,fibers,

gels,sponges

Concentration,

solvent

10lM,water

10–20lg

,

water

0.125lg

lL21

solutionor

100lgdry

weightfilm

5mgdry

weight

Films:

15–780

lgcm

22

Films:

15–780

lgcm

22

Films:

15–780

lgcm

22

Films:

15–780

lgcm

22

Films:

15–780

lgcm

22

Inform

ation

Mass

Size

Secondaryand

tertiary

structure

Denaturation

temperature,

heatcapacity

Atomic

composition

Hydrophobicity

Molecular

topography

Topography

Topography

Major

limitations

Massrange,

specialized

equipment

Denaturation

Deconvolution

softwarelimited

Largemass

needed

Specialized

equipment

Hydration,

surface

smoothness

Specialized

equipment

Artifacts

dueto

dehydration

Resolutionof

features

268 ABRAHAM ET AL.


migrate into the gel the current is interrupted, b-mercapto-

ethanol is then added to the wells and incubated, causing

the collagen helices to unwind at rates related to the disul-

fide content of the specific collagens. When this method

was applied to collagens isolated from human skin samples,

the migration of a1[III] chains was delayed when compared

to a1[I] chains, allowing resolution.53

Circular Dichroism. Circular dichroism (CD) utilizes the

differential absorption of circular polarized light in an

asymmetrical environment to assess structure.54 The amide

bonds of a protein in highly ordered regions such as a heli-

ces and b sheets have specific optical activity due to orien-

tation.54 CD has commonly been employed as a technique

to characterize the helical content of collagen.24,54,56 The

helical nature of collagen is responsible for the important

structural properties of tissues and in scaffolds used in tis-

sue-engineering. For example, CD has been used to confirm

collagen incorporation and structure in polymer–collagen

electrospun matrices designed as scaffolds for soft tissue-en-

gineering applications.57 CD has been used to assess the sus-

ceptibility of collagen to ultraviolet light based on loss of

helicity,58 to confirm the presence of collagen helical content

for collagen-like peptides56,59 and to study the enzymatic hy-

drolysis of collagen due to MMP-related reactions.51 CD has

also been used to characterize variation in collagen structure

in specific skeletal diseases such as osteogenesis imper-

fecta.60,61 Thermal denaturation melts the collagen, thus dis-

rupting the triple helix, and is usually an irreversible process

because of the complex self-assembly involved in proper

collagen chain associations and registry.62

The helical state of collagen in biomaterial and disease

applications can have significant effect on the cellular

remodeling responses. In fibroblastic matrix remodeling

in vitro, the rates of remodeling are greater, and cell health

is improved on denatured (wound-like) collagen versus

non-denatured (native-like) collagen.19 These observations

suggest that the presence of denatured collagen in a tissue-

engineering matrix might promote active remodeling neces-

sary for integration of implants. In several collagen disease

states including osteoporosis, osteogenesis, and bone metas-

tases, the remodeling of collagen plays an important role in

the pathology of the disease.46,63,64 In osteoarthritis, the

reduction in mechanical properties of subchondral bone has

been associated with an increase in denatured collagen.65

Since the helical content of collagen is critical to cell

responses both in vitro and in vivo, establishing the helical

content of collagen biomaterials is necessary. A summary

of collagen biomaterials used in tissue engineering charac-

terized by CD is shown in Table V.

The quality of CD data depends on sample concentration

and temperature. Sample concentrations must be controlled

and should be low enough (\0.125 mg mL21 for collagen)

to avoid saturation of the detector. Sufficient temperature

control is also required to avoid denaturation under experi-

mental conditions. Ellipticity data noting the angle of

polarization of light, reported in millidegrees, can be con-

verted to mean residue ellipticity [degree cm2 dmol21];

however, the molecular mass of the sample is required for

further conversion to molar ellipticity [dL mol21 dm21].

Since most commercially available collagens do not specify

molecular mass, this must be determined experimentally in

order to report mean residue ellipticity data (see section

above on molecular mass determination). Despite the com-

plications associated with collecting and comparing data on

collagen structure by CD, this remains a powerful tool to

assess helicity and thus degree of naturation/denaturation of

a sample used in biological studies. Thus, ellipticity, deter-

mined by CD, provides a measure of this structural feature,

and thus an assessment of the native/denatured state of the

collagen preparation.

Several methods for calculating helix content are avail-

able. For example, fhelix 5 [h]obs222/(240,000[1 2 2.5/chain

length]).66 Data conversion from millidegrees to mean resi-

due ellipticity using [h]222 5 h/(molar concentration 3 15

residues), in deg cm2 dmol21, allowing the calculation of

helix content as [h]222(observed)/[h]222

(max), where [h]222(max) is given

by 240,000(1 2 2.5/n), and n is the number of amino

acids in the peptide (Chakrabartty et al. 1991).71 Other

calculations rely on deconvolution programs to evaluate

helical content.73,74 In each case, the accuracy of the quan-

titative assessment of helicity depends strongly on the ac-

TABLE IV. Measures of Molecular Mass of Collagens

Collagen Material Technique/Findings Reference

Proteins synthesized on oocytes grown

with radio-labeled proline

Correlation of mass spectroscopy with polyacrylamide

gels for molecular weight analysis

48

Collagen extracted from rat tail SDS-PAGE for molecular size as a function of UV exposure 49

Sigma human type I collagen SDS-PAGE for molecular size as a function of MMP and

TIMP cleavages

50

Rat tail tendon type I collagen SDS-PAGE for molecular size as a function of MMP cleavages 51

Collagen extracted from human hip bones MALDI-TOF MS. to show the presence of C-telopeptides 42

Collagen extracted from fetal calf skin tissue Infrared (IR)-MALDI TOF MS detection of collagen triple helix 40

Type II collagen from fetal bovine cartilage SDS-PAGE and MS to characterize gelatinase B degradation

of collagen type II

46

N-terminal propeptide of human procollagen

(PINP) from amniotic fluid

Size determination of PINP by MS and SDS-PAGE 52



curacy of the solution concentration, molecular mass, and

amino acid content. These data can often be problematic

for collagens, as they are often not well defined. A major

limitation to the interpretation of CD data for collagens and

other fibrous proteins is that the current algorithms used for

conversion of signals to structural information are based on

globular proteins.

Differential Scanning Calorimetry. Differential scanning

calorimetry (DSC) provides direct determination of en-

thalpy (DH) by measuring the temperature dependence of

partial heat capacity.75 The difference in electrical energy

required to raise the temperature of the sample versus that

to raise the temperature of the reference solvent (buffer) is

normalized by the heating rate to calculate the difference

in heat capacity.75 With known masses and temperature

changes, the sample heat capacity and melting temperature

can be calculated.75

DSC is frequently used to characterize the bulk thermal

characteristics of a biomaterial,76 including crosslinked col-

lagen.77 Thermal properties of collagen-based scaffolds

provide information on transitions in the structural state,

reflecting initial primary (chemistry) sequence, structural

state, and degree of crosslinking, and also purity of sam-

ples. There is a wide variation in the protocols used to col-

lect data by DSC. Most often the researcher determines the

apparent Tm, as the thermal unfolding of large proteins like

collagen is usually irreversible. Incomplete drying of sam-

ples can lead to errors in determining the melting tempera-

ture (Tm).78 The rate of sample heating can impact thermal

transitions. For example, collagen-like peptides are heated

at rates no greater than 0.18C min21 for accurate determi-

nation of Tm. Thermal equilibration of collagen may take

as much as 40 min, requiring a heating rate of 0.0048Cmin21.62 Despite these issues, rates of heating used in

many DSC studies of collagen are often as fast at 108Cmin21.79 A summary of collagens analyzed by DSC is

shown in Table VI.

Chemical Information

X-ray Photoelectron Spectroscopy. X-ray photoelec-

tron spectroscopy (XPS) is commonly employed to charac-

terize the atoms present on the uppermost 10 nm of a

TABLE V. Circular Dichroism Assessments of Collagens and Associated References


Collagen-like helices from streptococcal proteins Unfolding of the helix was observable at 220 nm

after heat denaturation

66

Calf skin collagen and gelatin Lower intensity, redshifted CD for heat-denatured

collagen at 220 nm

67

Chick type I procollagen Tm of 428 C 58

Fetal calf skin collagen Characterization of collagen helix reduction with

denaturation

68

Collagen-like peptides Confirmation of the presence of triple helix 56

Collagen type I from bovine skin or rat tendon Characterization of collagen helix reduction with

denaturation

51

Bovine calf skin type I collagen and peptides Confirmation of the presence of triple helix 55

Collagen-like peptides Changes in helicity with side group modifications 59

Calf skin collagen Characterization of collagen and collagen peptides,

reduction of helix with heat treatment

69

Calf skin collagen also prepared with crosslinks Characterization of collagen helix reduction with

denaturation and SDS-PAGE

70

Sigma bovine type I, electrospun with polymers CD spectroscopy of released collagen confirmed collagen

incorporation and preservation of collagen structure

57

TABLE VI. Differential Scanning Calorimetry Assessments of Collagens


Isenglass collagen from fish, bovine hide

collagen

DSC for Tm’s 80

Collagen extracted from bovine tendon Higher denaturation temperatures for crosslinked

collagen scaffolds

33

Collagen helix-like peptides DSC for enthalpy change and Tm for various peptides 56

Rat skin collagen Evaluation of crosslinking, age-related changes 76

Rat tail tendon, albino rats Evaluation of collagen film and solution thermal transitions

as a function of UV irradiation

81

Adult bovine femur bone Tm changes as a function of c-irradiation 82

Collagen extracted from bovine muscle Tm changes as a function of sample moisture content 78

Human amnion Thermal characterization as a function of crosslinking 77

270 ABRAHAM ET AL.


material’s surface.83 After bombarding the material with X-

rays, measuring the emitted photoelectron numbers and

energies provides the basis for determination of the

amounts and chemical identities, respectively, of the atoms

present on the material’s surface.83,84 Because of the crude

preparation steps necessary to isolate collagen from animal

tissue, it is important to characterize the chemical composi-

tion to assess contaminants, material homogeneity such as

in film formation in tissue culture wells, and surface modi-

fications with adsorbed or chemically coupled growth, se-

rum, or adhesion factors. The elemental composition of

collagen biomaterials can be determined by XPS,85,86 and

this technique can also be used to confirm the biosynthesis

of collagen by cells.87 XPS has been used to determine the

presence of absorbed serum proteins on tissue-engineering

surfaces,88 and for the presence of a collagen coating to

improve the biocompatibility of titanium implants for bone

growth.89 A summary of collagens characterized by XPS is

shown in Table VII.

Contact Angle. The hydrophobic character of biomate-

rial surfaces influences cell adhesion and spreading, and

these surfaces are often characterized using water contact

angle measurements. Furthermore, contact angle is com-

monly employed as an indicator of surface chemical modi-

fication reactions to track successful chemical coupling

reactions, reflective of a change in surface hydrophobicity/

hydrophilicity. Using an contact angle goniometer, the

angle of contact of a small drop (sessile drop method) of

fluid placed on the surface of interest can be measured.92

Contact angle data for prepared collagen surfaces helps

predict and explain cell attachment data. Fibroblasts adhere

and proliferate preferentially on hydrophilic surfaces with

contact angles below 578.93,94 Surfaces with a contact angle

of 708 and a collagen-grafted polyethylene water contact

angle of 438 supported optimal fibroblast proliferation.95

Contact angles of ultrapure water on 1.0 mg mL21 Cellgen

IPG type I collagen cast films varied from 418 to 718.96

Collagen coatings applied to poly(e-caprolactone) films for

use as implants displayed contact angles that confirmed

changes in hydrophobicity upon grafting collagen to the

surface.91 The contact angle has also been related to the

synthesis of new collagen, with an increase in contact angle

from 428 to 1168.95 A summary of collagen material in tis-

sue-engineering applications characterized by contact angle

is shown in Table VIII.

Morphological Information

Surface Morphology. Cells respond to surface morphol-

ogy or roughness. In order to characterize the surface to-

pology of a collagen-based biomaterial, a variety of surface

imaging tools can be used including atomic force micros-

copy (AFM), scanning electron microscopy (SEM), envi-

ronmental scanning electron microscopy (ESEM), and light

microscopy. Each technique offers advantages and disad-

vantages associated with sample preparation and resolution,

thus often combinations of several microscopy techniques

are considered. The collection of SEM data requires appro-

priate sample preparation such as sputter coating with gold,

which can dampen surface resolution. The soft nature of

collagen can result in cracking of the coating during imag-

ing. Therefore, ESEM is often a more suitable choice, since

the problems with SEM are avoided, and resolution is usu-

ally sufficient although not as good as SEM.

Atomic Force Microscopy. Nanometer-scale resolution

is achieved with AFM, providing input on scales related to

surface receptors and protein interactions. Contact imaging

via tapping mode using the microscopic probe on the sur-

face of a sample is accompanied by measurements of force

deflection of the cantilever on which the probe is mounted,

to generate the readout of surface morphology. AFM has

been used to characterize the surface roughness of collagen

coated with polystyrene and oxidized polystyrene,80 and to

characterize surface roughness with addition of collagen

films on poly(e-caprolactone).91 The presence of collagen

castings from 0.5 mg mL21 bovine collagen intended to

mask poly-e-caprolactone hydrophobicity increased mean

surface roughness (Ra) from 46 to 60 nm.91 AFM has also

been used to pattern surfaces with collagens and collagen

peptides in the dip pen lithography mode with line resolu-

TABLE VII. X-ray Photoelectron Spectroscopy Analysis of Collagens


Sigma type I calf skin bovine collagen Film coating of polystyrene surfaces via C/N/O bond energies 90

Collagen films on poly(e-caprolactone) Presence of collagen films by [N]/[O] ratios 91

Type I bovine collagen (KNC SemedS collagen powder)

from Kensey Nash

Collagen-coated titanium surface [C]69.2 [O]17.1 [N]12.6

Collagen source material [C]69.1 [O]17.5 [N]11.7 89

TABLE VIII. Contact Angle Assessment of Collagens


Collagen-grafted polyethylene Water contact angle 438 6 38 95

Poly(e-caprolactone) grafted with Sigma calf skin

collagen type I

Hydrophobicity characterization, collagen-grafted polymer 458 91

Collagen extracted from rat tail Changes in contact angle with crosslinking 97



tion to 30- to 50-nm line widths.74 A brief summary of col-

lagens characterized by AFM is shown in Table IX.

Scanning Electron Microscopy. Depending on the spe-

cific model, SEM magnification of 3,0003 to 30,0003 can

be achieved and can be used to image the collagen sub-

strate and cells grown on these surfaces. A sputter-coated

thin layer of gold is required to facilitate imaging of the

surface. For example, the shape of human lung fibroblasts,

IMR-90 cells, growing on collagen has been related to cell

age using SEM, with older cells exhibiting a larger more

irregular morphology.103 Formulas have been developed

relating ratios of maximum to minimum cell length to char-

acterize cell morphology in the study of stromal cell

spreading.104 In collagen scaffolds engineered for artificial

dermis applications, SEM has been used to quantitate the

pore sizes of the scaffolds as well as the extent of collagen

crosslinking.33,105 The orientation of osteoblasts along pat-

terned collagen surfaces was examined using SEM to iden-

tify patterns conducive to bone formation.97 The adhesion

and spreading of bone marrow stem cells on silk biomate-

rial fibers for ligament repair has been imaged directly by

SEM.106 A summary of collagen biomaterials characterized

by SEM is shown in Table X.

Environmental Scanning Electron Microscopy. ESEM

uses a high vacuum, high humidity chamber to image sam-

ples without sputter-coating. Both native and denatured col-

lagen-cell samples have been imaged. For example, to

characterize self-assembled fibrils of collagen composites

for bone-tissue engineering, ESEM was used.108 In tissue-

engineering arterial constructs with collagen coatings,

ESEM was used to image cells.111 A summary of collagen

material in tissue-engineering applications characterized by

ESEM is shown in Table XI.

Light Microscopy. More routine observations of colla-

gen-cell constructs are frequently made with light micros-

copy. For example, light microscopy was used to image the

growth of chondrocytes on a collagen type I/III matrix

designed to improve regenerative capacity of hyaline artic-

ular cartilage.113 Light microscopy techniques are common

and provide at least a gross morphological assessment of

material features and cell interactions as a starting point for

the assessment of biological responses, such as cell adhe-

sion, spreading, and replication (Table XII).

CASE STUDY—CELLULAR REMODELINGCOLLAGEN BIOMATERIALS

To illustrate the importance of characterization of collagen

biomaterials, the relationship between collagen biomaterials

and matrix remodeling by human cells will be described.

These studies illustrate the importance of understanding

collagen structure related to cellular aging,19 the retention

TABLE IX. Atomic Force Microscopy Analysis of Collagens


Sigma type I calf skin collagen 67-nm banding and 150 nm diameter collagen fibrils via AFM 98

Collagen films on poly(e-caprolactone) Increased roughness (Ra 5 60 nm) with addition of collagen films 91

Collagen from bovine vertebrae 67-nm banding and 50–200 nm diameter collagen fibrils via AFM 99

Type I bovine skin collagen AFM to confirm build up of film coatings 100

Dentin collagen fibers Size distribution and repeat distances 101

Bovine dermal collagen Alignment of collagen fibers with AFM tip 102

Sigma type I calf skin bovine collagen Film coating of polystyrene surfaces via AFM roughness 90

TABLE X. Scanning Electron Microscopy Analysis of Collagens


Sigma type I collagen–chitosan matrices SEM morphology characterization of collagen crosslinking 107

Collagen extracted from bovine tendon Pore size 50–150 lm, porosity rate 94% 33

Type I collagen extracted from equine tendon SEM to characterize the morphology of the spray-dried

collagen composite for bone-tissue engineering

108

Collagen extracted from rat tail Cell alignment and orientation in comparison to collagen

crosslinking

97

Collagen type I from bovine Achilles tendons Collagen crosslink morphology dependency on freeze drying

temperature

39

Porcine temporomandibular joint disc SEM characterization of collagen fibers 109

2.6% collagen gel (Matrix Pharmaceutical) as

an adenovirus delivery vehicle

SEM to characterize the contact of how alveolar bone with the

dental implant surface

110

Type I collagen from bovine tendon Assessment of scaffold crosslinking in the presence of various

amino acids

111

272 ABRAHAM ET AL.


of differentiation potential of human stem cells toward

bone116 and adipose tissue, impact on phagocytosis,30 and

impact on rates of matrix remodeling to generate new

extracellular matrices.30 These biological outcomes and the

impact of collagen structure/morphology and chemistry on

these outcomes highlights the importance of appropriate an-

alytical characterization of biomaterial substrates for the

study of biological relevance.

Collagen Preparation

Details regarding reagents and related background can be

found in the earlier referenced papers. Rat tail collagen

type I was purchased from Roche Chemicals (Indianapolis,

IN) and collagen films were prepared as we have previously

reported.19 Briefly, collagen was dissolved at 2–88C in ster-

ile filtered 0.1% GAA at 5–10 mg mL21 over at least

3 days for complete dissolution. Once dissolved, the solu-

tion of collagen is diluted to working concentrations just

before use to generate the nondenatured (native) biomate-

rial surfaces. Denaturation is accomplished by a 60-min

treatment in a 508C water bath and confirmed by CD.20

Surface morphology, because of changes in collagen con-

centration, suggests that positive fibroblast growth14 occurs

on surfaces formed from collagen at 78 lg cm22, and addi-

tional concentrations were also applied to tissue culture

plastic (TCP) wells for study. The materials are dried at

room temperature in a vacuum drying oven, typically for

12–48 h, until no liquid remains. Also used in the collagen

evaluation gel (Figure 2) were type I bovine collagen from

Sigma (St. Louis, MO) and human placental collagen from

Calbiochem (San Diego, CA). After the initial assessments

of these various collagen sources, primarily by SDS-PAGE

for this case study, we selected just one collagen commer-

cial source for the remaining characterization assessments,

with a few exceptions, along the lines of the guide. The

exception to this plan was light microscopy.

Cells

IMR-90 human lung primary fibroblasts were purchased

from the American Type Culture Collection (ATCC, Mana-

ssas, VA) and cultured at 378C and 5% CO2 in 20% fetal

bovine serum, 77% eagle minimum essential medium

(MEM), 1% penicillin–streptomycin liquid, 1% L-gluta-

mine-(200 mM), and 1% MEM nonessential amino acids

solution (10 mM). Cells were split at confluence �1:10.

Cells with fewer than 30 cumulative population doubling

levels (PDL) were designated ‘‘young,’’117 and cells with

more than 48 cumulative PDLs were designated as

‘‘aged.’’118 Metabolism of several proteins is twofold

higher in very young cells (PDL 5 22) versus old cells (PDL

5 48).118 Cells were harvested at 70–80% confluence for

inoculation of the collagen surfaces. The IMR-90 cells were

selected for their high rates of collagen synthesis and distinct

morphological changes that occur with aging.119–122

Molecular Mass

The collagen samples were analyzed on an Applied Biosys-

tems Voyager-DE Pro MALDI mass spectrometer in linear

mode (Tufts University Core Protein Chemistry Facility,

Boston, MA). Sample preparation included 10-min heating

at 438C in 8M urea followed by buffer exchange to water

by dialysis. Matrices were DHB and sinipinic acid depend-

ing on molecular mass of the collagen sample, such as the

TABLE XI. Environmental Scanning Electron Microscopy Analysis of Collagens

Collagen Material Technique/Findings Ref.

Sigma rat tail type I ESEM to determine extent of cell coverage on collagen endothelial

arterial graft constructs

111

Cell generated ESEM evaluation of collagen fibrotic bundles in the formation of new

tissue

112

Type I collagen extracted from equine

tendon

ESEM to characterize the self-assembled fibrils of collagen composite

for bone-tissue engineering

108

TABLE XII. Light Microscopy Analysis of Collagens

Collagen Material Technique/Findings Ref.

2.6% collagen gel (Matrix Pharmaceutical) as

an adenovirus delivery vehicle

Light microscopy with histological staining to note the

formation of hew bone

110

Extracted collagen type II from porcine costa Micrographs of chondrocyte attachment to various ratio

polymer:collagen scaffolds

114

Type I/III collagen bilayer Light microscopy to show layering and porosity of

bilayer membrane

115

Rat tail type I collagen from BD Biosciences Examination of composite layers of dermal tissue

engineering construct

35



presence of telopeptides or other contaminants. Figure 1

shows MALDI data for the collagen, with expected masses

at 94,366.41, 47,365.12, and 31,585.99. Calculated theoreti-

cal values for a1(I) 93,915 Da and a2(I) 94,910.7 Da are

expected to vary with species, extent of glycosylation, and

extent of pepsin digestion.40 Collagen 1 alpha chain subu-

nits dissociate under heat treatment (necessary to dissociate

the three alpha chains for MALDI) giving masses roughly

around 31,100 and 45,500, identified by Dreiseward et al. as

a31 and a2140. Molecular masses of C-telopeptides of the a1chain of type 1 collagen are reported as follows: type I colla-

gen teloPeptide (ICTP) (a1Ca1Ca1H) 10,279 Da, divalent

a1Ca1H 5967 Da, divalent a1Ca2H 6,037 Da, monovalent

a1C smaller 3,730 Da and larger 4,326 Da, and histidinohy-

droxylysinonorleucine HHL crosslinked a1Ca2Ha1H (skin)

7,024 Da.42 The N-terminal propeptide of procollagen type I

(PINP) masses have been reported from 14,313 to 14,360.8

Da.52 No evidence of telopeptides was observed by MALDI

in the collagen samples prepared earlier.

Sodium Dodecyl Sulfate Polyacrylamide GelElectrophoresis

To characterize collagen molecular mass for comparison

with MALDI and to attain more quantitative information

(via densitometry), the samples were run by SDS-PAGE

(Figure 2). Type 1 collagens from three vendors (Roche

type I rat tail collagen, Sigma type I bovine collagen, and

Calbiochem type I human collagen) were compared and

were run as untreated solutions and also after treatment

with collagenase. The sources of collagen from Sigma and

Roche had fewer small peptide bands on the gels, indica-

tive of degradation in the solution samples. Upon partial

digestion with collagenase, all three sources of collagens

showed a reduction in the typical collagen bands (sizes of

Figure 2. Sodium dodecyl sulfate polyacrylamine gel for source col-lagen. The gel shows lanes 1–10 (left to right): 1, mark 12 standard;

2, Sigma bovine collagen; 3, roche rat tail collagen; 4, Calbiochem

human placental collagen; 5, mark 12 standard; 6, collagenase

digested Sigma bovine collagen; 7, collagenase digested Sigma bo-vine collagen; 8, collagenase digested roche rat tail collagen; 9, col-

lagenase digested Calbiochem human placental collagen; and 10,

Calbiochem collagenase (vs. type I collagen). The schematic bar tothe right denotes the expected positions and chain combinations

for collagen. The Sigma and Roche collagen samples are relatively

pure and nondegraded. The reduction of all of the main bands of all

three collagens with collagenase supports the presence of collagenin the sample.

Figure 1. Mass spectroscopy of collagen. Collagen MALDI 4000–100,000 Da shows the major col-lagen peaks with no evidence of telopeptides. The a chains �94 kDa are visible, and the scan con-

firms the presence of the expected size. Small amounts of typical cleavage products from MMP-1

activity (�[1/4] and [3/4] size) are present. The absence of telopeptide products or other size achains indicates a relatively pure collagen sample.

274 ABRAHAM ET AL.


roughly 100 kDa)52 and an accompanying appearance of

lower molecular mass bands on the gel. The differences in

purity of the three commercial sources of collagen suggest

that additional purification may be appropriate for some of

these source materials depending on the nature of the bio-

logical studies to be conducted.

Circular Dichroism

CD data were collected using a Jasco J 710 Spectropo-

larimeter (Easton, MD) at 258C from 190 nm to 260 nm

with 0.05-nm step resolution, 10 nm min21 collection, an

accumulation rate of 4, response of 16 s, band width at

1.0 nm, and sensitivity of 50 millidegrees. Data were con-

sidered valid for the range of the instrument when the

‘‘HT’’ (Jasco labeling of voltage) photomultiplier voltage

was below 650 V. CD for collagen solutions was collected

at a concentration of 0.125 mg mL21. For collagen films,

drops of �100 lL of 0.5 lg lL21 collagen were dried on

quartz Suprasil (QS) 0.01-mm flat cuvette plates from

Hellma (Plainview, NY) for analysis.

CD was used to examine helical content as a function

of heat denaturation (temperature melts), as a reflection of

structure and chemistry. For both native and denatured

(1 h, 508C heat treated) collagen, the helical content of

the collagen solutions and the films was confirmed by CD

(Figure 3). The CD curve for the nondenatured collagen

shows the expected profile for a helical collagen molecule

including maxima and minima at 221 and 197 nm, respec-

tively. Both of the denatured collagen samples show a sig-

nificant decrease in the positive peak at 221 nm as well as

a significant reduction in the negative peak at 197 nm.

These data confirm the helical content of the nondena-

tured collagen and also indicate a significant reduction of

helical content for denatured collagen that had been heat-

treated. There was no significant difference in helicity as

a function of the concentration at which the collagen was

heat-treated. These same data are presented in Table XIII

after deconvolution using the CDNN program.73 Using a

back propagation network model with a single hidden

layer between input and output, the CDNN program cal-

culated five different secondary structure fractions (helix,

parallel and antiparallel beta-sheet, beta-turn, and random

coil).73

Although Figure 3 shows a reduction in ellipticity at

h222 as is expected for the denatured collagen samples,

Table XII demonstrated the complication of using deconvo-

lution programs based on globular proteins to evaluate CD

data of collagen samples. The CDNN program calculates

less helical content in the nondenatured samples compared

to the denatured samples, an incorrect conclusion based on

visual inspection of Figure 3. Quantification of alpha helix

and helix reduction requires knowledge of the molar con-

centration and the number of residues in the sample.71,72

Errors in estimating these values may also lead to some

errors in applying the CDNN algorithms. The more likely

source of error is in the reference proteins associated with

the CDNN program. The calculations of secondary struc-

ture depend on comparisons to model proteins (globular)

embedded in the CDNN program.73 For accurate ‘‘auto-

mated’’ calculation of molecule shape, a program with

fibrillar reference molecules is needed.

CD was applied directly to collagen films (Figure 4)

composed of 100 lL of 0.5 lg lL21 (50 lg) collagen-driedover a surface area of �0.8 cm diameter. For the �0.25-

cm2 film area examined, film density was �199 lg cm22.

The spectra for these films had the profile expected for col-

lagens. Likely because of film opacity there was a shift of

the spectra with the maxima shifted to 225 nm and the

minima shifted to 206 nm. However, there remained clear

evidence for significant reduction in helical content for the

films prepared from denatured collagen when compared to

the films prepared from nondenatured collagens. Using the

CDNN program on the film data yielded similar results to

those for the liquid collagen samples.

TABLE XIII. Deconvolution of Circular Dichroism Data UsingCDNN74 Program

Secondary

Structure

Type

% Structurea

Nondenatured

Denatured at

0.5 mg mL21Denatured at

1.0 mg mL21

% helix 5.9 12.4 10.6

% antiparallel 61.8 42.5 47.8

% parallel 4.6 7.4 6.9

% beta turn 20.8 16.3 16.2

% random coil 6.9 21.5 18.6

aEach CD sample was run at 0.125 mg mL21. The concentrations listed above

are the concentrations at which the samples were during the 508C denaturing heat

treatment.

Figure 3. Circular dichroism on collagen solutions. Shown are the

elipticity data for collagen on solutions of 0.125 mg mL21 collagen.

Samples were (—) Denatured at 0.5 mg mL21, (- -) Denatured at1.0 mg mL21, and (h) Nondenatured. Typical alpha helical protein

structure is observed for the nondenatured collagen. The expected

reductions in CD amplitude at 195 and 221 nm are seen for the

denatured collagen samples. This helical reduction is observed forsamples irrespective of the concentration of the sample during

denaturation.



Differential Scanning Calorimetry

Several attempts were made to characterize collagen ther-

mal transitions using a TA Instruments temperature modu-

lated DSC, TA2920 MDSC (New Castle, DE). For cooling,

a TA Instruments liquid nitrogen cooling accessory

(LNCA) was used. Dry nitrogen gas was purged into the

TMDSC cell at a flow rate of 20 mL min21. The standard

DSC was carried out with a heating rate of 58C min21

from 220 to 2008C and a cooling rate of 208C min21.

Roughly 5 mg of a 0.5 mg mL21 sample of dried nondena-

tured collagen film was added to an aluminium DSC pan.

A similar weight empty reference pan was used as a con-

trol (Figure 5). Although a thermal transition is suggested

by the data, integration for calculation of Tm is not possi-

ble. This limitation is due to the level of noise in the data,

a function both of the difficulty of adding sufficient weight

of collagen into a sample pan and also the sensitivity of

the instrument. As described earlier, with a slower heating

rate (not available on all DSC systems) or a more advanced

DSC system, improved thermal transitions could be deter-

mined, as are reported in the literature.

X-ray Photoelectron Spectroscopy

Sample surfaces were characterized using a Surface Sci-

ence Instruments (Mountain View, CA) Model SSX-100

XPS. Each sample was subjected to triplicate elemental

scans: 1000 nm, resolution 4, window 100 eV, at varied

positions in the well. Scans were conducted with a charge

neutralizer flood gun at 5 eV and with a nickel wire mesh

over the sample surface to prevent charging. After survey

scans to identify elements present, environmental scans for

C, N, and O were conducted. XPS was used to verify the

C/N/O ratios expected for uncontaminated TCP and colla-

gen films.29 To assess contaminants in the collagen prepa-

ration, the presence of the expected collagen C/N/O ratios,

and film coverage over all areas of sampling, XPS analysis

was conducted on denatured and nondenatured collagen

films [Figure 6(a,b)]. The elemental scans showed only car-

bon (C), nitrogen (N), and oxygen (O). The ratios of C, N,

and O were those expected for TCP and collagen. Detec-

tion is possible to about 0.1 atomic %, with accuracy of

element concentrations at less than 10 lM.123 All sample

concentrations of collagen, denatured and nondenatured,

showed similar C/N/O ratios to confirm that the collagen

matrices were free from significant contamination with sili-

cone, which can sometimes be a problem in collagen prep-

arations, and that the coatings on the plates were

continuous.

Contact Angle

Static contact angles were measured using a Rame-Hart

NRL CA contact angle Gonimeter (Mountain Lakes, NJ).

Measurements were made minimally in triplicate by the

sessile drop method. Hydrophobicities of denatured and

nondenatured collagen films of concentrations 15.6 lgcm22 to 780 lg cm22, were assessed via water contact

angle measurements (Figure 7). From collagen concentra-

tions 32.5–468 lg cm22, there was an increase in contact

angle with increasing collagen concentration for both the

denatured and the nondenatured samples. The increase in

hydrophobicity with increasing concentration of collagen

may be related to the accompanying increase in surface

roughness. Both the surface roughness and the hydropho-

bicity increases with concentration may be related to the

better cell growth and survival as observed for cells grown

on 78 and 156 lg cm22 collagen matrices.19 An increase

Figure 4. Circular dichroism on collagen solutions. Shown are the

elipticity data for collagen films, where 100 lL of 0.5 mg mL21 ofcollagen was dried directly on quartz Suprasil (QS) 0.01-mm flat

cuvette plates: (- -) Denatured (—) Gelatin (h) Nondenatured. Typical

alpha helical protein structure is observed for the nondenatured

collagen. The expected reductions in CD amplitude near 195 and221 nm are seen for the denatured collagen sample. The slight

offset in wavelength is expected due to the width of the cuvette on

which the films were dried. These data confirm that the dried colla-

gen films used in many tissue-engineering studies have similar heli-cal characteristics to their counterpart solutions.

Figure 5. Differential scanning calorimetry shown is the heat flowdata for DSC of roughly 5 mg of a 0.5 mg mL21 collagen sample.

Data were collected in a TA Instruments temperature modulated

DSC, TA2920 MDSC (New Castle, DE). The graph demonstrates the

challenges in collecting data for collagen samples due to equipmentlimitations or sample preparation/amounts.

276 ABRAHAM ET AL.


in hydrophobicity found with increasing collagen concen-

tration may relate to the phenomena of ‘‘pillars’’ in surface

topology leading to increased surface roughness.124 These

data may help explain the poorer cell growth observed on

higher collagen concentrations.19

Atomic Force Microscopy

AFM imaging was performed in tapping mode on a Dimen-

sion 3100 Nanoscope III with tapping mode etched silicon

probes (TESP), SMP, and DNP20 contact mode probes

(Digital Instruments, Santa Barbara, CA). TESP probes

have a cantilever length of 225 lm and a spring constant

of 1–5 N m21. Rotated TESP AFM tips have the same

spring constants and cantilever lengths as the TESP probes,

except that the tips rotated at a 158 angle to allow for bet-

ter visualization of high-aspect ratio features. Imaging was

achieved with 5- to 100-lm scan widths at rate of 0.5–

1.0 Hz. Data were collected on undisturbed collagen sam-

ples in phosphate buffered saline (PBS) in a 35-mm dish.

Large differences in surface roughness as a function of col-

lagen concentration were observed. The phase data surface

images typical of a 780 lg cm22 denatured collagen sam-

ple are shown in Figure 8. From the angled image showing

a portion of the surface in an edge on view, it can be

observed that the collagen surface has extensive roughness.

From the perspective of the cell, this 100 lm by 100 lmcollagen surface offers a bed of blunt spikes. The increas-

ing roughness observed with increasing collagen concentra-

tion corresponded to decreasing cell viability on the highest

collagen concentrations.19

To determine whether the increase in hydrophobicity

with increasing concentration correlated to surface rough-

ness, AFM was conducted with samples of varying concen-

trations. The AFM height images for 0.1% GAA on TCP

(GAA/TCP) and on denatured collagen concentrations of

15.6 to 780 lg cm22 are shown in Figure 9(a–g) . Each

scan is for a 100-lm square area. The height scales are

200 nm. The GAA on TCP plate surface indicates a regular

pattern of small spikes. All 35-mm TCP plates wet, dry,

and with and without GAA, showed similar patterns. Even

at the lowest collagen concentrations the collagen films

filled the valleys of the TCP topology and create a

smoother surface. Also visible in the collagen film images

are a series of more frequent surface topology spikes with

increasing concentration. The 15.6 and 31.5 lg cm22 sam-

ples show increasing smoothing of the TCP topology and a

small number of 50-nm to 200-nm spikes. For the samples

prepared from 78 and 156 lg cm22, there was no evidence

of the TCP topology, and the 50-nm to 200-nm spikes

associated with the collagen film are more frequent. The

Figure 6. (a) X-ray photoelectron spectroscopy on denatured collagen films. Surface content of the

following atoms is shown (j) Carbon, ( ) Nitrogen, (h) Oxygen. TC is tissue culture plastic. GAA istissue culture plate with 0.1% GAA (the collagen solution buffer) as control. The presence of colla-

gen-associated atoms in samples at each film thickness confirms that for each area of the film

tested, the plate is fully covered with collagen. The absence of significant content of noncollagen

atoms indicates a lack of contamination of the samples. (b) X-ray photoelectron spectroscopy onnondenatured collagen films. Surface content of the following atoms is shown (j) Carbon, ( ) Nitro-

gen, ( ) Oxygen.

Figure 7. Contact angle on collagen films. Shown is the water con-tact angle for the following surfaces: (h) tissue culture plastic; ( )

glacial acetic acid; ( ) denatured collagen; (j) nondenatured colla-

gen. The increase in hydrophobicity with collagen concentrationmay be related with increasing surface roughness at higher collagen

concentrations and illustrates the challenges in assessing surface

energy of films where surface morphology (smoothness) is an issue.



468 and 780 lg cm22 collagen samples have mountain-like

spikes up to and exceeding 200 nm in their topology cover-

ing most of the film surface. This increasing surface rough-

ness observed visually can also be quantitated using root

mean square (RMS) roughness calculations in the Digital

Instruments (DI) software (Figure 10). RMS roughness

increased significantly from 29 for GAA on TCP to 71 for

the denatured collagen films at 780 lg cm22. The increas-

ing surface roughness may help explain cell growth and

survival on the samples prepared from the solutions of col-

lagen containing 78 and 156 lg cm22.19 To confirm that

surface topology was due to the collagen film, any colla-

gen-related structure should be altered by heat treatment at

658C and to a greater extent at 858C. In Figure 11(a–c) a

156 lg cm22 denatured collagen sample is shown before

and after 65 and 858C heating intended to reduce surface

Figure 8. AFM image of collagen surface roughness. Digital Instruments Nanoscope, (left) 100-lmscan size, 0.5003-Hz scan rate, 256 samples, phase data image, 908 data scale; (right) 30-lm scan

size, 1.0-Hz scan rate, 256 samples, phase data image, 908 data scale. The surface roughness and

scale of surface topology of a high concentration collagen film is observed.

Figure 9. (a–g) Denatured collagen (0, 16, 31, 78, 156, 468, 780 lg cm22) surface roughness. Digi-

tal Instruments Nanoscope, 100-lm scan size, 0.5003-Hz scan rate, 512 samples, height dataimage, 200-nm data scale. The increasing surface roughness of collagen films with increasing colla-

gen concentration is observed. [Color figure can be viewed in the online issue, which is available at

www.interscience.wiley.com.]

278 ABRAHAM ET AL.


roughness due to the collagen structure by ‘‘melting out’’

the topology. The heating steps resulted in statistically sig-

nificant reduced roughness. These observations support the

hypothesis that the roughness observed in nonheated films

was due to the collagen structure.

Scanning Electron Microscopy

Cells that had been grown on collagen surfaces were fixed

with 2.5% glutaraldehyde in 0.1M sodium cacodylate, then

washed with 0.1M sodium cacodylate.107 Samples were

dehydrated by soaking in a graduated series of alcohol

washes. Samples were sputter-coated using a Polaron

SC502 Sputter Coater (Fisson Instruments, UK). SEM

images were collected using a JEOL LSM-840-A SEM

(Peabody, MA). Some of the denatured collagen film sam-

ples collapsed under the force of the sputter coating and

the vacuum of the SEM imaging; thus, these films lacked

sufficient structural integrity for this analysis. A sufficient

fraction of the films survived processing to permit the col-

lection of images. The roughness of nondenatured collagen

samples for GAA on TCP and for the collagen samples

prepared from 78, 156, 312, and 780 lg cm22 are shown

both with and without cells via SEM imaging (Figure 12).

Images are shown at 3,0003 and 1,0003 to depict both

single cells and groups of cells. Both PDL 33 and PDL 48

cells grown on nondenatured collagen are shown. For both

the image rows without cells, increased surface topology

roughness can be observed with increasing collagen con-

centration. For both the PDL 33 and the PDL 48 cells, bet-

ter adhesion to the collagen films, more contact points

(suggesting matrix phagocytosis), and fewer irregular cells

shapes were observed at the middle collagen concentrations.

The cells grown on 156 lg cm22 nondenatured collagen

were observed with the fewest of the age-related morphol-

ogy indicators, demonstrated fewer senescence related fea-

tures and also supported by assessments of cell morphology,

biochemistry, and transcript measures related to these age-

related outcomes.23,108 These observations correspond to

quantitation of senescence-associated b-galactosidase assays

for cell function.19 Since cell death and low proliferation

rates are a common problem in cell expansion and use on

biomaterial matrices, identifying collagen matrix properties

that lead to improved cell heath may allow for the design of

more successful tissue engineering constructs.

The SEM images of the collagen surfaces support, along

with the AFM images, increased surface roughness of the

collagen surfaces with increasing concentration used in film

preparation. The SEM images also allow assessment of cell

morphology that is not available via AFM. IMR-90 fibro-

blasts showed significant changes in cell morphology with

age. Increases in cell size, cell surface roughness, decreased

proliferation rates, and increased senescence-associated b-galactosidase expression are all associated with aging in

IMR-90 cells.125 From the SEM images, it can be con-

cluded that the cells grown on the highest concentrations of

nondenatured collagen were more phenotypically aged than

cells grown on the 156 lg cm22 nondenatured collagens.

Figure 10. Collagen surface roughness quantitation of AFM data.

Route mean square (RMS) roughness as quantified by the DigitalImaging AFM software was calculated for 100-lm areas, then aver-

aged over three samples at each collagen concentration. The

increase in RMS roughness with increasing collagen concentration

is shown. Statistical analysis was preformed for 3–10 samples ateach concentration.

Figure 11. (a–c) Denatured collagen (156 lg cm22) surface roughness—(a) nonheated (b) reduced

by heating at 658C, and (c) reduced by heating at 858C. Digital Instruments Nanoscope, 100-lmscan size, 0.5003-Hz scan rate, 512 samples, height data image, 200-nm data scale. The reduction

in surface roughness with heating is shown to demonstrate that the surface roughness observed isrelated to collagen structure. [Color figure can be viewed in the online issue, which is available at

www.interscience.wiley.com.]



Figure 12. Scanning electron microscopy on nondenatured collagen films. For all six rows ofimages, the collagen concentration increases (left to right) 0.1% GAA and 0, 78, 156, 312, and

780 lg mL21 collagen. The rows (top to bottom) show 10-lm size images of collagen with no cells,

10-lm size passage 19 ‘‘old’’ cells, 10 lm size passage 13 ‘‘young’’ cells, 30 lm size images of col-lagen with no cells, 30 lm size Passage 19 ‘‘old’’ cells, and 30 lm size passage 13 ‘‘young’’ cells.

Increasing surface roughness is observed as a function of increasing collagen concentration. Cells

appear ‘‘younger’’ (smaller, smoother edges, fewer processes) on the lower collagen concentrations

versus the higher collagen concentrations. Cells similarly appear ‘‘younger’’ on the denatured colla-gen versus the nondenatured collagen.

280 ABRAHAM ET AL.


Environmental SEM

ESEM images were collected on a FEI model Quanta 200

ESEM (Hillsboro, OR) with a tungsten filament and a pelt-

ier stage. Working distances were 6–10 mm, tilt was 258,temperature was 48C, and pressure was �2.75 Torr. ESEM

was used to observe cellular interactions with collagen mat-

rices with as little sample processing (fewer artifacts) as

possible. ESEM images for GAA on TCP and collagen

samples from 156, 468, and 780 lg cm22 with cells are

shown in Figure 13. For all collagen concentrations, the

cells grown on denatured collagen samples are more regu-

larly shaped and appear more attached to the underlying

collagen matrices, and may appear more closely in contact

with the substrate due to phagocytosing of the collagen ma-

trix. The cells grown on the denatured collagen are simi-

larly healthy when compared to those on the GAA on

tissue culture plastic wells. The cell in the 780 lg cm22

image is particularly notable for its nonattached appear-

ance, as if it is trying to remove itself from the surface.

The images show some of the collagen surface characteris-

tics as well as the cellular interactions with the collagen

substrates. This provides some assessment of the morphol-

ogy of the IMR-90 cells in relation to cell age, which may

be helpful in selecting collagens for tissue-engineering mat-

rices with the goal of promoting better cell health and sur-

vival. Compared with SEM, ESEM offers benefits in

avoiding sputter coating with metal that can generate arti-

facts, and generates fewer vacuum-induced artifacts. How-

ever, there is generally a loss in resolution with ESEM,

which for cell characterization studies is usually not an

issue in terms of gross morphological assessments.

Optical Microscopy

Cell images were captured using a Zeiss Axiovert S100

microscope (Thornwood, NY) equipped with a Sony

Exwave HAD 3CCD color video camera (Shinagawa, To-

kyo, Japan). Images were processed with Scion Image for

windows v4.0.2 software (Fredrick, MD). Image overlays

for fluorescence images used Adobe Photoshop 5.0 or Corel

Photo-Paint 10 software. In order to determine the extent

of cell growth and the general morphology of the cells,

light microscopy images were collected. Several light mi-

croscopy images of IMR-90 cells on Sigma and Roche type

I collagen at various concentrations are shown in Figure 14.

Although the light microscope images are not as detailed

as those from AFM and SEM, they provide immediate,

low-cost images, that can be collected in most laboratories.

Figure 13. Environmental scanning electron microscopy on collagen-cell surfaces. (A) P21 cells,

0.1% glacial acetic acid, (B) P21 cells, denatured collagen, 156 lg cm22, (C) P21 cells, denatured

collagen, 468 lg cm22, (D) P21 cells, denatured collagen, 780 lg cm22, (E) P21 cells, nondenatured

collagen, 156 lg cm22, (F) P21 cells, denatured collagen, 468 lg cm22, (G) P21 cells, nondenaturedcollagen, 780 lg cm22. Similar to SEM, increasing surface roughness is observed as a function of

increasing collagen concentration. Cells appear ‘‘younger’’ (smaller, smoother edges, fewer proc-

esses) on the lower collagen concentrations versus the higher collagen concentrations. Cells simi-larly appear ‘‘younger’’ on the denatured collagen versus the nondenatured collagen. In ESEM, the

lack of a gold coating allows for better visualization of the cellular reaction to the collagen surfaces.

Individual cells can be observed to be spreading on ‘‘favorable’’ collagen surfaces or contracting

from ‘‘nonfavorable’’ collagen surfaces.



Additionally, light microscopy images can be taken without

destructive preparation of the samples, allowing for further

cell culture postimaging.

Case Study—Conclusions

Several analytical tools were used to determine details of col-

lagen structure, morphology, and chemistry in this ‘‘case

study.’’ These methods were used to assess increases in colla-

gen roughness and the observance of cell morphologies with

relationship to helicity and topography. The cells grown on

denatured collagens appeared healthier than those grown on

nondenatured collagens and higher collagen substrate con-

centrations. We have also related these results to the cell

aging and collagen trafficking results reported elsewhere.19,30

SUMMARY

The characterization of biomaterial matrices is essential to

the design of intelligent tissue engineered matrices as well

as to provide comparative data among studies from differ-

ent laboratories. Cell responses to collagen matrices depend

on many features of these biomaterials, including secondary

and tertiary structure, chemical composition, hydrophobic-

ity, and surface roughness, among others. Techniques that

provide more thorough characterization of the matrices are

crucial to identifying optimal collagen-based biomaterials

for matrix fabrication and to relate the data to cell biology.

We have described the benefits and limitations of several

of the methods of characterization that can be applied to

collagen biomaterials and provide an initial basis for intra-

and interstudy comparisons. These types of biomaterial

characterizations are beneficial to guide the design of bio-

material tissue-engineering matrices and for the evaluation

of cellular responses to these matrices.

We have retained a focus in the analytical guide and in

the case study on commercial sources of collagens to pro-

vide a starting point for assessments that can be considered.

Many researchers prefer to isolate their own collagen, such

as from tendons or rat tails. The isolation procedures for

these extractions are well-described in the literature. Once

carried out, similar analytical tools as described in this pa-

per, as a guide to assessments of the isolated collagens, can

be considered for these tissue-derived sources of materials.

In a similar fashion, variations in the presence of telopepti-

des, contaminating ECM components, or crosslinking pro-

cedures, are some issues that may be encountered, which

can at least in part be assessed with the tools outlined in

this guide.

REFERENCES

1. Burgeson RE, Nimni ME. Collagen types. Molecular structureand tissue distribution. Clin Orthop 1992;282:250–272.

2. Patino MG, Neiders ME, Andreana S, Noble B, Cohen RE.Collagen: An overview. Implant Dent 2002;11:280–285.

3. Friess W. Collagen—Biomaterial for drug delivery. Eur J PharmBiopharm 1998;45:113–136.

4. Burke JF, Yannas IV, Quinby WC Jr, Bondoc CC, Jung WK.Successful use of a physiologically acceptable artificial skinin the treatment of extensive burn injury. Ann Surg 1981;194:413–428.

5. Eaglstein WH, Falanga V. Tissue engineering and the devel-opment of Apligraf, a human skin equivalent. Cutis 1998;62(1Suppl):1–8.

6. Meaney Murray M, Rice K, Wright RJ, Spector M. The effectof selected growth factors on human anterior cruciate liga-ment cell interactions with a three-dimensional collagen-GAGscaffold. J Orthop Res 2003;21:238–244.

7. Angele P, Kujat R, Nerlich M, Yoo J, Goldberg V, JohnstoneB. Engineering of osteochondral tissue with bone marrowmesenchymal progenitor cells in a derivatized hyaluronan-gel-atin composite sponge. Tissue Eng 1999;5:545–554.

Figure 14. Light microscopy images shown are IMR-90 cells on (A) 10 mg mL21 Sigma collagen,

(B) 5 mg mL21 Sigma collagen, (C) 1 mg mL21 Sigma collagen, (D) 0.1 mg mL21 Sigma collagen,(E) tissue culture plastic, (E) 1 mg mL21 Roche Collagen, and (F) 1 mg mL21 Roche Collagen. Light

microscopy is a convenient tool to show cell numbers and general morphology, while little informa-

tion is gained about the collagen surface. [Color figure can be viewed in the online issue, which is

available at www.interscience.wiley.com.]

282 ABRAHAM ET AL.


8. Brekken RA, Sage EH. SPARC, a matricellular protein: Atthe crossroads of cell-matrix. Matrix Biol 2000;19:569–580.

9. Blair HC, Zaidi M, Schlesinger PH. Mechanisms balancingskeletal matrix synthesis and degradation. Biochem J 2002;364(Pt 2):329–341.

10. Hay ED. Cell Biology of Extracellular Matrix. New York:Plenum; 1991. xvii, 468 pp.

11. Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P.Molecular Biology of the Cell. New York: Garland Science;2002.

12. Ricard-Blum S, Dublet B, van der Rest M. UnconventionalCollagens. Grenoble Cedex: Oxford University Press; 2000.155 pp.

13. Badii F, Howell NK. Elucidation of the effect of formalde-hyde and lipids on frozen stored cod collagen by FT-Ramanspectroscopy and differential scanning calorimetry. J AgricFood Chem 2003;51:1440–1446.

14. Awad HA, Butler DL, Harris MT, Ibrahim RE, Wu Y, YoungRG, Kadiyala S, Boivin GP. In vitro characterization of mes-enchymal stem cell-seeded collagen scaffolds for tendonrepair: Effects of initial seeding density on contractionkinetics. J Biomed Mater Res 2000;51:233–240.

15. Zimmermann WH, Melnychenko I, Eschenhagen T. Engi-neered heart tissue for regeneration of diseased hearts. Bioma-terials 2004;25:1639–1647.

16. Viravaidya K, Shuler ML. The effect of various substrates oncell attachment and differentiation of 3T3-F442A preadipo-cytes. Biotechnol Bioeng 2002;78:454–458.

17. Bornstein MB. Reconstituted rattail collagen used as substratefor tissue cultures on coverslips in Maximow slides and rollertubes. Lab Invest 1958;7:134–137.

18. Michalopoulos G, Pitot HC. Primary culture of parenchymalliver cells on collagen membranes. Morphological and bio-chemical observations. Exp Cell Res 1975;94:70–78.

19. Abraham LC, Vorrasi J, Kaplan DL. Impact of collagen struc-ture on matrix trafficking by human fibroblasts. J BiomedMater Res A 2004;70:39–48.

20. Volloch V, Kaplan D. Matrix-mediated cellular rejuvenation.Matrix Biol 2002;21:533–543.

21. Kleinman HK, McGarvey ML, Liotta LA, Robey PG, Trygg-vason K, Martin GR. Isolation and characterization of type IVprocollagen, laminin, and heparan sulfate proteoglycan fromthe EHS sarcoma. Biochemistry 1982;21:6188–6193.

22. Bornstein P, Traub W. The Chemistry and Biology of Colla-gen. New York, NY: Academic Press; 1979.

23. Gallop PM, Seifter S. Preparation and properties of solublecollagens. Meth Enzymol 1963;6:635–641.

24. Niyibizl C, Fietzek PP, van der Rest M. Human placenta typeV collagens. Evidence for the existence of an alpha 1(V)alpha 2(V) alpha 3(V) collagen molecule. J Biol Chem 1984;259:14170–14174.

25. Kordula T, Banbula A, Macomson J, Travis J. Isolation andproperties of stachyrase A, a chymotrypsin-like serine proteinasefrom Stachybotrys chartarum. Infect Immun 2002;70:419–421.

26. Einbinder J, Schubert M. Binding of mucopolysaccharidesand dyes by collagen. J Biol Chem 1951;188:335–341.

27. Trentham DE, Townes AS, Kang AH. Autoimmunity to typeII collagen an experimental model of arthritis. J Exp Med1977;146:857–868.

28. Miller EJ. Isolation and characterization of a collagen fromchick cartilage containing three identical a chains. Biochemis-try 1971;10:1652–1659.

29. Abraham LC, Dice JF, Lee K, Kaplan DL. Phagocytosis andremodeling of collagen matrices. Exp Cell Res 2007;313:1045–1055.

30. Abraham LC, Dice JF, Finn PF, Mesires NT, Lee K, KaplanDL. Extracellular matrix remodeling—Methods to quantifycell–matrix interactions. Biomaterials 2007;28:151–61.

31. Basle MF, Grizon F, Pascaretti C, Lesourd M, Chappard D.Shape and orientation of osteoblast-like cells (Saos-2) areinfluenced by collagen fibers in xenogenic bone biomaterial.J Biomed Mater Res 1998;40:350–357.

32. Shimizu T, Yamato M, Kikuchi A, Okano T. Cell sheet engi-neering for myocardial tissue reconstruction. Biomaterials2003;24:2309–2316.

33. Zhang L, Ma D, Wang F, Zhang Q. The modification of scaf-fold material in building artificial dermis. Artif Cells BloodSubstit Immobil Biotechnol 2002;30:319–332.

34. Ng KW, Khor HL, Hutmacher DW. In vitro characterizationof natural and synthetic dermal matrices cultured with humandermal fibroblasts. Biomaterials 2004;25:2807–2818.

35. Torkian BA, Yeh AT, Engel R, Sun CH, Tromberg BJ, WongBJ. Modeling aberrant wound healing using tissue-engineeredskin constructs and multiphoton microscopy. Arch Facial PlastSurg 2004;6:180–187.

36. Friess W. Collagen in drug delivery and tissue engineering.Adv Drug Deliv Rev 2003;55:1529–1530.

37. Knowles GC, McKeown M, Sodek J, McCulloch CA. Mecha-nism of collagen phagocytosis by human gingival fibroblasts:Importance of collagen structure in cell recognition and inter-nalization. J Cell Sci 1991;98(Pt 4):551–558.

38. Diener A, Nebe B, Luthen F, Becker P, Beck U, NeumannHG, Rychly J. Control of focal adhesion dynamics by mate-rial surface characteristics. Biomaterials 2005;26:383–392.

39. Buijtenhuijs P, Buttafoco L, Poot AA, Daamen WF, van Kup-pevelt TH, Dijkstra PJ, de Vos RA, Sterk LM, GeelkerkenBR, Feijen J, Vermes L. Tissue engineering of blood vessels:Characterization of smooth-muscle cells for culturing on col-lagen-and-elastin-based scaffolds. Biotechnol Appl Biochem2004;39(Pt 2):141–149.

40. Dreisewerd K, Rohlfing A, Spottke B, Urbanke C, Henkel W.Characterization of whole fibril-forming collagen proteins oftypes I, III, and V from fetal calf skin by infrared matrix-assisted laser desorption ionization mass spectrometry. AnalChem 2004;76:3482–3491.

41. Schonherr E, Witsch-Prehm P, Harrach B, Robenek H, Rau-terberg J, Kresse H. Interaction of biglycan with type I colla-gen. J Biol Chem 1995;270:2776–2783.

42. Eriksen HA, Sharp CA, Robins SP, Sassi ML, Risteli L, Ris-teli J. Differently cross-linked and uncross-linked carboxy-ter-minal telopeptides of type I collagen in human mineralisedbone. Bone 2004;34:720–727.

43. Vorm O, Reopstorff P, Mann M. Improved Resolution andvery high senisitivity in MALDI TOF of matrix surfacesmade by fast evaporation. Anal Chem 1994;66:3281–3287.

44. Gusev A, Wilkinson W, Proctor A, Hercules D. Improvementof signal reproducibility and matrix/comatrix effects inMALDI analysis. Anal Chem 1995;67:1034–1041.

45. Demers LM, Costa L, Lipton A. Biochemical markers andskeletal metastases. Cancer 2000;88(12 Suppl):2919–2926.

46. Van den Steen PE, Proost P, Grillet B, Brand DD, Kang AH,Van Damme J, Opdenakker G. Cleavage of denatured naturalcollagen type II by neutrophil gelatinase B reveals enzymespecificity, post-translational modifications in the substrate,and the formation of remnant epitopes in rheumatoid arthritis.Faseb J 2002;16:379–389.

47. Wei L, Sun XJ, Wang Z, Chen Q. CD95-induced osteoar-thritic chondrocyte apoptosis and necrosis: Dependency onp38 mitogen-activated protein kinase. Arthr Res Ther 2006;8:R37.

48. Bonner WM, Laskey RA. A film detection method for trit-ium-labelled proteins and nucleic acids in polyacrylamidegels. Eur J Biochem 1974;46:83–88.

49. Miles CA, Ghelashvili M. Polymer-in-a-box mechanism forthe thermal stabilization of collagen molecules in fibers. Bio-phys J 1999;76:3243–3252.



50. Ellerbroek SM, Wu YI, Stack MS. Type I collagen stabiliza-tion of matrix metalloproteinase-2. Arch Biochem Biophys2001;390:51–56.

51. Tam EM, Moore TR, Butler GS, Overall CM. Characteriza-tion of the distinct collagen binding, helicase and cleavagemechanisms of matrix metalloproteinase 2 and 14 (gelatinaseA and MT1-MMP): The differential roles of the MMP hemo-pexin c domains and the MMP-2 fibronectin type II modulesin collagen triple helicase activities. J Biol Chem 2004;279:43336–43344.

52. Brandt J, Krogh TN, Jensen CH, Frederiksen JK, Teisner B.Thermal instability of the trimeric structure of the N-terminalpropeptide of human procollagen type I in relation to assaytechnology. Clin Chem 1999;45:47–53.

53. Sykes B, Puddle B, Francis M, Smith R. The estimation oftwo collagens from human dermis by interrupted gel electro-phoresis. Biochem Biophys Res Commun 1976;72:1472–1480.

54. Greenfield NJ. Methods to estimate the conformation of pro-teins and polypeptides from circular dichroism data. AnalBiochem 1996;235:1–10.

55. Consonni R, Zetta L, Longhi R, Toma L, Zanaboni G, TenniR. Conformational analysis and stability of collagen peptidesby CD and by 1H- and 13C-NMR spectroscopies. Biopolymers2000;53:99–111.

56. Mizuno K, Hayashi T, Peyton DH, Bachinger HP. Hydroxyla-tion-induced stabilization of the collagen triple helix. Acetyl-(glycyl-4(R)-hydroxyprolyl-4(R)-hydroxyprolyl)(10)-NH(2)forms a highly stable triple helix. J Biol Chem 2004;279:38072–38078.

57. Stankus JJ, Guan J, Wagner WR. Fabrication of biodegradableelastomeric scaffolds with sub-micron morphologies. J BiomedMater Res A 2004;70:603–614.

58. Hayashi T, Curran-Patel S, Prockop DJ. Thermal stability ofthe triple helix of type I procollagen and collagen. Precautionsfor minimizing ultraviolet damage to proteins during circulardichroism studies. Biochemistry 1979;18:4182–4187.

59. Babu IR, Ganesh KN. Enhanced triple helix stability of colla-gen peptides with 4R-aminoprolyl (Amp) residues: Relativeroles of electrostatic and hydrogen bonding effects. J AmChem Soc 2001;123:2079–2080.

60. Peltonen L, Palotie A, Hayashi T, Prockop DJ. Thermal sta-bility of type I and type III procollagens from normal humanfibroblasts and from a patient with osteogenesis imperfecta.Proc Natl Acad Sci USA 1980;77:162–166.

61. Vomund AN, Braddock SR, Krause GF, Phillips CL. Potentialmodifier role of the R618Q variant of proalpha2(I)collagen intype I collagen fibrillogenesis: In vitro assembly analysis. MolGenet Metab 2004;82:144–153.

62. Leikina E, Mertts MV, Kuznetsova N, Leikin S. Type I colla-gen is thermally unstable at body temperature. Proc NatlAcad Sci USA 2002;99:1314–1318.

63. Wang X, Li X, Bank RA, Agrawal CM. Effects of collagenunwinding and cleavage on the mechanical integrity of thecollagen network in bone. Calcif Tissue Int 2002;71:186–192.

64. Ellerbroek SM, Wu YI, Overall CM, Stack MS. Functionalinterplay between type I collagen and cell surface matrix met-alloproteinase activity. J Biol Chem 2001;276:24833–24842.

65. Day JS, Van Der Linden JC, Bank RA, Ding M, Hvid I,Sumner DR, Weinans H. Adaptation of subchondral bone inosteoarthritis. Biorheology 2004;41:359–368.

66. Xu Y, Keene DR, Bujnicki JM, Hook M, Lukomski S. Strep-tococcal Scl1 and Scl2 proteins form collagen-like triple heli-ces. J Biol Chem 2002;277:27312–27318.

67. Jenness DD, Sprecher C, Johnson WCJ. Circular dichroism ofcollagen, gelatin, and poly(proline) II in the vacuum ultravio-let. Biopolymers 1976;15:513–521.

68. Brown EM, Dudley RB, Elsetinow AR. A conformationalstudy of collagen as affected by tanning procedures. J AmLeather Chem Assoc 1997;92:225–233.

69. Rossi A, Zuccarello LV, Zanaboni G, Monzani E, Dyne KM,Cetta G, Tenni R. Type I collagen CNBr peptides: Speciesand behavior in solution. Biochemistry 1996;35:6048–6057.

70. Yamauchi K, Takeuchi N, Kurimoto A, Tanabe T. Films ofcollagen crosslinked by S��S bonds: Preparation and charac-terization. Biomaterials 2001;22:855–863.

71. Chakrabartty A, Kortemme T, Baldwin RL. Helix propensitiesof the amino acids measured in alanine-based peptides with-out helix-stabilizing side-chain interactions. Protein Sci 1994;3:843–852.

72. Cochran DA, Doig AJ. Effect of the N2 residue on the stabil-ity of the a -helix for all 20 amino acids. Protein Sci 2001;10:1305–1311.

73. Bohm G, Muhr R, Jaenicke R. Quantitative analysis of proteinfar UV circular dichroism spectra by neural networks. ProteinEng 1992;5:191–195.

74. Wilson DL, Martin R, Hong S, Cronin-Golomb M, MirkinCA, Kaplan DL. Surface organization and nanopatterning ofcollagen by dip-pen nanolithography. Proc Natl Acad SciUSA 2001;98:13660–13664.

75. Jelesarov I, Bosshard HR. Isothermal titration calorimetry anddifferential scanning calorimetry as complementary tools toinvestigate the energetics of biomolecular recognition. J MolRecogn 1999;12:3–18.

76. Flandin F, Buffevant C, Herbage D. A differential scanningcalorimetry analysis of the age-related changes in the thermalstability of rat skin collagen. Biochim Biophys Acta 1984;791:205–211.

77. Kumar TR, Shanmugasundaram N, Babu M. Biocompatiblecollagen scaffolds from a human amniotic membrane: Physi-cochemical and in vitro culture characteristics. J Biomater SciPolym Ed 2003;14:689–706.

78. Rochdi A, Foucat L, Renou JP. Effect of thermal denaturationon water–collagen interactions: NMR relaxation and differentialscanning calorimetry analysis. Biopolymers 1999;50:690–696.

79. Knott L, Bailey AJ. Collagen biochemistry of avian bone:Comparison of bone type and skeletal site. Br Poult Sci 1999;40:371–379.

80. Hickman D, Sims TJ, Miles CA, Bailey AJ, de Mari M,Koopmans M. Isinglass/collagen: Denaturation and functional-ity. J Biotechnol 2000;79:245–257.

81. Sionkowska A, Kaminska A. Thermal helix–coil transition inUV irradiated collagen from rat tail tendon. Int J Biol Macro-mol 1999;24:337–340.

82. Kubisz L, Mielcarek S, Jaroszyk F. Changes in thermal andelectrical properties of bone as a result of 1 MGy-dose c-irra-diation. Int J Biol Macromol 2003;33:89–93.

83. Hanley L, Kornienko O, Ada ET, Fuoco E, Trevor JL. Sur-face mass spectrometry of molecular species. J Mass Spec-trom 1999;34:705–723.

84. Merrett K, Cornelius RM, McClung WG, Unsworth LD, Shear-down H. Surface analysis methods for characterizing polymericbiomaterials. J Biomater Sci Polym Ed 2002;13:593–621.

85. Dewez JL, Doren A, Schneider YJ, Rouxhet PG. Competitiveadsorption of proteins: Key of the relationship between sub-stratum surface properties and adhesion of epithelial cells.Biomaterials 1999;20:547–559.

86. Dewez JL, Schneider YJ, Rouxhet PG. Coupled influence ofsubstratum hydrophilicity and surfactant on epithelial cell ad-hesion. J Biomed Mater Res 1996;30:373–383.

87. Zambonin G, Losito I, Triffitt JT, Zambonin CG. Detection ofcollagen synthesis by human osteoblasts on a tricalcium phos-phate hydroxyapatite: An X-ray photoelectron spectroscopyinvestigation. J Biomed Mater Res 2000;49:120–126.

284 ABRAHAM ET AL.


88. Healy KE, Ducheyne P. Hydration and preferential molecularadsorption on titanium in vitro. Biomaterials 1992;13:553–561.

89. Morra M, Cassinelli C, Cascardo G, Cahalan P, Cahalan L,Fini M, Giardino R. Surface engineering of titanium by colla-gen immobilization. Surface characterization and in vitro andin vivo studies. Biomaterials 2003;24:4639–4654.

90. Pamula E, De Cupere V, Dufrene YF, Rouxhet PG. Nanoscaleorganization of adsorbed collagen: Influence of substratehydrophobicity and adsorption time. J Colloid Interface Sci2004;271:80–91.

91. Cheng Z, Teoh SH. Surface modification of ultra thin poly(epsilon-caprolactone) films using acrylic acid and collagen.Biomaterials 2004;25:1991–2001.

92. Kwok DY, Gietzelt T, Grundke K, Jacobasch H-J, NeumannAW. Contact angle measurements and contact angle interpre-tation, Part 1: Contact angle measurements by axisymmetricdrop shape analysis and a goniometer sessile drop technique.Langmuir 1997;13:2880–2894.

93. Altankov G, Grinnell F, Groth T. Studies on the biocompati-bility of materials: Fibroblast reorganization of substratum-bound fibronectin on surfaces varying in wettability. J BiomedMater Res 1996;30:385–391.

94. Brocchini S, James K, Tangpasuthadol V, Kohn J. Structure–property correlations in a combinatorial library of degradablebiomaterials. J Biomed Mater Res 1998;42:66–75.

95. Tamada Y, Ikada Y. Fibroblast growth on polymer surfaces andbiosynthesis of collagen. J BiomedMater Res 1994;28:783–789.

96. Higuchi A, Tamiya S, Tsubomura T, Katoh A, Cho CS,Akaike T, Hara M. Growth of L929 cells on polymeric filmsprepared by Langmuir-Blodgett and casting methods. J Bio-mater Sci Polym Ed 2000;11:149–168.

97. Ber S, Torun Kose G, Hasirci V. Bone tissue engineering onpatterned collagen films: An in vitro study. Biomaterials 2005;26:1977–1986.

98. Paige MF, Rainey JK, Goh MC. A study of fibrous long spac-ing collagen ultrastructure and assembly by atomic force mi-croscopy. Micron 2001;32:341–353.

99. Hassenkam T, Fantner GE, Cutroni JA, Weaver JC, MorseDE, Hansma PK. High-resolution AFM imaging of intact andfractured trabecular bone. Bone 2004;35:4–10.

100. Zhang J, Senger B, Vautier D, Picart C, Schaaf P, Voegel JC,Lavalle P. Natural polyelectrolyte films based on layer-bylayer deposition of collagen and hyaluronic acid. Biomaterials2005;26:3353–3361.

101. Habelitz S, Balooch M, Marshall SJ, Balooch G, Marshall GW Jr.In situ atomic force microscopy of partially demineralized humandentin collagen fibrils. J Struct Biol 2002;138:227–236.

102. Jiang F, Khairy K, Poole K, Howard J, Muller DJ. Creatingnanoscopic collagen matrices using atomic force microscopy.Microsc Res Tech 2004;64:435–440.

103. Cristofalo VJ, Pignolo RJ. Replicative senescence of humanfibroblast-like cells in culture. Physiol Rev 1993;73:617–638.

104. Qiu Q, Sayer M, Kawaja M, Shen X, Davies JE. Attachment,morphology, and protein expression of rat marrow stromalcells cultured on charged substrate surfaces. J Biomed MaterRes 1998;42:117–127.

105. Ma L, Gao C, Mao Z, Zhou J, Shen J. Enhanced biologicalstability of collagen porous scaffolds by using amino acids asnovel cross-linking bridges. Biomaterials 2004;25:2997–3004.

106. Chen J, Altman GH, Karageorgiou V, Horan R, Collette A,Volloch V, Colabro T, Kaplan DL. Human bone marrow stro-mal cell and ligament fibroblast responses on RGD-modifiedsilk fibers. J Biomed Mater Res A 2003;67:559–570.

107. Tan W, Krishnaraj R, Desai TA. Evaluation of nanostructuredcomposite collagen–chitosan matrices for tissue engineering.Tissue Eng 2001;7:203–210.

108. Tampieri A, Celotti G, Landi E, Sandri M, Roveri N, FaliniG. Biologically inspired synthesis of bone-like composite:Self-assembled collagen fibers/hydroxyapatite nanocrystals.J Biomed Mater Res A 2003;67:618–625.

109. Detamore MS, Orfanos JG, Almarza AJ, French MM, WongME, Athanasiou KA. Quantitative analysis and comparative re-gional investigation of the extracellular matrix of the porcinetemporomandibular joint disc. Matrix Biol 2005;24:45–57.

110. Dunn CA, Jin Q, Taba M Jr, Franceschi RT, Bruce RutherfordR, Giannobile WV. BMP gene delivery for alveolar bone engi-neering at dental implant defects. Mol Ther 2005;11:294–299.

111. Baguneid M, Murray D, Salacinski HJ, Fuller B, Hamilton G,Walker M, Seifalian AM. Shear stress preconditioning and tis-sue engineering based paradigms for generating arterial sub-stitutes. Biotechnol Appl Biochem 2004;39:151–157.

112. Ferrando JM, Vidal J, Armengol M, Huguet P, Gil J, ManeroJM, Planell JA, Segarra A, Schwartz S, Arbos MA. Earlyimaging of integration response to polypropylene mesh in ab-dominal wall by environmental scanning electron microscopy:Comparison of two placement techniques and correlation withtensiometric studies. World J Surg 2001;25:840–847.

113. Ehlers EM, Fuss M, Rohwedel J, Russlies M, Kuhnel W,Behrens P. Development of a biocomposite to fill out articularcartilage lesions. Light, scanning and transmission electronmicroscopy of sheep chondrocytes cultured on a collagen I/IIIsponge. Anat Anz 1999;181:513–518.

114. Hsu SH, Tsai CL, Tang CM. Evaluation of cellular affinityand compatibility to biodegradable polyesters and type-II col-lagen-modified scaffolds using immortalized rat chondrocytes.Artif Organs 2002;26:647–658.

115. Cherubino P, Grassi FA, Bulgheroni P, Ronga M. Autologouschondrocyte implantation using a bilayer collagen membrane: Apreliminary report. J Orthop Surg (Hong Kong) 2003;11:10–15.

116. Mauney JR, Kaplan DL, Volloch V. Matrix-mediated reten-tion of osteogenic differentiation potential by human adultbone marrow stromal cells during ex vivo expansion. Bioma-terials 2004;25:3233–3243.

117. Garfinkel S, Wessendorf JH, Hu X, Maciag T. The human dip-loid fibroblast senescence pathway is independent of interleu-kin-1 a mRNA levels and tyrosine phosphorylation of FGFR-1substrates. Biochim Biophys Acta 1996;1314:109–119.

118. Chen KY, Chang ZF. Age dependency of the metabolic conver-sion of polyamines into amino acids in IMR-90 human embry-onic lung diploid fibroblasts. J Cell Physiol 1986;128:27–32.

119. Low RB, Hildebran JN, Absher PM, Stirewalt WS, Arnold J.Comparison of the use of isotopic proline vs. leucine to mea-sure protein synthesis in cultured fibroblasts. Connect TissueRes 1986;14:179–185.

120. Hildebran JN, Airhart J, Stirewalt WS, Low RB. Prolyl-tRNA-based rates of protein and collagen synthesis in humanlung fibroblasts. Biochem J 1981;198:249–258.

121. Kulju KS, Lehman JM. Increased p53 protein associated withaging in human diploid fibroblasts. Exp Cell Res 1995;217:336–345.

122. Mu XC, Higgins PJ. Differential growth state-dependent regu-lation of plasminogen activator inhibitor type-1 expression insenescent IMR-90 human diploid fibroblasts. J Cell Physiol1995;165:647–657.

123. Briggs D. Surface Analysis of Polymers by XPS and StaticSIMS. Cambridge, UK: Cambridge University Press; 1998.

124. Patankar NA. Mimicking the lotus effect: Influence of doubleroughness structures and slender pillars. Langmuir 2004;20:8209–8213.

125. Cristofalo VJ. Cellular biomarkers of aging. Exp Gerontol1988;23:297–307.



Abraham-2008-Guide to collagen ch

Documents

Transcript of Abraham-2008-Guide to collagen ch