friction and wear measurments of bovine articular

FRICTION AND WEAR MEASURMENTS OF BOVINE ARTICULAR CARTILAGE AGAINST NON-NATIVE MATERIALS

Sevan R. Oungoulian

Submitted in partial fulfillment of the

requirements for the degree

of Doctor of Philosophy

in the Graduate School of Arts and Sciences

COLUMBIA UNIVERSITY

2015

©2015

Sevan R. Oungoulian

All rights reserved

ABSTRACT

FRICTION AND WEAR MEASURMENTS OF BOVINE ARTICULAR CARTILAGE AGAINST NON-

NATIVE MATERIALS

Sevan R. Oungoulian

The three studies reported in this thesis investigate the friction and wear properties of articular cartilage.

The aim of the first study presented was to examine the functional properties and biocompatibility

of glutaraldehyde-fixed bovine articular cartilage for potential use as a resurfacing material in joint

arthroplasty. Treated cartilage disks were fixed over a range of glutaraldehyde concentrations and

incubated, along with an untreated control group, at 37 °C for up to 28 days. The equilibrium

compressive modulus increased nearly twofold in the treated samples when compared to day 0 control,

and maintained that property value from day 1 to day 28; the minimum friction coefficient did not change

significantly with fixation and incubation time, whereas the time constant for the frictional response

decreased twofold at most. Live explants co-cultured with fixed explants showed no qualitative difference

in cell viability over 28 days of incubation. Cartilage-on-cartilage frictional measurements were performed

under the configuration of migrating contact for a subset of treated explants over a period of 28 days

exhibited either no significant difference or slightly lower friction coefficient values than the untreated

control group. These results suggest that a properly titrated glutaraldehyde treatment can reproduce the

desired functional properties of native articular cartilage and maintain these properties for at least 28 days

in-vitro.

The aim of the second study was to determine whether the latest-generation particle analyzers

are capable of detecting cartilage wear debris generated during in vitro loading experiments that last 24 h

or less, by producing measurable content significantly above background noise levels otherwise

undetectable through standard biochemical assays. Immature bovine cartilage disks were tested against

glass using reciprocal sliding under unconfined compression creep for 24 h. Control groups were used to

assess various sources of contamination. Results demonstrated that cartilage samples subjected to

frictional loading produced particulate volume significantly higher than background noise and

contamination levels at all tested time points. The particle counter used was able to detect very small

levels of wear), whereas no significant differences were observed in biochemical assays for collagen or

glycosaminoglycans among any of the groups or time points.

The aim of the final study was to measure the wear response of immature bovine articular

cartilage tested against glass or alloys used in hemiarthroplasties. Two cobalt chromium alloys and a

stainless steel alloy were selected for these investigations. The surface roughness of one of the cobalt

chromium alloys was also varied within the range considered acceptable by regulatory agencies.

Cartilage disks were tested in a configuration that promoted loss of interstitial fluid pressurization to

replicate conditions believed to occur in hemiarthroplasties. Results showed that considerably more

damage occurred in cartilage samples tested against smooth stainless steel and rough cobalt chromium

alloys compared to smooth glass, and smooth cobalt chromium alloys. The two materials producing the

greatest damage also exhibited higher equilibrium friction coefficients. Cartilage damage occurred

primarily in the form of delamination at the interface between the superficial tangential zone and the

transitional middle zone, with much less evidence of abrasive wear at the articular surface. These results

suggest that cartilage damage from frictional loading occurs as a result of subsurface fatigue failure

leading to the delamination. Surface chemistry and surface roughness of implant materials can have a

significant influence on tissue damage, even when using materials and roughness values that satisfy

regulatory requirements.

i

TABLE OF CONTENTS

LIST OF FIGURES ....................................................................................................................................... iv

LIST OF TABLES .......................................................................................................................................... v

Chapter 1: INTRODUCTION ................................................................................................................... 1

1.1 Cartilage Lubrication ..................................................................................................................... 1

1.1.1 Boundary Lubrication ................................................................................................................ 1

1.1.2 Interstitial Fluid Pressurization .................................................................................................. 1

1.2 Cartilage Wear .............................................................................................................................. 2

1.3 Hemiarthroplasty ........................................................................................................................... 2

Chapter 2: GLUTARALDEHYDE FIXED CARTILAGE AS A RESURFACING MATERIAL ................... 4

2.1 Introduction .................................................................................................................................... 4

2.2 Materials and Methods .................................................................................................................. 6

2.2.1 Study Design ............................................................................................................................. 6

2.2.2 Sample Harvest ......................................................................................................................... 6

2.2.1 Glutaraldehyde Treatment ........................................................................................................ 7

2.2.2 Mechanical Testing ................................................................................................................... 8

2.2.3 Biocompatibility Testing ............................................................................................................ 9

2.3 Results ........................................................................................................................................ 10

2.3.1 Study 2-1 ................................................................................................................................. 10

2.3.2 Study 2-2 ................................................................................................................................. 11

2.4 Discussion ................................................................................................................................... 11

2.5 Acknowledgements ..................................................................................................................... 13

2.6 Figures ........................................................................................................................................ 14

ii

Chapter 3: CARTILAGE WEAR DEBRIS MEASUREMENTS VIA PARTICLE ANALYZER ................ 23

3.1 Introduction .................................................................................................................................. 23

3.2 Materials and Methods ................................................................................................................ 23

3.2.1 Sample Harvest ....................................................................................................................... 23

3.2.2 Solution Preparation ................................................................................................................ 23

3.2.3 Mechanical Testing ................................................................................................................. 24

3.2.4 Particulate Analyzer ................................................................................................................ 24

3.2.5 Particulate Assay..................................................................................................................... 25

3.2.6 Particulate Imaging.................................................................................................................. 25

3.2.7 Statistical Analysis................................................................................................................... 26

3.1 Results ........................................................................................................................................ 26

3.1.1 Particulate Number.................................................................................................................. 26

3.1.2 Particulate Volume .................................................................................................................. 26

3.1.3 Particulate Assay and Imaging ................................................................................................ 26

3.2 Discussion ................................................................................................................................... 26


3.5 Figures ........................................................................................................................................ 29

Chapter 4: INFLUENCE OF BEARING MATERIAL AND ROUGHNESS ON CARTILAGE

DEGENERATION ....................................................................................................................................... 34

4.1 Introduction .................................................................................................................................. 34

4.2 Materials and Methods ................................................................................................................ 35

4.2.1 Material Preparation ................................................................................................................ 35

4.2.1 Sample Harvest ....................................................................................................................... 36

iii


4.2.3 Particulate Testing................................................................................................................... 37

4.2.4 Biochemical Testing ................................................................................................................ 38

4.2.5 Histological Staining ................................................................................................................ 38

4.3 Results ........................................................................................................................................ 38

4.3.1 Material Roughness Testing ................................................................................................... 38


4.3.3 Particulate Testing................................................................................................................... 39

4.3.4 Biochemical Testing ................................................................................................................ 39

4.3.5 Histological Staining ................................................................................................................ 39

4.4 Discussion ................................................................................................................................... 39


4.6 Figures ........................................................................................................................................ 41

Chapter 5: CONCLUSION .................................................................................................................... 48

5.1 Motivation .................................................................................................................................... 48

5.2 Glutaraldehyde Fixed Cartilage As A Resurfacing Material ........................................................ 48

5.3 Cartilage Wear Debris Measurements Via Particle Analyzer ..................................................... 49

5.4 Influence of Artificial Bearing Material on Cartilage Degeneration ............................................. 49

Chapter 6: REFERENCES .................................................................................................................... 51

iv

LIST OF FIGURES

Figure 2-1: Schematic representation of bovine calf articular cartilage harvest location ........................... 14

Figure 2-2: Representative biocompatibility culture dish with viable and glutaraldehyde treated cartilage 15

Figure 2-3: Representative FEBio curve-fit of experimental stress-relaxation test ..................................... 16

Figure 2-4: Summary of mechanical properties for Study 2-1 .................................................................... 17

Figure 2-5: Qualitative and quantitative biocompatibility from chondrocyte viability ................................... 18

Figure 2-6: Frictional properties for Study 2-2 ............................................................................................ 19

Figure 2-7: India ink staining of representative Study 2-2 tibial plateau strips ........................................... 20

Figure 2-8: Intact immature bovine knee condyles ..................................................................................... 21

Figure 2-9: Potential clinical approach for bioprosthetic arthroplasties ...................................................... 22

Figure 3-1: Schematic for TEST, CTRL, ENVR (cross-sectional views), and BASE .................................. 30

Figure 3-2: Wear particulate measurements for bovine articular cartilage explants................................... 31

Figure 3-3: Wear particulate confocal z stack and angled z stack.............................................................. 32

Figure 3-4: Particulate size (A) and volume (B) distribution for representative 24 h TEST solution .......... 33

Figure 4-1: Schematic representation of bovine calf articular cartilage harvest location ........................... 41

Figure 4-2: Frictional properties of cartilage against different counterface materials ................................. 42

Figure 4-3: Representative qualitative macroscopic comparison of cartilage tissue .................................. 43

Figure 4-4: Summary of cartilage mechanical properties and wear measurements .................................. 44

Figure 4-5: Summary of cartilage biochemical properties .......................................................................... 45

Figure 4-6: Representative histological images of top region ..................................................................... 46

v

LIST OF TABLES

Table 3-1: Biochemical assay measurements of cartilage wear debris ...................................................... 29

Table 4-1: Roughness measurements for orthopaedic material platens .................................................... 47

vi

ACKNOWLEDGEMENTS

My graduate experience here at Columbia University has been extraordinarily rewarding and fulfilling.

There are many people who have contributed to and helped shape this experience.

I would like to express gratitude to Dr. Gerard Ateshian. To have worked under his guidance has

been a great privilege. His unwavering sense of academic rigor and discipline set a high bar that I have

strived to reach in my graduate career. His kindness, patience, and encouragement along the way

provided a model of managerial action. The persistence of his intellectual curiosity coupled with his desire

for deep understanding is quite extraordinary. Of much personal importance, I would like to thank him for

his trust in the exploration of new scientific ideas.

I would like to thank members of my doctoral committee for their valuable insights and comments

to this work: Dr. Clark Hung, Dr. Kristin Myers, Dr. Christopher Ahmad, and Dr. Jeffrey Kysar. In

particular, I would like to thank Dr. Clark Hung for his discussions and insight to tissue and biochemical

interactions at the cellular level. I have thoroughly enjoyed our discussions and know that this body of

research would not be complete without your valuable advice.

Dr. Michael Albro was an outstanding role model first as a graduate student and eventually as a

post-doctoral researcher at Columbia. Mike has been both a source of tremendous help, friendship, and

inspiration. I feel very fortunate for having been able to work with him.

I would like to thank my close friends and current colleagues, Brian Jones, Robert Nims, Alex

Cigan, Krista Durney, Chieh Hou, and Brandon Zimmerman. I will deeply miss working beside you and

wish you all the very best. I would also like to thank Vikram Rajan, Dr. Matteo Caligaris, Dr. Clare Canal,

Dr. Hui Chen, Charlie Yongpravat, Dr. Elizabeth Oswald, Andrea Tan, Sonal Sampat, Adam Nover,

Brendan Roach, Amy Silverstein, and Kyoko Yoshida, it has been a pleasure knowing and working with

all of you. Thanks to Keith Yeager for providing a great collaborative partner in the area of engineering

design, for granting me wide use of facilities, and for his support and access to valuable professional

opportunities. I have also been fortunate in having worked with tremendous undergraduate and graduate

student research volunteers who have taught me as much about being a mentor as I hope to have taught

them about rubbing together bits of cartilage. Thank you for making this experience at Columbia

memorable.

vii

My friends and family deserve so very much appreciation for all of their continued love and

support: to my parents Semik and Laura Oungoulian for their unyielding encouragement for success in all

of my pursuits, to Ani and Shaunt Oungoulian for their much appreciated sense of humor, and to Sarah

Dymkar Brandt, for her incredible warmth, love, and care.

1

CHAPTER 1: INTRODUCTION

1.1 Cartilage Lubrication

1.1.1 Boundary Lubrication

The primary function of articular cartilage is to serve as the bearing material in diarthrodial joints,

transmitting loads while minimizing friction and wear. The friction coefficient of cartilage has been

characterized extensively in the literature, using standard measurements of normal and tangential forces

acting across a sliding interface [1-8]. In the mid 20th century, the idea that cartilage tribology was

mediated by the mechanism of boundary lubrication was proposed by Sir John Charnley [9]. Boundary

lubricants are typically thought of as molecules that coat the surface of a bearing and effect a low friction

coefficient due to molecular-level repulsive forces [10]. Many investigators since have explored the

tribological function of different synovial fluid constituents, such as hyaluronan [11-17], lubricin [7, 8, 18-

22], and surface-active phospholipids [13, 14, 23-29]. Homology was discovered between lubricin [30, 31]

and superficial zone protein, a more recently identified molecule that is expressed by chondrocytes in the

superficial region of the tissue [23, 32]. This discovery suggests that cartilage constituents may also

contribute to boundary lubrication.

1.1.2 Interstitial Fluid Pressurization

The concept that pressurization of the interstitial fluid of articular cartilage could play a dominant role in

cartilage lubrication has been the subject of more recent theoretical and experimental work. Analysis has

helped formulate a theoretical framework that would embody this mechanism for friction coefficient

reduction. It has been proposed that the joint articular surfaces come in direct contact, but that the

pressurized interstitial fluid of this highly porous tissue supports most of the contact load. Cartilage

porosity is especially high at the articular surface, so the contact area over which collagen are in direct

contact with apposing collagen fibers is very low [33]; with load transfer between collagen and fluid, or

fluid and fluid, occurring over the bulk of the remaining contact area. This results in a very low friction

coefficient while the interstitial fluid pressure is elevated. The friction coefficient will rise as the portion of

2

the load supported by the pressurized interstitial fluid diminishes, and as the portion of the load supported

by direct contact between solid constituents increases. Agreement between theoretical predictions and

and experiment, much of which was performed previously in this lab, is very strong [33-38].

Important insight into the reason for low in-vivo friction can be gained from the investigation of this

interstitial fluid pressurization mechanism. It has since been shown that under conditions of steady motion

between curved articular cartilage surfaces, that interstitial fluid pressurization remains elevated [39].

Under conditions that promote a continuously migrating contact area, the friction coefficient remains low

[39]. This finding is significant when considered in the context that during normal function of diarthrodial

joints, conditions exist that promote migrating contact areas on at least one of the apposing articular

surfaces. This has been demonstrated in experimental studies of articular contact during normal activities

of daily living in many different diarthrodial joints [40-44].

1.2 Cartilage Wear

As mentioned previously, the friction coefficient of cartilage has been characterized extensively in the

literature [1-8]. However, characterization of cartilage wear and damage has proved to be more difficult,

and a standard method for measuring microscopic wear has yet to be agreed upon. Qualitative

observations of cartilage wear debris have been made [45-52]; however, fewer studies report quantitative

measurements of cartilage wear. The primary quantitative approaches proposed to date include

biochemical assaying of cartilage and test solutions [53-57], characterization of changing articular layer

thickness [54, 58-60], and changes in surface roughness [7, 57, 61-65]. Each of these methods has

limitations in determining the minute amounts of wear expected in the normal in-vivo environment.

Without a standard method for quantitative measurement, the complex relationship between tissue

damage and wear and friction coefficient is still poorly understood, and determining the underlying

physical and biological events that lead to these conditions remains elusive [3, 66-69]. It is generally

assumed that low friction is associated with low wear and damage, but as seen with the first generation of

artificial hips, this association is not straightforward [70-72].

1.3 Hemiarthroplasty

3

The surgical procedure in which half of a damaged synovial joint is replaced with a non-native bearing is

called hemiarthroplasty. According to the Centers for Disease Control, over 360,000 hemiarthroplasty

procedures are performed each year in the US, [73]. Hemiarthroplasties are less invasive, require less

surgical time and result in lower blood loss than total arthroplasties [74, 75], however the value over total

joint arthroplasty still subject to debate [74-82]. Some indications for hemiarthroplasty include fracture of

the femoral or humeral head [83-85] and fixation failure due to osteonecrosis [76, 86, 87], while

contraindications include disease or damage to the apposing joint surface [80]. Hemiarthroplasty patient

outcomes are sometimes less successful than total arthroplasties, and often require revision, due in part

to accelerated erosion of the articular surface [59, 88-92] although reasons for this acceleration in

damage is unclear.

Extending the insight gained by our understanding of the importance of migrating contact areas on

maintaining an elevated interstitial fluid pressure in articular cartilage, and resulting low friction coefficient,

to joint hemiarthroplasties might perhaps help explain this accelerated wear. It is possible that traditional

hemiarthroplasties compromise joint lubrication by reducing the migrating contact pattern.

4

CHAPTER 2: GLUTARALDEHYDE FIXED CARTILAGE AS A

RESURFACING MATERIAL

2.1 Introduction

Total joint arthroplasty is a commonly applied and generally effective surgical treatment for debilitating

arthritic conditions, accounting for over 1 million surgical procedures in the US alone during 2010

according to the CDC [73]. The development of an alternative resurfacing material to impermeable metals

or ultra-high molecular weight polyethylene remains an important engineering achievement that can

potentially improve the long-term clinical outcome of arthroplasty procedures by providing a more

cartilage-like bioprosthetic replacement [93, 94]. Recent theoretical developments and corroborating

experimental evidence support the idea that interstitial fluid pressurization of the porous native cartilage is

critical to the maintenance of low friction [38, 95, 96]. These findings suggest that joint resurfacing with a

porous biologically stable material has the potential to outperform traditional nonporous implant materials,

because of improved tribological properties. Though porous hydrogels and electrospun polymers have

been considered [63, 97-101], chemically fixed cartilage allografts or xenografts may exhibit better

functional properties [102-106].

Bioprosthetic heart valves have been successfully used clinically since the 1960s [107-109].

These valves typically consist of human or porcine heart valve, or bovine pericardial tissue that has been

chemically treated. Glutaraldehyde is most commonly used for fixation and sterilization, though it is

sometimes supplemented with alcohol or gamma radiation (when using low glutaraldehyde

concentrations) [110, 111]. Glutaraldehyde causes cross-linking of proteins in the valve tissue

extracellular matrix, which has been shown to alter mechanical properties [110, 112-115], though fixed

valves are able to maintain a satisfactory level of function when implanted, as evidenced from their

clinical success. Fixation under a pressure differential has also been advocated, in order to alter both the

configuration under which cross-linking occurs and the resulting stress-strain response of the fixed tissue

[113, 116-118].

5

Similar treatment has been advocated for musculoskeletal connective tissues [119-122], most

notably for the knee meniscus [103-105, 123-128]. These studies have similarly demonstrated that

glutaraldehyde treatment may alter mechanical properties [103, 128], though proper titration of the

concentration may control this variation [103, 104]. Animal studies have been performed which use whole

menisci as bioprosthetic replacements to the native meniscus [123, 125, 126] or as osteochondral

allografts [106, 124, 127], showing encouraging biomechanical function and integration. An additional

human case study was reported to have successfully used a glutaraldehyde fixed bovine meniscus

xenograft for the treatment of avascular necrosis of the scaphoid proximal pole, with satisfactory

performance reported after five years without the occurrence of inflammation [127]. Fewer studies have

been reported on the glutaraldehyde fixation of articular cartilage: One study demonstrated that fixation

increased the resistance of the tissue to degradation by collagenase [129], and another study

demonstrated decreased wear and improved tissue stabilization in vitro, following fixation [130].

Previous basic science findings have clearly established the structure-function relationships of

articular cartilage that promote elevated interstitial fluid pressurization: The tissue must be highly porous

but exhibit low permeability, to prevent rapid loss of interstitial fluid pressurization [35, 36, 131]; it must

exhibit a higher modulus in tension than compression in order to enhance this pressurization, by

restricting the ability of the tissue to expand laterally when compressed axially [34, 131-133]. In particular,

it is known from theory [134] and experiments [39] that elevated interstitial fluid pressurization and

decreased friction can be achieved when the dimensionless Peclet number for cartilage sliding contact is

much greater than unity (Pe>>1). The Peclet number is given by Pe=Vh/HAk, where V is the velocity of

the migrating contact area, h is a characteristic dimension of the contacting articular layers (cartilage

thickness or contact radius), HA is its aggregate modulus and k its hydraulic permeability. It is also known

that interstitial fluid pressurization is enhanced when the tensile modulus of the porous material is much

greater than its compressive modulus [34, 35, 131, 135]. Finally, it is known that the friction coefficient

decreases with increasing porosity [33, 38].

Therefore, though glutaraldehyde treatment is known to alter the mechanical and transport

properties of soft tissues, these alterations may be titrated by adjusting the glutaraldehyde concentration

[103, 104] to control the alteration in Pe such that it remains significantly above unity. In this case,

6

interstitial fluid pressurization can remain elevated. Many alternative materials, such as hydrogels, do not

share all the critical structure-function relationships found in articular cartilage, which are needed to

promote elevated interstitial fluid pressurization as outlined above. Furthermore, methods to improve their

functionality using alternative hydrogel formulations and fiber reinforcement may be constrained by

biocompatibility issues.

The aim of this study was to examine the functional properties and biocompatibility of glutaraldehyde-

fixed bovine articular cartilage over several weeks of incubation at body temperature to investigate its

potential use as a resurfacing material in joint arthroplasty. The hypothesis was that successful fixation

treatments may preserve functional properties of native tissue over long-term culture at body

temperature.

2.2 Materials and Methods

2.2.1 Study Design

Two studies were performed, each with glutaraldehyde treatment and incubation time as factors. In Study

2.1, the effects of glutaraldehyde stabilization on mechanical properties and tissue biocompatibility were

examined over a period of 28 days. Compressive and tensile moduli, hydraulic permeability, friction

parameters, and biocompatibility (via cell viability) were compared over time (1, 7, 14, or 28 days) to

optimize glutaraldehyde treatment concentration (0% control, 0.02, 0.20, or 0.60% glutaraldehyde) for

tissue maintenance. In Study 2-2, findings from Study 2-21 were applied and additional mechanical and

qualitative tests were performed to evaluate functional performance over time (1, 7, 14, or 28 days) at the

optimal glutaraldehyde treatment concentration (0% control and 0.20% glutaraldehyde) with more

physiologically relevant friction test conditions.

2.2.2 Sample Harvest

Experiments were conducted using articular cartilage explants harvested from matched left-right pairs of

healthy tibial plateaus and opposing condyles of 2-3 months old calf knee joints obtained from a local

abattoir. In Study 2-1, full thickness cartilage discs (4 mm) were excised from the tibial plateaus of eight

joints (Figure 2-1A). Samples were randomized to friction testing, mechanical testing, and biocompatibility

7

groups. Those samples in the friction and mechanical testing groups were placed with the articular

surface down on a freezing stage microtome (Leica Instruments #SM2400, Nusslock, Germany),

embedded and frozen in a water soluble sectioning gel (Thermo Scientific #1310APD, Rockford, IL), then

trimmed of bone and deep zone tissue to obtain samples inclusive of an intact superficial zone with a final

thickness of 1.1 ± 0.03 mm. These testing samples were then randomized into various treatment groups

and test type. Samples were rinsed with and transferred to filtered phosphate buffered saline (PBS)

supplemented with an isothiazolone based biocide (Sigma-Aldrich #46878-U, St. Louis, MO). Viable

explants for biocompatibility testing were manually trimmed of bone and deep zone tissue then

transferred to dishes containing Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10%

fetal bovine, amino acids (1 X minimum essential amino acids, 1 X nonessential amino acids), buffering

agents (10 mM HEPES, 10 mM sodium bicarbonate, 10 mM TES, 10 mM BES), and antibiotics (100

U/mL penicillin, 100 µg/mL streptomycin) and cultured at 37° C and 15% CO2 (Sigma Aldrich, St. Louis,

MO).

In Study 2-2, full thickness cartilage strip-shaped explants (28 mm 10 mm ~1 mm), one from

each medial and lateral sides of the tibial plateau, were harvested along with the opposing femoral

condyles from nine joints (Figure 2-1B). Tibial plateau strips were trimmed of bone and deep zone tissue

as described in Study 2-1 to obtain samples inclusive of an intact superficial zone and with a final

thickness of 1.1 ± 0.06 mm. Full thickness cartilage discs (10 mm) were punched from the condyles for

use as opposing articulating faces. Samples were rinsed with and transferred to filtered PBS with biocide

in the same manner as listed for Study 2-1. Condyle discs were randomized within each joint, wrapped in

PBS soaked gauze and stored at -20 °C for a period no greater than 30 days until use.

2.2.1 Glutaraldehyde Treatment

All glutaraldehyde solutions were prepared from 8% stock glutaraldehyde (Sigma Aldrich #G7526, St.

Louis, MO) and PBS. In Study 2-1, specimens were randomized to four glutaraldehyde treatment groups

(0% control, 0.02, 0.20, or 0.60% glutaraldehyde, n=20 per group per time point, of which 8 were used for

friction testing, 8 for mechanical testing, and 4 for biocompatibility) and placed in 1000 mL (80 samples

per solution concentration) of the appropriate glutaraldehyde supplemented PBS solution for 24 h at 8 °C

8

[103] . After treatment, samples were rinsed with and transferred to 1000 mL sterile PBS with biocide for

a 48 h desorption, rinsed again, and transferred to fresh PBS with biocide where they remained at 37°C

for the remainder of the incubation time (1, 7, 14, or 28 days). Sample groups were rinsed and stored

separately to prevent potential cross contamination. For this study, a total of 340 specimens were tested

(20 at day 0 and 80 at each of days 1, 7, 14 and 28). In Study 2-2, tibial plateau strips were randomized

to two treatment groups (0% control, or 0.20% glutaraldehyde, n = 9 per group, for a total of 18

specimens) and treated with glutaraldehyde according to the protocol listed for Study 2-1 and transferred

to fresh PBS with biocide where they remained at 37°C for the remainder of the incubation time (with

friction tests repeated on the same samples at 1, 7, 14, and 28 days). Samples were randomized by tibial

plateau side, with either treatment or control assigned to either the medial or lateral side for each joint.

2.2.2 Mechanical Testing

A custom two-axis loading device equipped with a six-degrees-of-freedom load cell (JR3 Inc. #20E12A4,

Woodland, CA) was used for all mechanical testing. The device was controlled with custom LabVIEW

software (National Instruments Corporation #LabVIEW 2010, Austin, TX) and an associated motion

controller (National Instruments Corporation #7354, Austin, TX) to provide continuous measures of

specimen thickness, position, applied loads, and reaction loads.

In Study 2-1, unconfined compression stress-relaxation tests were performed on a subset of

samples (n=8 per group from each incubation time point) to evaluate the equilibrium compressive (E-Y)

and tensile moduli (E+Y) and hydraulic permeability (k). For each sample, a tare strain was applied (10%

strain with 4800 s relaxation time). After equilibrium was reached, a stress relaxation test was performed

(to 15% strain from pre-tare reference configuration, with 2400 s relaxation time). Stress-relaxation data

were curve-fitted as previously described [136]. Briefly, finite element models were constructed in FEBio

[137] that incorporated the geometry of the specimens. The cartilage was modeled as a biphasic material

containing intrinsically incompressible fluid and solid phases [138]; the composite porous solid phase

consisted of a compressible neo-Hookean solid ground matrix reinforced by a continuous, random

distribution of fibril bundles sustaining tension only, with a linear stress–strain response; the hydraulic

permeability was assumed constant [136, 139]. A least-square parameter optimization routine in FEBio

9

was used to curve-fit the stress-relaxation data and extract the specimen E-Y, k, and (fibril tensile

modulus) values. These values were substituted into a finite element analysis of equilibrium uniaxial

tension to produce the tissue tensile modulus E+Y [136].

Friction measurements against smooth glass (Edmund Optics Inc. #27-834, Barrington, NJ) were

performed on another subset of samples (n=8), using a creep loading configuration (4.45 N) with

reciprocal sliding motion (±5.0 mm at 1 mm/s, Figure 2-1A) [38]. The temporal response of the friction

coefficient was evaluated from the normal and frictional load responses over 3600 s. The minimum

friction coefficient, µmin, equilibrium friction coefficient, µeq, and time constant to reach equilibrium, tµ, were

calculated as described previously [38]. Samples were submerged in PBS throughout all tests and

allowed to recover for a duration equal to or greater than the test time. A two-way ANOVA on the effects

of treatment and incubation time, followed by post-hoc testing using Tukey correction, was performed on

the moduli, hydraulic permeability, and friction parameters.

In Study 2-2, quantitative friction measurements and qualitative articular surface staining was

performed on tibial plateau strip samples. Tibial plateau strips were affixed to 60 mm petri dishes (Becton

Dickinson #351007, Franklin Lakes, NJ) with 6 µl of cyanoacrylate (Henkel Corporation #Loctite 420,

Rocky Hill, CT). Custom indexing features were attached to each dish allowing for repeatable positioning

at subsequent test time points. Friction measurements were performed against the untreated matching

condyle explant in a creep configuration with a prescribed 4.45 N load and sliding with translation range

of ±5 mm and velocity of (1 mm/s) at room temperature (Figure 2-1B). The response of the friction

coefficient was evaluated from the normal and frictional load responses over 3600 s. The initial friction

coefficient, µinitial, and final friction coefficient, µfinal, were calculated as described previously [39]. Samples

were submerged in PBS throughout all tests and allowed to recover for a duration equal to or greater than

the test time. A two-way ANOVA on the effects of treatment and incubation time, followed by post-hoc

testing using Tukey correction, was performed on friction parameters.

2.2.3 Biocompatibility Testing

Viable explants (n=4 per treatment group for each incubation time point) in Study 2-1 were co-cultured

with an equal number of fixed explants for each glutaraldehyde concentration (Figure 2-2). Samples were

10

cultured in supplemented DMEM, changed three times a week. At each time point, viable samples from

each treatment group were sliced in half and stained for chondrocyte viability (Invitrogen #L3224,

Eugene, OR). Sample cross sections were scanned on a confocal microscope (Leica Microsystems #TCS

SP5, Buffalo Grove, IL) and combined for assessment. Viable and non viable cells were counted with the

use of software after image processing [140] (NIH #ImageJ V1.44p, Bethesda, MD). Cross sectional

specimen areas were measured using sample outlines and viable to non viable cell ratios were

normalized to sample cross sectional image area. A two-way ANOVA on the effects of treatment and

incubation time, followed by post-hoc testing using Tukey correction, was performed on the cell viability

ratio.

2.3 Results

2.3.1 Study 2-1

Glutaraldehyde treated samples showed an immediate change in appearance similar to that reported for

other tissues [103, 125, 128]. Treated samples appeared to yellow with increasing glutaraldehyde

treatment concentration and felt firmer to the touch. Curve-fitting of the stress-relaxation response to the

theoretical model produced fits with coefficients of determination that did not differ among treatment

groups or time points, producing an average and standard deviation of R2=0.987 ± .011 (representative fit

in Figure 2-3). The resulting E-Y displayed a dose-dependent increase with concentration immediately

upon fixation (p ≤ 0.034, Figure 2-4), followed by maintenance of compressive stiffness over the study

duration for the higher 0.20% and 0.60% glutaraldehyde concentrations (p ≤ 0.040) and a maintenance of

native (day 0 control) levels for the 0.02% concentration (p ≥ 0.446). With the exception of day 7 at the

0.20% glutaraldehyde concentration, the difference between same day control and higher 0.20% and

0.60% glutaraldehyde concentrations (p ≤ 0.036) was also maintained over the test duration. E+Y

displayed a similar increase with glutaraldehyde concentration with significant differences detected

between 0.60% glutaraldehyde and both day 0 and same day control values at day 7, 14, and 28 (p ≤

0.015, Figure 2-4). No differences were detected between any other factor level combinations (p ≥ 0.229).

No significant differences were detected between any factor level combinations for k (p ≥ 0.577).

11

Significant increases in µeq from native values were observed immediately following glutaraldehyde

treatment, at the higher 0.20% and 0.60% treatment concentrations (p ≤ 0.003, Figure 2-4), although

values returned close to native levels by day 14 (p ≥ 0.360). A downward trend in the time constant tµ was

observed with incubation time, with a significant difference from native values detected only for the 0.02%

treatment concentration at day 14 and day 28 (p ≤ 0.016, Figure 2-4). No significant differences were

detected across glutaraldehyde concentrations or incubation times for µmin (p ≥ 0.473, Figure 2-4). No

qualitative or quantitative differences were detected in cell viability at any time point for any tested

glutaraldehyde concentration (p ≥ 0.848, Figure 2-5).

2.3.2 Study 2-2

Friction coefficient measurements in the migrating contact configuration provided an additional measure

of long-term performance in a more physiologic loading condition. Initial friction coefficient µinitial remained

near native values for the 0.20% glutaraldehyde treated tibial plateau strips (p ≥ 0.783, Figure 2-6). At day

14, control group values were different from both native and same day 0.20% glutaraldehyde treated

group values, although the difference between same day values was not detected at day 28. Final friction

coefficient µfinal values mirrored µinitial values with 0.20% glutaraldehyde treated group maintaining near

native values over the test duration (p ≥ 0.288, Figure 2-6), and a significant difference detected

between day 14 control group and native levels (p ≤ 0.0267, Figure 2-6). After testing on day 28, surface

fibrillation was assessed qualitatively with india ink staining (Figure 2-7). All nine of the control group

samples displayed evidence of fibrillation over the contact area, while none of the glutaraldehyde-treated

test group samples showed signs of superficial zone fibrillation.

2.4 Discussion

Glutaraldehyde fixation of bovine articular cartilage xenografts offers the potential for creating a highly

functional resurfacing material for hemiarthroplasties in humans, much like the use of glutaraldehyde-

treated bovine heart valves has provided a practical treatment for heart disease. The results of this study

demonstrate that glutaraldehyde fixation of cartilage over a range of concentrations produces only

moderate changes in its mechanical properties (Figure 2-4 & Figure 2-6) and is biocompatible with live

12

tissue in co-culture (Figure 2-5). The equilibrium tensile and compressive moduli increases twofold, as

expected from the crosslinking effect of the glutaraldehyde treatment; however, the hydraulic permeability

remains unchanged; the equilibrium friction coefficient returns to control values after 7 days; the minimum

friction coefficient is nearly unchanged, and the time constant for the frictional response decreases

twofold at most, consistent with the twofold increase in the moduli and the resulting effect on the

characteristic time constant for loss of interstitial fluid pressurization [141]. When loaded in a migrating

contact configuration, glutaraldehyde fixed tissue performs almost identically to native tissue (Figure 2-6).

This outcome is consistent with the fact that the frictional response under migrating contact is dependent

on the tissue’s ability to sustain interstitial fluid pressurization continuously, which is achievable when the

Peclet number remains significantly greater than unity [39]; based on the measured material properties of

control and glutaraldehyde tissue, a sliding velocity of V=1 mm/s and an estimated contact radius of a~2

mm, the Peclet number Pe=V a/E+Y k decreased from ~130 to ~65 between control and treated samples,

remaining significantly greater than unity. Furthermore, it is found that glutaraldehyde-treated tissue is

better able to withstand repeated friction loading than untreated tissue (Figure 2-7).

Previous studies have reported results from the use of autografts for cartilage resurfacing

applications, but donor tissue supply is limited [142-147] and problems still exist with donor site morbidity

and incongruence [148]. Commercial hydrogel product testing has shown inadequate connection to

subchondral bone [149, 150]. When compared to alternative porous resurfacing materials such as

hydrogels or the current generation of engineered cartilage constructs, glutaraldehyde-treated cartilage

exhibits mechanical properties much more similar to native tissue. Most importantly, results of the 28-day

long incubation of fixed tissues at body temperature reported here suggest that they may be used

successfully under in vivo conditions.

Taken together, these highly encouraging results suggest that a properly titrated glutaraldehyde

treatment (0.20% in this case) can reproduce the desired functional properties of native articular cartilage,

and maintain these properties for at least 28 days while being incubated at body temperature. The

results of this study motivate further investigations of the potential use of glutaraldehyde-fixed xenografts

for joint resurfacing. Immature cartilage from bovine joints can be easily harvested nearly whole, by

‘popping’ it off the underlying bone as illustrated in Figure 2-8. The resulting articular layer has a clean

13

and smooth underlying surface. This fortuitous ability to harvest whole articular layers from immature

joints may facilitate their use as resurfacing materials, following glutaraldehyde treatment, with a suitable

biocompatible adhesive to attach it to a substrate (such as medical octyl cyanoacrylates) (Figure 2-9). In

a carefully controlled supply of bovine joints from animals raised for this specific purpose, animals can be

sacrificed at an age when the size of the joint (such as the bovine humeral head diameter) matches the

desired dimensions.

2.5 Acknowledgements

Research reported in this section was supported by the National Institute of Arthritis and Musculoskeletal

and Skin Diseases of the National Institutes of Health under Award Number R01 AR043628.

14

2.6 Figures

Figure 2-1: Schematic representation of bovine calf articular cartilage harvest location and friction testing schematic.

(A) Study 2-1 includes articular cartilage discs harvested from the tibial plateau, fixed with glutaraldehyde, and tested

against a smooth glass counterface. (B) Study 2-2 includes articular cartilage strips harvested from the tibial plateau,

fixed with glutaraldehyde, and tested against unfixed convex discs from the opposing condyles.

15

Figure 2-2: Representative biocompatibility culture dish with viable and glutaraldehyde treated cartilage explants.

Explants were cultured for 28 days in cell culture media at 37° C and 15% CO2. Samples from each treatment group

were sectioned and stained for chondrocyte viability at 1, 7, 14, and 28 days.

16

Figure 2-3: Representative FEBio curve-fit of experimental stress-relaxation test (5% strain) used to obtain model

parameters for E-Y, k, and ξ (0% control sample at day 14, R2=0.988).

17

Figure 2-4: Summary of mechanical properties for Study 2-1 as a function of glutaraldehyde concentration. µeq =

equilibrium friction coefficient; tµ = time constant for the rise of the friction coefficient to its equilibrium value; µmin =

minimum friction coefficient, typically achieved at the earliest time point of the friction response; E-Y = equilibrium

compressive modulus; E+Y = equilibrium tensile modulus; k = hydraulic permeability. Significant differences (p ≤ 0.05)

from native day 0 values indicated with †, differences between groups indicated with *.

18

Figure 2-5: Qualitative and quantitative biocompatibility from chondrocyte viability in sectioned co-cultured explants at

day 1 and day 28 for 0.0% control, 0.02, 0.2, and 0.6% glutaraldehyde treated explants. Samples from each

treatment group were sectioned and stained for chondrocyte viability at 1, 7, 14, and 28 days. No significant

differences were detected between groups for the ratio of viable to non viable cells normalized to tissue area.

19

Figure 2-6: Frictional properties for Study 2-2, migrating contact configuration. µinitial = friction coefficient at start of

test; µfinal = friction coefficient at end of test (1 h). Significant differences (p ≤ 0.05) from day 0 values indicated with †,

differences between groups indicated with *.

20

Figure 2-7: India ink staining of representative Study 2-2 tibial plateau strips shows surface fibrillation of the control

(CTRL) group without any apparent superficial zone damage to the 0.2% glutaraldehyde (GLUT) treated sample

group after friction testing at day 28.

21

Figure 2-8: Intact immature bovine knee condyles (A). ‘Popped’ condylar articular layer (B). Cross-section of popped

articular layer, showing the underlying surface (C).

22

Figure 2-9: Potential clinical approach for bioprosthetic arthroplasties, using glutaraldehyde-fixed xenografts. (A)

Natural shoulder joint. (B) A glutaraldehyde-treated bovine articular layer may be affixed to a hemispherical stainless

steel substrate supported on a metal stem, and inserted into the humerus in analogy to standard hemiarthroplasties.

23

CHAPTER 3: CARTILAGE WEAR DEBRIS MEASUREMENTS

VIA PARTICLE ANALYZER

3.1 Introduction

Although qualitative observations of cartilage wear debris have been made [45-52]; quantitative

measurements of cartilage wear have proven to be more challenging, with only a few studies having

reported such measurements. The primary quantitative approaches proposed to date include biochemical

assaying of cartilage and test solutions [54-57, 151], characterization of changing articular layer thickness

[54, 58-60], and changes in surface roughness [7, 57, 61, 62, 64, 65, 101]. One study examining

polyethylene wear debris in hip arthroplasty reported the use of an automated particle analyzer [152]. The

aim of this study was to test whether latest-generation particle analyzers are capable of detecting

cartilage wear debris generated during in vitro loading experiments that last 24 h or less, by producing

measurable content significantly above background noise levels.



Articular cartilage cylindrical explants were harvested from the tibial plateau of 2-3 month old calf knee

joints (n=9) obtained from a local abattoir. Full thickness cartilage discs (4 mm) were excised, placed

with articular surface down on a freezing stage microtome (Leica Instruments #SM2400, Nusslock,

Germany), embedded and frozen in a water soluble sectioning gel (Thermo Scientific #1310APD,

Rockford, IL), and trimmed of bone and deep zone tissue to obtain samples inclusive of an intact

superficial zone with a final thickness between 1.2 to 1.4 mm (n=9 per treatment group). After

preparation, samples were stored at -20° C for a period no greater than 30 days until use. Cartilage

sample pairs harvested from adjoining regions were randomized into test and control groups to produce a

study design with repeated measure.

3.2.2 Solution Preparation

24

Phosphate buffered saline solution (PBS) supplemented with a protease inhibitor cocktail (Thermo

Scientific #78438, Rockford, IL) and bactericide (Sigma-Aldrich #46878-U, St. Louis, MO) was used as a

base solution for all testing. Base solution was passed through a 0.1 µm pore filter (Millipore

#SLVV033RS, Billerica, MA) to eliminate background particulate, and care was taken to prevent

contamination. Four solution groups were prepared, one each for the loaded cartilage wear (TEST),

unloaded cartilage control (CTRL), environmental contamination control (ENVR), and base solution

contamination control (BASE) groups (Figure 3-1).


Test and control cartilage samples were thoroughly rinsed with base solution prior to testing and

submerged in 10 mL of base solution, which was aspirated and replaced with fresh solution at 1, 2, 6, and

24 h time points. Test cartilage samples were subjected to friction using a smooth glass slide. The glass

slide surface roughness was measured, Ra = 0.003±0.002 µm, using an optical profilometer with a 10X

objective and 1.81 X 1.36 mm scan area (Zygo #NV5000, Middlefield, CT). Measurements were

performed in an unconfined compression creep configuration with a prescribed 4.48 N load (equivalent to

0.36 MPa) and intermittent sliding with translation range of ±10 mm and sliding velocity of 1 mm/s at room

temperature, as used previously for the TEST group [7]. Control cartilage samples were placed in dishes

without loading platens to determine the combined effects of natural particulate shedding from cartilage

and environmental contamination for the CTRL group. Dishes without cartilage samples were used to

determine environmental solution contamination for the ENVR group and were sampled in the same way

as TEST and CTRL groups. The BASE solution particulate content was also assessed to verify

cleanliness over time Figure 3-1.

3.2.4 Particulate Analyzer

Solutions were tested with a particle sizing and counting analyzer configured to measure particulates from

0.6 to 60μm in diameter using two separate aperture tubes (30 μm and 100 μm) with a 300 μL total

aspiration volume (Beckman Coulter #Multisizer 4, Brea, CA). After testing with the 100 µm aperture tube,

solutions were then screened through a 10 µm filter (Spectrum Labs #148102, Rancho Dominguez, CA)

prior to testing with the 30 µm aperture, to prevent clogging (the 10 µm filters are first cleaned by copious

25

flushing with BASE solution). Counting analyzer calibration verification for both particulate size and

number was performed per manufacturer specifications. Solution samples were placed in clean

containers for testing, and agitated throughout test duration to keep particulates in suspension during

characterization. For each group, cumulative particulate content was calculated at each time point by

adding results from preceding time points.

3.2.5 Particulate Assay

To concentrate wear particulate in preparation for biochemical assay, remaining sample solutions were

centrifuged at 14,000 RCF for 45 minutes. The supernatant was removed and replaced with filtered

deionized water to decrease the overall salt content prior to lyophilization. This centrifugation rinse cycle

was repeated twice for each solution sample prior to lyophilization. Wear particulate lyophilisate was

digested with Proteinase K for 12 hours at 60° C in 0.1 M sodium acetate containing 5 mM EDTA, 1 mM

iodacetamide, 10 µg/mL pepstatin A, and 50mM Tris. Minimal digestion solution volumes were used (250

µL) to maximize particulate concentration for increased assay sensitivity. Solution digests were then

assayed for sulfated glycosaminoglycan (GAG) content using the 1,9-dimethylmethylene blue dye-binding

assay [153]. Hydroxyproline content was determined from aliquots of the Proteinase K digest product.

Samples were hydrolyzed by heating with equal volume of 12M HCl at 107° C for 18 hours and dried. The

hydrolysates were assayed for hydroxyproline content using a colorimetric assay [154]. Collagen content

was calculated from the aliquot fraction and hydroxyproline to total collagen ratio (7.64) [155].

3.2.6 Particulate Imaging

Representative samples were obtained for TEST, CTRL, and ENVR solutions at the 24 h time point and

concentrated according to the method above. Solutions were then incubated in 20 μM dichlorotriazinyl

aminofluorescein (Invitrogen #D16, Eugene, OR) for 30 min then mixed one to one with a water soluble

mounting gel to prevent particulate motion during imaging (Sigma-Aldrich #C 0612, St. Louis, MO). Gel

solutions were mounted on coverslips for confocal imaging and sealed to prevent evaporation [156]. Z

image stacks were obtained on a confocal microscope (Leica Microsystems #TCS SP5, Buffalo Grove, IL)

26

and combined (NIH #ImageJ V1.44p, Bethesda, MD) for qualitative visualization. Before and after testing,

images were taken of the TEST cartilage tissue samples to assess potential macroscopic damage.

3.2.7 Statistical Analysis

A two-way analysis of variance was performed for the factors of treatment (TEST, CTRL, ENVR, BASE)

and time (1, 2, 6, 24 h) using repeated measures, with type I error set at =0.05 and significance set at

p≤0.05. Post-hoc testing of the means used Tukey correction. Analyses were performed for particulate

number, particulate volume, GAG content, and collagen content (SAS Institute #SAS V9.2, Cary, NC).

3.1 Results

3.1.1 Particulate Number

TEST solution cumulative wear particulate number was found to be significantly greater than CTRL,

ENVR, and BASE solutions at all time points tested (p≤0.005, Figure 3-2A). No differences were detected

between CTRL, ENVR, and BASE solutions at any time points (p≥.97).

3.1.2 Particulate Volume

TEST solution cumulative wear particulate volume, similar to particulate number, was found to be

significantly greater than CTRL, ENVR, and BASE solutions at the 2, 6, and 24 hour time points (p<.042)

(Figure 3-2B). No differences were detected between CTRL, ENVR, and BASE solutions at any time

points (p≥.90).

3.1.3 Particulate Assay and Imaging

No differences were detected between TEST, CTRL, ENVR, and BASE solutions for either GAG (p≥.87)

or collagen content (Table 3-1, p≥.93). Qualitative visualizations from confocal imaging regarding particle

geometry (Figure 3-3) matched size distributions obtained via particle analyzer (Figure 3-4A, Figure

3-4B).

3.2 Discussion

27

The reported measurements of this study clearly demonstrate that the cartilage samples subjected to

frictional loading produce particulate content that is significantly higher than background noise and

contamination levels (Figure 3-2A, Figure 3-2B). The experimental design of this study accounted for

shedding of cartilage debris in the absence of loading, as may occur from natural enzymatic degradation

or other similar mechanisms. The design also accounted for contamination from the testing environment,

such as dust particles from the air or debris from the dishes and fluid handling equipment. By enforcing a

clean testing environment, and minimizing enzymatic degradation using protease inhibitors, it was found

that environmental contamination was negligible at all time points, in comparison to the wear produced

from frictional loading. Confocal images provide direct visual evidence of the debris characterization from

the particle counter (Figure 3-3).

Though the amount of cartilage wear observed in the TEST group was significantly higher than

background noise and contamination, it should be noted that it represented only a minute amount of loss

from the cartilage sample. As noted from Figure 3-2B, the particulate volume at 24 h averaged less than

2106 µm

3, which is less than 0.02 percent of the total volume of cartilage (1610

9 µm

3). This outcome is

not surprising, considering the relatively short 24 h duration of loading used in this test, when compared

to the normal lifetime of cartilage. It is thus remarkable that the particle analyzer used in this study is able

to detect such small but significant wear levels, otherwise not detected using standard biochemical

assays (Table 3-1). This high sensitivity will considerably facilitate future studies of the wear response of

cartilage under a variety of testing conditions, allowing investigators to test various wear-related

hypotheses.

The lubrication regime achieved in the testing conditions of this study is a combination of

interstitial fluid pressurization and boundary lubrication [157, 158]. As shown in prior studies, upon

application of the load, the interstitial fluid in the tissue pressurizes and supports most of the applied load,

producing a low friction coefficient [38]. As the pressure subsides, the load becomes supported by the

solid matrix, thus resulting in a boundary lubrication regime and a high friction coefficient [38]. For a 4 mm

diameter plug, the characteristic time for this loss of fluid pressurization is ~1400 s [141]. Therefore, for

the experiments in this study, boundary lubrication prevailed over most of the testing duration.

28

Furthermore, the magnitude of wear reported in this study is only representative of the specific testing

conditions used here. Other experimental conditions will likely produce differing amounts of wear.

In summary, the results of this study establish unequivocally that latest-generation particle

analyzers are capable of detecting very low wear levels in cartilage experiments conducted over a period

no greater than 24 h.


Research reported in this section was supported by the National Institute Of Arthritis And Musculoskeletal

And Skin Diseases of the U.S. National Institutes of Health under Award Number R01AR043628. The

authors would like to thank Mr. Aleksander V. Navratil and Dr. Elon J. Terrell for training and use of the

optical profilometer.

29

3.5 Figures

Table 3-1: Biochemical assay measurements of cartilage wear debris showing no detected difference in either

glycosaminoglycan (A, p≥.87) or collagen (B, p≥.93) content among the TEST, CTRL, ENVR, or BASE groups at any

time point (n=8 per group, 1, 2, 6, 24 h).

30

Figure 3-1: Schematic for TEST, CTRL, ENVR (cross-sectional views), and BASE (15mL centrifuge tube) solution

configurations.

31

Figure 3-2: Wear particulate measurements for bovine articular cartilage explants showing significant (* p ≤ 0.05)

increases in particulate number (A) and volume (B)when compared to unloaded CTRL, ENVR, and BASE groups at

respective time points (n=9 per group, 1, 2, 6, 24 h). No difference between CTRL, ENVR, or BASE groups was

detected (A: p≥.97, B: p≥.90).

32

Figure 3-3: Wear particulate confocal z stack and angled z stack showing qualitative agreement between observed

size distribution and particle analyzer measurements for representative 24 h TEST solution.

33

Figure 3-4: Particulate size (A) and volume (B) distribution for representative 24 h TEST solution showing a high

number of micron sized particles.

34

CHAPTER 4: INFLUENCE OF BEARING MATERIAL AND

ROUGHNESS ON CARTILAGE DEGENERATION

4.1 Introduction

Hemiarthroplasty is a surgical procedure that replaces half of a diarthrodial joint with an artificial bearing,

leaving the apposing articular cartilage layer intact. According to the Centers for Disease Control, there

are over 360,000 hemiarthroplasty operations performed each year in the US, [73]. Common indications

for hemiarthroplasty include acute bone fracture of the proximal femoral head of the hip [85], or proximal

humeral head of the shoulder [83, 84]; failed internal fixation from osteonecrosis of the diarthrodial joint

head [76, 86, 87]; and low activity expectations [79]. In all cases, hemiarthroplasty is indicated only if the

non-resurfaced native articular layer shows no significant signs of disease or damage [80].

Hemiarthroplasties are less invasive, require less surgical time and result in lower blood loss than total

arthroplasties [74, 75]. The value of hemiarthroplasty over total joint arthroplasty however is still subject to

debate [76-78], most notably for displaced femoral neck fractures [74, 79, 80], and the treatment of

primary shoulder osteoarthritis [75, 81, 82]. Patient outcomes are sometimes shown to be less successful

than with total arthroplasties, and often require revision, due in part to erosion of the articular surface [59,

88-92]. There is a general uncertainty about the cause of seemingly accelerated cartilage wear in

hemiarthroplasties.

The low friction coefficient of healthy cartilage, resulting primarily from boundary lubrication and

fluid load support, has been characterized extensively in the literature [1, 2, 4-6, 157, 159, 160]. In

particular, it is known from theory [39, 134] and experiments [39] that elevated interstitial fluid

pressurization and decreased friction can be achieved. Surface lubricating molecules acting as boundary

lubricants, such as lubricin, are known to be important as well [161]. Qualtiative and quantitative methods

for wear measurement via particulate analysis [48-50, 52, 162], biochemical assay [56, 57, 163], histology

[46, 53, 163], electron microscopy [47], magnetic resonance and radiological methods have been

developed. However, the complex relationship between friction coefficient and tissue damage and wear is

still poorly understood, and determining the underlying physical and biological events that lead to these

35

conditions remains elusive [3, 66-69]. It is generally assumed that low friction is associated with low wear

and damage, but as seen with the first generation of artificial hips, this association is not straightforward

[70-72]. Further understanding of this relationship between friction, damage, and wear is crucially

important to the common clinical procedure of hemiarthoplasty.

Material surface chemistry and geometry generally play an important role in tribology, and

advancements in material science and surface preparation have produced materials, surface coatings,

and polishing techniques for artificial joint bearing surfaces thought to minimize the degradative effects of

wear and damage to native articular cartilage in the in-vivo environment. Some investigators have

suggested that using different artificial materials for resurfacing the joint might produce less wear [164-

168]. However, a comprehensive set of quantitative data is lacking, many of the previous in-vitro studies

apply loading conditions that encourage accelerated wear, and use artificial material counterface

geometry that doesn’t meet FDA guidelines for surface roughness of artificial joint components in contact

with native tissue [167, 168]. This deviation from accepted guidelines for surface roughness of implanted

artificial joint bearing components, severely limits the potential clinical impact of those resulting material

performance comparisons.

The aim of this study was to explore use of materials used in hemiarthroplasty to glass, a material

commonly used in in-vitro studies. Two different cobalt chromium alloys along with a stainless steel alloy

accepted by the FDA for use as bearing counterfaces were selected for comparison. Material surface

preparation was conducted and tested in accordance to FDA guidelines to ensure that surface roughness

would be comparable to implanted joint components. A test configuration that promotes loss of fluid

pressurization over time was specifically selected to determine whether the equilibrium friction coefficient

is responsible for wear and damage. The hypothesis was that low equilibrium friction coefficient would be

associated with low levels of cartilage tissue wear.


4.2.1 Material Preparation

36

Cobalt-chromium low carbon (CoCrLC), cobalt-chromium high carbon (CoCrHC), and 316 low vacuum

melt stainless steel (316SS) orthopaedic material samples were obtained as cylindrical rod sections to be

used as friction platens (M. Vincent & Associates, Bloomington, MN). Platen samples were ground flat

and parallel using a surface grinder (Acer #AGS-1020AHD, Anaheim, CA). Platen samples were held in a

custom jig to maintain surface flatness, then ground and polished using successively finer silicon carbide

abrasives and final alumina oxide slurry polish steps. A group of CoCrLC platen samples was withheld

from the final polishing step to create a group with a rougher surface (CoCrLC-Ra25nm). Uncoated

optical borosilicate glass discs were used for a glass platen group (Edmund Optics #43-892, Barrington

NJ). Surface roughness measurements were performed before and after cartilage friction testing with an

optical profilometer to ensure surface finish compliance to both ISO 21534:2007, and ASTM F2033

standards with average roughness Ra less than 0.05µm for all platen samples. Six measurements were

taken on each surface.


Experiments were conducted using articular cartilage explants harvested from matched left-right pairs of

healthy tibial plateaus and opposing condyles of 2-3 months old calf knee joints obtained from a local

abattoir. Full thickness cartilage discs (4 mm) were excised from the tibial plateaus of eight joints,

placed with the articular surface down on a freezing stage microtome (Leica Instruments #SM2400,

Nusslock, Germany), embedded and frozen in a water soluble sectioning gel (Thermo Scientific

#1310APD, Rockford, IL), then trimmed of bone and deep zone tissue to obtain samples inclusive of an

intact superficial zone with a final thickness of 1.6 ± 0.05 mm (Figure 4-1). Samples were then

randomized into various treatment groups and test type. Samples were rinsed with and transferred to

filtered phosphate buffered saline (PBS) supplemented with an isothiazolone based biocide (Sigma-

Aldrich #46878-U, St. Louis, MO) and protease inhibitors.


A custom two-axis loading device equipped with a six-degrees-of-freedom load cell (JR3 Inc. #20E12A4,

Woodland, CA) was used for all mechanical testing. The device was controlled with custom LabVIEW

37

software (National Instruments Corporation #LabVIEW 2010, Austin, TX) and an associated motion

controller (National Instruments Corporation #7354, Austin, TX) to provide continuous measures of

specimen thickness, position, applied loads, and reaction loads.

Friction measurements against all platens were performed on samples (n=8), using an

unconfined compression creep loading configuration (2.22 N) with reciprocal sliding motion (±5.0 mm at 1

mm/s, Figure 4-1) [38]. Sample solutions were collected for wear particulate samples as described

previously [162]. The temporal response of the friction coefficient was evaluated from the normal and

frictional load responses over 4 h. The friction coefficient, µ was calculated as described previously [38].

Samples were submerged in PBS throughout all tests and allowed to recover for a duration equal to or

greater than the test time. Before and after test sample wet weights and macroscopic images were taken.

A two-way ANOVA on the effects of treatment and test time, followed by post-hoc testing using Tukey

correction, was performed on minimum friction coefficient µmin, friction coefficient at 98% creep

displacement (µ98%), and friction coefficient at 4 h (µ4h).

Unconfined compression stress-relaxation tests were performed on samples (n=8) after friction

testing to evaluate the equilibrium compressive modulus (EY). For each sample, a tare load was applied

(0.025N with 360 s relaxation time). After equilibrium was reached, a stress relaxation test was performed

(to 5% strain from pre-tare reference configuration, with 1200 s relaxation time). A one-way ANOVA on

the effects of treatment, followed by post-hoc testing using Tukey correction, was performed on swelling

ratio (after test to before test wet weight ratio) and compressive modulus.

4.2.3 Particulate Testing

Friction test bath solutions were tested with a particle sizing and counting analyzer configured to measure

particulates from 0.6 to 60μm in diameter using two separate aperture tubes (30 μm and 100 μm) with a

300 μL total aspiration volume (Multisizer 4, Beckman Coulter, Brea, CA). Counting analyzer calibration

verification for both particulate size and number was performed per manufacturer specifications. Solution

samples were placed in clean containers for testing, and agitated throughout the test duration to keep

particulates in suspension during characterization. Solution preparation and sample rinsing procedures

were performed as described previously [162]. A one-way ANOVA on the effects of treatment, followed by

38

post-hoc testing using Tukey correction, was performed on total particulate number and total particulate

volume.

4.2.4 Biochemical Testing

After mechanical testing, samples were bisected and weighed, and half of each construct was lyophilized

and digested in 0.5 mg/mL proteinase K solution (Fisher Scientific) overnight at 56 °C [155]. Sulfated

GAG content was determined by dimethylmethylene blue spectrophotometric assay [153, 169], collagen

content was assessed by orthohydroxyproline assay of acid-hydrolysates of proteinase K digests [169]

assuming a 1:7.64 OHP-to-collagen mass ratio [155]. Biochemical contents were normalized to tissue

sample pre-test weights. A one-way ANOVA on the effects of treatment, followed by post-hoc testing

using Tukey correction, was performed on GAG and collagen content.

4.2.5 Histological Staining

Remaining bisected samples were fixed, sectioned, and stained for collagen with safranin-o [169].

4.3 Results

4.3.1 Material Roughness Testing

No differences in individual platen group surface roughness were detected after friction testing (p ≤ 0.007,

Table 4-1). As intended, differences between the rough CoCrLC-Ra25nm group and all other platen

groups were detected both before (p ≤ 0.003, Table 4-1) and after (p ≤ 0.002, Table 4-1) friction testing.


No difference in minimum friction coefficient (µmin) was detected between any of the tested counterfaces

(Figure 4-2A, p ≥ 0.72). Differences were detected in friction coefficient at 98% creep displacement (µ98%)

and at 4 hours (µ4h) between the 316SS group and Glass, CoCrHC, and CoCrLC (p ≤ 0.01); and between

the CoCrLC – Ra 25nm and Glass, CoCrHC, and CoCrLC (Figure 4-2B, p ≤ 0.005). No differences were

detected between Glass, CoCrHC, and CoCrLC (Figure 4-2B, p ≥ 0.27) or between 316SS and CoCrLC –

39

Ra 25nm (Figure 4-2B, p ≥ 0.57). After friction testing, gross swelling in the top region was evident for

many of the samples in the 316SS and CoCrLC – Ra 25nm groups (Figure 4-3).

Swelling ratio differences were detected between the 316SS group and Glass, CoCrHC, and

CoCrLC (Figure 4-4A p ≤ 0.01), and between the CoCrLC–Ra 25nm and Glass, CoCrHC, and CoCrLC

(Figure 4-4A p ≤ 0.003).

4.3.3 Particulate Testing

Differences between Glass and CoCrLC-Ra25nm, and Glass and 316SS, were detected for both wear

particulate number (Figure 4-4C p ≤ 0.023) and particulate volume (Figure 4-4D p ≤ 0.043), however no

differences were detected between any other groups (p ≥ 0.27).

4.3.4 Biochemical Testing

Post friction test GAG content differences were detected between CoCrLC-Ra25nm and Glass, CoCrLC,

and 316SS groups (Figure 4-5A p ≤ 0.041), although no other differences were detected between groups

(Figure 4-5B p ≥ 0.98).

4.3.5 Histological Staining

After sectioning and staining, nearly all samples exhibited swelling of the top layer (Figure 4-6A). Some

samples in groups exhibiting high swelling ratio (316SS and CoCrLC-Ra25nm) showed delamination of

the top layer (Figure 4-6B).

4.4 Discussion

The results of this experimental study demonstrate that artificial bearing materials commonly used in

hemiarthroplasty may induce damage of the apposing cartilage surface, even when prepared to FDA

approved surface roughness levels and when tested using physiologic loads. These results suggest that

316SS, when compared to CoCr, should not be used as a bearing surface against articular cartilage,

contradictory to results previously published [168]. Swelling ratio tended to increase with equilibrium

friction coefficient, although the difference in wear particulate generation was not as apparent. Swelling

40

ratio more than doubled when surface roughness doubled from approximately 0.013µm to 0.027µm for

the CoCrLC to CoCrLC-Ra25nm groups, even though both were well under the 0.050µm limit required by

the FDA. Decreases in equilibrium compressive modulus mirrored swelling of the top layer (Figure 4-4).

Previous studies have shown that the layer just beneath the superficial zone of both neonatal bovine and

adult human cartilage has a low shear modulus [170, 171]. Macroscopic images show general swelling of

the top region (Figure 4-3), and while microscopic histological sections show little surface fibrillation,

failure of the region just beneath the surface near the depth of lowest shear properties [170, 171] is seen

(Figure 4-6). This could be taken to suggest that cartilage degradation associated with osteoarthritis might

initiate from the damage of this layer as opposed to wear at the surface.

Taken together, these results suggest that selection of artificial joint bearing material and surface

preparation may have an effect on the long term success of hemiarthroplasties, and that equilibrium

friction coefficient is indeed related to tissue damage. The results motivate further investigations on the

difference between articular cartilage wear and damage, and of current guidelines for the surface

preparation of orthopaedic bearing materials.


Research reported in this section was supported by the National Institute of Arthritis and Musculoskeletal

and Skin Diseases of the National Institutes of Health under Award Number R01 AR043628.

41

4.6 Figures

Figure 4-1: Schematic representation of bovine calf articular cartilage harvest location and friction testing schematic.

Articular cartilage discs harvested from the tibial plateau, trimmed of subchondral bone, and tested against flat

counterfaces of different material composition and roughness.

42

Figure 4-2: Frictional properties of cartilage against different counterface materials and roughness. Minimum friction

coefficient (µmin) (A) and a comparison of friction coefficient at time of 98% creep displacement and final time at 4h

between tissue friction test counterface materials of different composition and roughness (B). Significant differences

(p ≤ 0.05) between groups indicated with *.

43

Figure 4-3: Representative qualitative macroscopic comparison of cartilage tissue samples before and after friction

testing for glass and 316SS counterface materials. Top, front, and side views are shown. After friction testing,

swelling of the upper region is evident and highlighted by arrows in the 316SS after test images.

44

Figure 4-4: Summary of cartilage mechanical properties and wear measurements. Swelling ratio (A) (after test to

before test wet weight ratio), equilibrium compressive modulus EY (B), and wear debris particulate number (C) and

volume measurements (D) for cartilage tissue friction testing. Significant differences (p ≤ 0.05) between groups

indicated with *.

45

Figure 4-5: Summary of cartilage biochemical properties. GAG content (A) and collagen content after friction testing

normalized to sample initial wet weight. Significant differences (p ≤ 0.05) between groups indicated with *.

46

Figure 4-6: Representative histological images of top region. Cartilage tissue samples after friction testing for glass

(A) and 316SS (B) counterface materials stained with safranin-o showing damage and delamination of top layer for

316SS tested sample.

47

Table 4-1: Roughness measurements for orthopaedic material platens before and after tissue friction testing. No

differences were detected on a per material basis before and after testing for all materials, and between smooth

materials (Glass, CoCrHC, CoCrLC, and 316SS, p ≥ 0.98) or on a per material basis before and after testing (p ≥

0.99). Differences between the rough CoCrLC Ra 25nm group and all others were detected (p ≤ 0.01).

Glass CoCrHC CoCrLC CoCrLC 316SSRoughness - Ra Ra 25nmBefore Test [µm] 0.008 ± 0.005 0.011 ± 0.004 0.013 ± 0.003 0.027 ± 0.010 0.010 ± 0.004

After Test [µm] 0.010 ± 0.003 0.012 ± 0.003 0.012 ± 0.002 0.025 ± 0.008 0.012 ± 0.005

48

CHAPTER 5: CONCLUSION

5.1 Motivation

This work has been motivated by the desire to further understand degradation of articular cartilage. Most

of the past literature has focused on measurement of friction coefficient as a proxy for damage, however

understanding of osteoarthritis cannot be improved without better understanding of the mechanisms for

cartilage degradation and their relationship to friction. More specifically, the studies described in this

thesis have been motivated by the following questions:

- Can a porous material capable of sustaining interstitial fluid pressurization, such as

glutaraldehyde fixed articular cartilage, be used for functional resurfacing of damaged

articular cartilage?

- Are low levels of articular cartilage wear, associated with in-vivo conditions, possible to

measure using readily available particulate analysis equipment?

- Is material selection and the associated surface roughness characteristics important for the

performance of artificial joint components used in hemiarthroplasties?

5.2 Glutaraldehyde Fixed Cartilage As A Resurfacing Material

It was seen that glutaraledhyde fixation of bovine articular cartilage over a range of concentrations

produced only moderate changes in its mechanical properties and is biocompatible with live tissue in co-

culture. When loaded in a migrating contact configuration, glutaraldehyde fixed tissue performed almost

identically to native tissue. Furthermore, it is found that glutaraldehyde-treated tissue is better able to

withstand repeated friction loading than untreated tissue. These encouraging results suggest that a

properly titrated glutaraldehyde treatment can reproduce the desired functional properties of native

articular cartilage, and maintain these properties for at least 28 days while being incubated at body

temperature.

This study represents a necessary step before proceeding with animal experiments. Chondrocyte

cell viability was the only measurement of biocompatibility reported here. In vivo, it is possible that

glutaraldehyde-fixed cartilage wear debris within the synovial capsule might provoke an immunologic

49

response that could lead to joint pain, cell death, or further tissue degradation. Furthermore, the explant

testing configuration employed in this in vitro study does not exactly reproduce the response of a whole

joint to physiological load magnitudes. Finally, the failure characteristics of glutaraldehyde-treated

cartilage remain to be investigated. In vivo investigations might address some of the implications of this

study regarding the immunologic response from residual glutaraldehyde diffusion and fixed cartilage

debris generation within the closed synovial capsule.

5.3 Cartilage Wear Debris Measurements Via Particle Analyzer

The measurements reported here clearly demonstrate that the cartilage samples subjected to frictional

loading produce wear particulate content that is significantly higher than background noise. Confocal

images provide direct visual evidence of the debris characterization from the particle counter. Though the

amount of cartilage wear observed in the friction test group was significantly higher than background

noise and contamination, it should be noted that it represented only a minute amount of loss from the

cartilage sample. The particulate volume after 24 h of testing averaged less than 0.02 percent of the total

volume of cartilage. This outcome is not surprising, considering the relatively short 24 h duration of

loading used in this test, when compared to the normal lifetime of cartilage. It is thus remarkable that the

particle analyzer used in this study is able to detect such small but significant wear levels, otherwise not

detected using standard biochemical assays. This high sensitivity will considerably facilitate future studies

of the wear response of cartilage under a variety of testing conditions, allowing investigators to test

various wear-related hypotheses. The results of this study establish unequivocally that latest-generation

particle analyzers are capable of detecting very low wear levels in cartilage experiments conducted over a

period no greater than 24 h.

5.4 Influence of Artificial Bearing Material on Cartilage

Degeneration

The results of this experimental study demonstrate that artificial bearing materials commonly used in

hemiarthroplasty may induce damage of the apposing cartilage surface, even when prepared to FDA

approved surface roughness levels and when tested using physiologic loads. Swelling tended to increase

50

with equilibrium friction coefficient, although the difference in wear particulate generation was not as

apparent. Swelling more than doubled when surface roughness doubled even though all tested materials

were well under the limit required by the FDA. Decreases in equilibrium compressive modulus mirrored

swelling of the top layer. Macroscopic images show general swelling of the top region, and while

microscopic histological sections show little surface fibrillation, failure of the region just beneath the

surface near the depth of lowest shear properties [170, 171] is seen. This could be taken to suggest that

cartilage degradation associated with osteoarthritis might initiate from the damage of this layer as

opposed to wear at the surface.

These results suggest that selection of artificial joint bearing material and surface preparation may

have an effect on the long term success of hemiarthroplasties, and that equilibrium friction coefficient is

related to tissue damage. The results motivate further investigations on the difference between articular

cartilage wear and damage, and reevaluation of current guidelines for the surface preparation of

orthopaedic bearing materials.

51

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