Case Western Reserve University

THE EFFECT OF GAMMA RADIATION STERILIZATION ON YIELD

PROPERTIES AND MICROSCOPIC TISSUE DAMAGE IN DENSE

CANCELLOUS BONE

STEPHANIE JOY DUX

Submitted in partial fulfillment of the requirements

For the degree of Masters of Science

Thesis Advisor: Dr. Christopher Hernandez

Department of Mechanical and Aerospace Engineering

CASE WESTERN RESERVE UNIVERSITY

January, 2010 CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

______

candidate for the ______degree *.

(signed)______(chair of the committee)

______

(date) ______

*We also certify that written approval has been obtained for any proprietary material contained therein. Table of Contents

List of Tables ...... iv

List of Figures ...... v

Acknowledgements ...... ix

List of Abbreviations ...... x

Glossary ...... xi

Abstract ...... xii

Introduction ...... 1

Bone Structure and Composition ...... 2

Cortical Bone and Gamma Radiation Sterilization ...... 3

Cancellous Bone and Gamma Radiation Sterilization ...... 5

Cancellous Bone Allograft vs. Cortical Bone Allograft ...... 6

Damage in Cancellous Bone ...... 7

Objectives/Focus of Study ...... 9

Methods ...... 11

Specimen Preparation ...... 11

Mechanical Testing ...... 13

Histological Preparation ...... 17

Histomorphometry ...... 18

Statistical analysis ...... 23

Results ...... 25

Mechanical Properties ...... 25

Microscopic Tissue Damage ...... 28

Discussion ...... 31

Appendices ...... 38 ii

Appendix I: SOP – Techniques Using the Butcher Saw (The Biro Model 11) ...... 38

Appendix II: SOP – Preparing and Slabbing Bovine Long Bones ...... 42

Appendix III: SOP – Templating and Cutting Trabecular Bone Parallelepipeds ...... 47

Appendix IV: SOP – Coring Trabecular Bone Specimens ...... 52

Appendix V: SOP – Polishing Trabecular Bone Specimens ...... 58

Appendix VI: SOP – Marrow Removal for Trabecular Bone Specimens ...... 61

Appendix VII: SOP – Gamma Radiation Sterilization of Trabecular Bone Specimens ...... 64

Appendix VIII: SOP – Staining Techniques for Trabecular Bone Cores ...... 68

Appendix IX: SOP – Mechanical Testing of Trabecular Bone Using Platens ...... 71

Appendix X: SOP – Calculating Modulus from Instron Data ...... 75

Appendix XI: SOP – Cutting Toe Region from Raw Data ...... 77

Appendix XII: SOP – Clear Embedding Protocol for Cylindrical Bovine Cancellous Bone Specimens ...... 79

Appendix XIII: SOP – Staining, Polishing and Mounting of Embedded Cylindrical Bovine Cancellous Bone Specimens ...... 84

Appendix XV: SOP – Microdamage Counting of Cancellous Bone ...... 89

Appendix XVI: Matlab Code for Plotting Stress – Strain Curves and Calculating Mechanical Properties (xltension2.m) ...... 96

Appendix XVII: Stress – Strain Curves for Control Specimens ...... 100

Appendix XVIII: Stress – Strain Curves for Irradiated Specimens ...... 102

Appendix XIX: Mechanical Properties Data ...... 104

Appendix XX: Microscopic Tissue Damage Data ...... 106

Works Cited ...... 108

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List of Tables

Table 1. The mean values for bone volume fraction and mechanical properties are shown for both study groups along with the results of ANOVA comparisons. ………26

Table 2. Results of linear regression models between mechanical properties and bone volume fraction (BV/TV) are shown (y = a + b(BV/TV)). Standard error for each coefficient is listed in parentheses. ……………………….…………………………...27

Table 3. Average amounts of microscopic tissue damage in the region of interest are shown (standard deviation in parentheses) along with the results of ANOVA comparisons between the two groups. ………………………………………………………………………………………….29

Table 4. Average amounts of microscopic tissue damage are shown for our data along with data reported in literature (standard deviation in parentheses). Moore and Gibson loaded bovine proximal tibia to 1.3% strain and Wang et al. loaded bovine proximal tibia to 2% strain. (N.R. = not reported) …………………………………………………….35

List of Figures

Figure 1. The mechanical testing arrangement is shown. Cylindrical bovine cancellous bone specimens were tested between platens with a 10mm extensometer attached directly to the specimen. The bottom platen pivots to adjust for slight differences in the specimen. The specimen is colored because it has been stained with xylenol orange to mark any pre-existing microscopic tissue damage. ……………………………………15

Figure 2. The figure above shows an example of stress – strain data gathered from compression testing. The zero point is found by extending a tangent line from the steepest slope on the stress – strain graph. The elastic modulus is found by fitting a quadratic fit to the first 0.2% of the strain data and the yield point is found using the 0.2% offset method. ………………………………………………………………………….17

Figure 3. Diffuse damage is the area of green stain in the center of the image indicated with the arrow. The blue areas represent bone and the black areas represent porous space. …………………………………………………….……………………..19

Figure 4. A region of cross hatched damage is present in this trabecula and indicated with the arrow. The blue areas represent bone and the black areas represent porous space. …………………………………………………………………………………...20

Figure 5. A microcrack is seen above as the thin, sharp, linear regions of green stain in the center of the image indicated with the arrow. The blue areas represent bone and the black areas represent porous space...... ………………………………………………...21

Figure 6. A microfracture is seen above in the center of the image as the fractured trabecula surrounded with green stain indicated by the arrow. The blue areas represent bone and the black areas represent porous space. ……………………………………...22

Figure 7. Elastic modulus is significantly related to bone volume fraction for all specimens. The elastic modulus-bone volume fraction relationship did not differ between the control and irradiated specimens (p = 0.73). ……………………………………….26

Figure 8. Yield strains for the irradiated group decrease with increasing bone volume fraction, but are independent of bone volume fraction for the control group. ………….27

Figure 9. This graphically represents the percent of diffuse damage found in cross hatched regions and found in regions separate from cross hatched. The control group has a higher ratio of diffuse damage found in cross hatched regions than the irradiated group (p = 0.045). Error bars represent standard error. …….………………………………...29

Figure 10. Crack density is significantly related to bone volume fraction for the irradiated group, but is independent of bone volume fraction for the control group. .….30

Figure 11. Elastic modulus is significantly related to bone volume fraction for all specimens. Our data agrees well with the relationship between bone volume fraction and elastic modulus found in another study, EBVTV 892 8657( / ) (Wang, Guyette et al. 2005). …………………………………………………………………………………..34

Figure 12. The above figure shows 15mm thick sagittal slices of bovine distal femur. Each slice has been wrapped in saline soaked gauze and plastic wrap and labelled in order. …………………………………………………………………………………...46

Figure 13. Radiograph of bovine distal femur. ………………………………………..50

Figure 14. The left image shows a radiograph of a 15mm thick sagittal slice of a bovine distal femur. The right image shows the sagittal slice “templated” (i.e. regions of longitudinally aligned trabeculae with dimensions 15 x 30mm are marked out). ……...50

Figure 15. The image labelled A). shows a stencil of a 15mm thick sagittal slice from a bovine proximal tibia. Areas with aligned trabeculae are cut out. The image labelled B). shows a 15mm thick sagittal slice from a bovine proximal tibia. The image labelled C). depicts the sagittal slice with the stencil overlayed on top of it. The image labelled D). shows the sagittal slice with the regions of aligned trabeculae traced on to it. The image labelled E). shows the parallelpipeds with aligned trabecule cut from the sagittal slice. ……………………………………………………………………………………………51

Figure 16. The above image shows the drill press set up to core cancellous bone specimens. The coring tool is in the drill press and the specimen is gripped in the vice between two pieces of Plexiglass and submerged in a waterbath. ……………………...56

Figure 17. The above image shows a cylidrical bovine cancellous bone specimen that has been removed from its parallelpiped using the Isomet. …………………………….57

Figure 18. The above image shows the custom polishing tool. ……………………….60

Figure 19. The image to the left shows bovine cancellous bone embedded in methyl methacrylate in a glass jar. The image to the right shows the embedded bovine cancellous bone after it has been broken out of the glass jar. …………………………..85

Figure 20. The schematic shows how to trim off the excess methyl methacrylate so that thin sections can be made using the Isomet. ……………………………………………85

Figure 21. The above schematic shows how to mark the first cut on the trimmed embedded specimen. ……………………………………………………………………86

Figure 22. A microscope slide without the region of interest marked is depicted in the left image. The orange area in the left image is the polished histological section of vi

bovine cancellous bone. The region of interest was defined as a central rectangular area 7mm in the longitudinal direction and 6mm wide and is pictured in the image on the right. …………………………………………………………………………………....88

Figure 23. The above figure represents the eyepiece grid seen under the microscope. The gray area represents the area that should be examined with the eyepiece grid as it overlays the microscope slide. When point counting is used the red dots represent the points that should be counted within this eyepiece grid. ……………………………….90

Figure 24. The figure above represents an example of how to move the eyepiece grid. The grid surrounded by the red outline represents the first placement of the eyepiece grid and the gray area inside the red outline should be examined. A point on the slide should then be picked so that the eyepiece grid can be moved down to the area surrounded by the blue outline. The blue box overlaps the red box by one row. The gray area inside the blue box should be examined. This process should be repeated and the eyepiece should be moved to the green box and then the purple box. ………………………………..…90

Figure 25. The schematic above represents an example of point counting. The grid represents that eyepiece grid and the blue shape represents what we are interested in counting. The red dots represent the points that we should count because they are in the internal 8x8 grid of the eyepiece and intersect with the blue shape. …………………. 91

Figure 26. The right image shows another example of diffuse damage (indicated by the arrow). The left image shows the example of diffuse damage out of focus (indicated by the arrow). Since the area of diffuse damage is still present out of focus we know that it is diffuse damage. ………………………………………………………………………93

Figure 27. The right image shows another example of cross hatching (indicated by the arrow). The left image shows the example of cross hatching out of focus (indicated by the arrow). Since the area of cross hatching is still present out of focus we know that it is cross hatching. …………………………………………………………………………..93

Figure 28. The right image shows another example of a microcrack (indicated by the arrow). The left image shows the example of a microcrack out of focus (indicated by the arrow). Since the microcrack is still present out of focus we know that it is a microcrack. …………………………………………………………………………………………....93

Figure 29. The right image shows another example of a microfracture (indicated by the arrow). The left image shows the example of a microfracture out of focus (indicated by the arrow). Since the microfracture is still present out of focus we know that it is a microfracture. …………………………………………………………………………...94

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Figure 30. The image above shows an example of a vessel (indicated by the arrow). We can determine that it is a vessel because there are two distinct walls. ………………….94

Figure 31. The right image shows an example of a vessel that may be mistaken for a microcrack (indicated by the arrow). The left image shows this example out of focus (indicated by the arrow). Since this example appears fuzzy (not sharp) in the right image and two walls can be seen in the left image (out of focus) we know that this is a vessel and not a microcrack. …………………………………………………………………...95

Figure 32. The right image shows an example of an oblique surface which could be mistaken as microcracks (indicated by the arrows). The left image shows this example out of focus (indicated by the arrows). In the left image the “microcracks” are no longer present and instead they appear to have depth. This indicates that is an oblique surface and not a microcrack. …………………………………………………………………...95

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Acknowledgements

The work presented here would not be possible without the combined effort of many individuals. The author would like to thank the Musculoskeletal Transplant

Foundation for funding this work. Also, the author would like to thank the Case School of Engineering-Alcoa Campus Partnership Program for supporting her early work on this project. The author would like to thank Katherine Ehlert, Daniel Ramsey, Eileen Chu,

Shangjin Li, and Jay Bensusan for their technical assistance. Finally, the author would like to thank John and Abby Dux, Michael Meyers, her roommates, lab colleagues, Clare

Rimnac, PhD, and Christopher Hernandez, PhD for their continued support and guidance.

List of Abbreviations

Bone Volume………………………………………………………………………...….BV

Bone Volume Fraction……………………………………………………………...BV/TV

Celsius…………………………………………………………………………………….C

Diffuse Damage Volume………………………………………………………………..DV

Elastic Modulus………………………………………………………………………...…E

Kilogray………………………………………………………………………………..kGy

Megapascal…………………………………………………………………………….MPa

Micrometer..…………………………………………………………………………… m

Millimeter………………………………………………………………………………mm

Millimolar………………………………………………………………………………mM

Newton……………………………………………………………………………………N

Stress……………………………………………………………………………………..

Strain……………………………………………………………………………………..

Total Volume……………………………………………………………………………TV

Ultraviolet……………………………………………………………………………....UV

Glossary

Bone volume fraction: The ratio of bone volume by the total volume of a specimen.

Cancellous bone: A type of bone that is a cellular solid made up of connected rods and plates (also known as trabecular bone).

Histology: The study of anatomy at the tissue level.

Microscopic tissue damage: Damage at the tissue level that is observed through histology (also known as microdamage).

The Effect of Gamma Radiation Sterilization on Yield Properties and Microscopic Tissue Damage in Dense Cancellous Bone

Abstract

STEPHANIE JOY DUX

Cancellous bone allografts are increasingly attractive because the high porosity allows for rapid in-growth and the stiffness closely matches that of the host bone.

Allografts are sterilized with gamma radiation which can affect the mechanical properties and damage processes of bone. The purpose of this study was to determine the effects of gamma radiation sterilization on yield properties of dense cancellous bone and identify differences in microdamage accumulation. Compression tests of cylindrical bovine proximal tibia specimens sterilized at a dose of 29.32 kGy and control specimens were conducted to 1.3% strain. Histological sections were obtained to examine microscopic tissue damage. Sterilization had no effect on the yield properties of cancellous bone, but irradiated specimens had higher residual strain and significantly more microscopic tissue damage in the form of microfractures. This suggests that sterilization changes the damage processes of cancellous bone and could affect the performance of an allograft following overloading.

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Introduction

Bone grafts are often used to repair critical sized defects and to promote spinal fusion. Graft tissue acquired from a donor cadaver, known as allograft, is commonly used in orthopedic surgery, accounting for as many as 35% of all bone grafting procedures (Boyce, Edwards et al. 1999). Cortical bone is the primary form of allograft used clinically due to its availability, strength, and application to repairing defects in the cortices. Cancellous bone is also used as allograft, most commonly after being ground into small chips (morselized). Structurally intact cancellous bone allografts are attractive for grafting because their high porosity allows for more rapid and complete in-growth of new bone and vascular tissue than is possible with large cortical bone allografts. In addition, the stiffness of cancellous bone allograft more closely matches the host cancellous bone, reducing the possibility of stress shielding (Balabhadra, Kim et al.

2004). Structurally intact cancellous bone allografts are frequently used in spinal fusions and the treatment of metaphyseal defects.

The safety of the patient is of utmost importance when using bone allograft. To reduce disease transmission from the donor to the recipient, gamma radiation sterilization is often used to sterilize bone allograft before implantation. A dosage of 25 kGy is the accepted dosage for eliminating disease transmission (Kennedy 2005) although many tissue banks use a higher dosage of 30 or 35 kGy (Dziedzic-Goclawska, Kaminski et al.

2005). While gamma radiation sterilization at this dosage greatly reduces the risk of disease transmission it can also alter the mechanical properties of bone (Anderson, Keyak et al. 1992; Hamer, Strachan et al. 1996; Currey, Foreman et al. 1997; Akkus and Rimnac

2001; Mitchell, Stawarz et al. 2004; Akkus and Belaney 2005), potentially contributing to 1

allograft failure and additional surgeries (Boyce, Edwards et al. 1999). Because the accumulation and propagation of microcracks and other kinds of damage are often associated with allograft failure (Thompson, Pickvance et al. 1993; Vander Griend 1994;

Thompson, Garg et al. 2000; Enneking and Campanacci 2001), any degradation of allograft mechanical performance associated with gamma radiation sterilization has the potential to influence allograft failure in clinical situations. For this reason it is important to understand how microscopic tissue damage occurs in cancellous bone that has undergone gamma radiation sterilization. Understanding how damage occurs in sterilized cancellous bone and how and if it differs from damage in non-irradiated cancellous bone could aid in predicting failure in cancellous bone allograft.

Bone Structure and Composition

Bone is a hard connective tissue that makes up the skeletal system of the body and is an adaptive material that will heal itself and will change in structure during growth to match habitual loading. Bones provide for hematopoiesis, mineral storage, protection of organs, and mobility (Bartel, Davy et al. 2006). Bone is made up of a mineral component

(predominately of hydroxyapatite), organic part (primarily type I collagen), and water

(Jee 2001).

Bone tissue is characterized as cortical bone (also referred to as compact bone) or cancellous bone (also referred to as trabecular bone). Cortical bone is found in the diaphysis of long bones and outer shell of metaphyses and cancellous bone is found in the epiphysis of long bones and vertebral bodies (Bartel, Davy et al. 2006). Cortical and cancellous bone have the same composition, but cortical bone is solid whereas cancellous

bone is a cellular solid made up of connected rods and plates. As a result, cancellous bone is much more porous and less stiff than cortical bone (Guo 2001; Keaveny 2001).

Cortical Bone and Gamma Radiation Sterilization

Gamma radiation sterilization is known to impair cortical bone strength and resistance to fracture. In cortical bone the effects of radiation doses at levels below, at, and above the clinical dosage have been studied under monotonic loading, impact loading, fatigue, and fracture resistance (Hamer, Strachan et al. 1996; Currey, Foreman et al. 1997; Akkus and Rimnac 2001; Mitchell, Stawarz et al. 2004; Akkus and Belaney

2005). These studies reported no effects of gamma radiation on cortical bone elastic modulus for radiation doses of 36 kGy or less, but at high doses (94.7kGy), small (6%) alterations in elastic modulus were identified (Hamer, Strachan et al. 1996; Currey,

Foreman et al. 1997; Akkus and Belaney 2005). The yield stress (or load at yield) was not altered by irradiation doses less than 35 kGy (Hamer, Strachan et al. 1996; Akkus and

Belaney 2005), but was found to be reduced at doses of 60 kGy (Hamer, Strachan et al.

1996). More recently it was demonstrated that the yield strain and elastic energy decreases with radiation doses of 36.4 kGy (Akkus and Belaney 2005).

Clinical doses of gamma radiation sterilization (25-35 kGy) have been associated with reductions in work to fracture, impact energy, ultimate stress, and bending strength of cortical bone tissue (Bright RW 1983; Hamer, Strachan et al. 1996; Currey, Foreman et al. 1997; Akkus and Rimnac 2001; Mitchell, Stawarz et al. 2004; Akkus and Belaney

2005; Akkus, Belaney et al. 2005). In all cases reductions in mechanical properties increase with increasing radiation dose (Hamer, Strachan et al. 1996; Currey, Foreman et

al. 1997). Fracture resistance, fatigue life, and fatigue crack propagation resistance were also decreased by gamma radiation doses of 25-35 kGy (Akkus and Rimnac 2001;

Mitchell, Stawarz et al. 2004; Akkus and Belaney 2005; Akkus, Belaney et al. 2005).

With regard to crack propagation, fracture surfaces of irradiated cortical bone showed less microdamage generation ahead of the crack tip than non-irradiated control specimens, suggesting that less energy was required for propagation of microcracks in irradiated specimens (Akkus and Rimnac 2001; Mitchell, Stawarz et al. 2004; Akkus,

Belaney et al. 2005).

Together the preceding studies suggest that gamma radiation sterilization does not modify elastic properties of bone tissue, but does make the tissue more brittle and less resistant to crack propagation. These findings are consistent with the idea that elastic properties of bone tissue are determined primarily by the mineral component in bone while post-yield properties are determined primarily by the organic component in bone

(Burstein, Zika et al. 1975; Wang, Bank et al. 2001). Gamma radiation sterilization has been shown to alter collagen through polypeptide chain cross linking by free radicals produced through water radiolysis (Dziedzic-Goclawska, Kaminski et al. 2005). This mechanism has been confirmed by studies showing that inhibiting the movement of water molecules in bone tissue reduces the negative effect that irradiation has the failure properties (Hamer, Stockley et al. 1999; Akkus, Belaney et al. 2005). Hamer et al. found that bone irradiated when frozen (i.e. reduced mobility of water molecules) was less brittle than bone irradiated at room temperature, but in both cases irradiated tissue was more brittle than non-irradiated bone. Akkus et al. used free radical scavengers to lessen the effect that gamma radiation would have on bone by reducing the amount of cross

linking (Akkus, Belaney et al. 2005). They found that using free radical scavengers could reduce the impairment of tissue properties as compared to bone tissue irradiated without the free radical scavengers, although not to the point where failure properties matched those of non-irradiated tissue.

Cancellous Bone and Gamma Radiation Sterilization

Little is known about how a standard dosage of gamma radiation sterilization affects the biomechanics of cancellous bone. It has been found that elastic modulus, failure stress, and failure strain are not affected by 30 kGy of gamma radiation in human cancellous bone (Anderson, Keyak et al. 1992; Vastel, Meunier et al. 2004). When normalizing for apparent density, Anderson found that radiation at a dose of 60 kGy significantly reduced the elastic modulus and failure stress of cancellous bone, but a dose of 51 kGy did not have a significant effect on these two properties. When performed following lipid extraction and freeze drying, irradiation at a dose of 25 kGy caused a significant drop in work to failure and ultimate strength, without alterations in elastic modulus, as compared to tissue submitted to lipid extraction and freeze drying alone

(Cornu, Banse et al. 2000).

These studies suggest that high levels of gamma radiation sterilization cause cancellous bone to fail in a more brittle manner, but do not support the idea that clinically used levels of gamma radiation sterilization are associated with impairment of cancellous bone ductility. This differs from cortical bone where degradation in ductility is observed at a standard clinical dose of gamma radiation sterilization. It is likely that gamma radiation sterilization causes the same effects on bone tissue within cancellous bone, but

that the complex structure of cancellous bone prevents large changes in apparent mechanical performance.

While these few studies evaluated the effect that gamma radiation sterilization has on the mechanical properties of cancellous bone not all aspects of mechanical performance were examined. Apparent density was not reported in the Cornu et al. and

Vastel et al. studies making it difficult to evaluate failure stress and elastic modulus since they are dependent on apparent density. Also, the effect that gamma radiation sterilization has on the yield properties of cancellous bone is not known.

Cancellous Bone Allograft vs. Cortical Bone Allograft

Due to its porous structure, failure of cancellous bone is very different from that of cortical bone. Cortical bone can fail by a microcrack propagating into a macrocrack that eventually travels through the length of the specimen (Vashishth, Behiri et al. 1997;

Vashishth, Tanner et al. 2000). However, because of its porous structure cancellous bone cannot fail by a single macrocrack. In cancellous bone, apparent failure can involve a combination of large deformation bending and buckling and trabecular micofracture

(Michel, Guo et al. 1993; van Rietbergen, Weinans et al. 1995; Muller, Gerber et al.

1998; Nazarian and Muller 2004; Gibson 2005). Failure of cancellous bone often involves increases in the number of damaged trabeculae prior to complete failure (Arthur

Moore and Gibson 2002). However, individual trabecula can be damaged at strains well below apparent yield (Morgan, Yeh et al. 2005). As a result, changes induced in cancellous bone due to gamma radiation sterilization may be different than those in cortical bone.

Damage in Cancellous Bone

Damage in bone can be evaluated quantitatively as a reduction in mechanical performance, or through histological analysis. When evaluated through mechanical performance, damage in cancellous bone is measured as a reduction in elastic modulus, strength, or an increase in residual strain (permanent deformation) (Keaveny, Wachtel et al. 1994; Keaveny, Wachtel et al. 1999; Arthur Moore and Gibson 2002; Wang, Guyette et al. 2005). In general it has been found that the total strain applied to a specimen is related to both a reduction in elastic modulus and strength (Keaveny, Wachtel et al. 1994;

Keaveny, Wachtel et al. 1999). However, unlike metals, residual strain in cancellous bone is much less than the plastic strain (Keaveny, Wachtel et al. 1999; Currey 2002). In addition, it has been found that damage can occur at small strains (well below apparent yield) in cancellous bone. Damage measured through reductions in modulus and residual strain were found to occur in cancellous bone at apparent strains as low as 0.2% strain

(Morgan, Yeh et al. 2005).

Naturally occurring microscopic tissue damage, identified through histology, has been widely studied in human bone tissue (Burr, Forwood et al. 1997; Vashishth, Koontz et al. 2000). Microscopic tissue damage can be classified visually into one of four categories: diffuse damage, cross hatched regions, microcracks, and microfractures (Burr and Stafford 1990; Wenzel, Schaffler et al. 1996; Wachtel and Keaveny 1997; Fazzalari,

Forwood et al. 1998; Vashishth, Koontz et al. 2000; Arthur Moore and Gibson 2002;

Wang, Guyette et al. 2005; Kummari, Davis et al. 2009). Microdamage is a general term encompassing all four types of microscopic tissue damage. Microdamage can be

examined in histological sections using basic fuschsin or calcium-chelating fluorescent labeling to distinguish microdamage from damage caused by histological preparation

(Burr and Stafford 1990; O'Brien, Taylor et al. 2002).

Microdamage has been observed in specimens loaded below apparent yield as well as well past apparent yield and ultimate strains (Wachtel and Keaveny 1997; Arthur

Moore and Gibson 2002; Nagaraja, Couse et al. 2005; Wang, Guyette et al. 2005). A number of studies have noted significant increases in microdamage following loading beyond apparent yield, although no predictive relationships between the amount of microdamage and loading have been generated (Wachtel and Keaveny 1997; Arthur

Moore and Gibson 2002; Nagaraja, Couse et al. 2005). Microfracture occurrence is rare and occurs less frequently than other types microdamage (diffuse damage, cross hatched regions, and microcracks) suggesting that in normal human cancellous bone tissue individual trabecula show considerable post-yield deformation prior to failure (Wachtel and Keaveny 1997; Arthur Moore and Gibson 2002; Hernandez, Tang et al. 2005;

Nagaraja, Couse et al. 2005; Bevill, Eswaran et al. 2006). Moore and Gibson and

Nagaraja et al. found that microfractures only occurred in specimens loaded beyond yield whereas Wachtel and Keaveny found microfractures in specimens prior to yield. As total applied strain increases the total amount of microscopic tissue damage (diffuse damage, cross hatched regions, microcracks, and microfractures) has been shown to increase

(Wachtel and Keaveny 1997; Arthur Moore and Gibson 2002). At the trabecular level, as compressive strain increases, microdamage will increasingly extend across a trabecula until fracture occurs (Arthur Moore and Gibson 2002; Nagaraja, Couse et al. 2005). At the specimen level, as compressive loading is increased, microdamage increases in

amount and spreads throughout the entire specimen with increasing strain (Arthur Moore and Gibson 2002; Wang, Guyette et al. 2005).

The amount of microscopic tissue damage has been positively correlated to total applied strain, but appears to be independent of apparent density (Wachtel and Keaveny

1997). Moore and Gibson found that the crack density and crack length normalized by area are related to a reduction in modulus due to loading. As the reduction in modulus increases so does the crack density and normalized crack length.

There is still much that is not known about damage in cancellous bone. There are no predictive models relating damage measured to mechanical properties (reductions in elastic modulus and strength, residual strain) and the amount of microscopic tissue damage. Also, there are no models relating total applied strain or plastic strain to the amount of microscopic tissue damage in cancellous bone. Finally, there have been no studies on damage accumulation in irradiated cancellous bone.

Objectives/Focus of Study

To date, no studies have investigated how gamma radiation sterilization influences yield properties and the amount and type of microscopic tissue damage in cancellous bone. Because prior work suggests that gamma radiation sterilization makes cortical bone tissue more brittle, it is possible that gamma radiation sterilization will modify the damage mode in cancellous bone from one dominated by the accumulation of microcracks to one dominated by trabecular microfractures. Additionally, because damage in the form of reduced elastic modulus and residual strains have been observed at low strains (Morgan, Yeh et al. 2005), the effects of gamma radiation sterilization may

also alter microdamage accumulation/type at low loads, potentially influencing yield stress and yield strain.

The long-term goal of this work is to improve patient outcomes following allograft surgery. The objectives of this current study were to 1) determine the effects of gamma radiation sterilization on the yield properties of dense cancellous bone; and 2) identify any differences in microdamage accumulation/type associated with gamma radiation sterilization.

Methods

Specimen Preparation

Nine fresh bovine tibiae were acquired (Tucker Packing Plant, Orrville, OH,

USA). Tibiae were wrapped in saline soaked gauze and stored at -20C until specimen preparation. After removing soft tissue from the tibias, 15mm thick sagittal slices from the proximal tibia were made using a butcher saw (Biro Model 111, Cleveland, OH,

USA, Appendix II) from the still frozen bone (Keaveny, Guo et al. 1994).

Contact radiographs were made of each slice (Faxitron, 3050-010, Field Emission

Corporation, McMinnville, OR, USA) (Keaveny, Guo et al. 1994). A stencil was created of each bone slice using a light box. In each stencil, areas of uniform trabeculae 15mm by

30mm with trabeculae aligned longitudinally were identified as potential specimens. For each potential specimen a parallelepiped encompassing the specimen was identified.

Using the stencil, the parallelepipeds were traced on the actual bone slice using a wax pencil. The parallelepipeds were then cut from the bone slices using the butcher saw

(Appendix III). Again, the cancellous bone was cut while frozen and caution was taken to prevent excessive heating of the bone tissue during cutting.

A diamond tipped coring tool with an inner diameter of 8.3mm (Starlite 102095,

Rosemont, PA, USA) was used in a drill press to achieve cylindrical cores from the parallelepipeds (Appendix IV). Cylindrical specimens were made by gripping a parallelepiped in a vise between two pieces of Plexiglas. Care was taken to ensure that the longitudinal axis of the parallelepiped was aligned with the drill press (Keaveny, Guo et al. 1994; Morgan, Yeh et al. 2001). As in other steps, cutting was performed at a modest rate to ensure that the cancellous bone was not heated during coring and the cores 11

were collected with the bone specimen submerged under water. The coring tool was extended to 1mm from the bottom of the parallelepiped. The cylindrical specimen was then removed from the parallelepiped using a low speed saw (Isomet 1000 Precision Saw,

Buehler, Evanston, IL, USA).

The ends of the specimens were lightly polished to provide flat parallel ends. A polishing wheel (Ecomet 6, Buehler, Evanston, IL, USA) with 240 grit and 400 grit polishing paper was used with a custom polishing jig to keep the specimens aligned during polishing (Appendix V). After polishing, the dimensions for each specimen were recorded with five measurements of the diameter and three measurements of the length.

Marrow was removed from the specimens using a low pressure water jet (Waterpik

WP60-W, Fort Collins, CO, USA, Appendix VI). Each specimen was visually examined to ensure that it was uniform in density and without obvious heterogeneities in microarchitecture. If a specimen did not meet these requirements it was excluded from the study.

In total, thirty-four specimens were created. Specimens were wrapped in saline soaked gauze and stored in air tight plastic bags in the freezer at 20 C prior to the experiment. Specimens had an average length to diameter ratio of 9:4.

The specimens were randomly divided into an irradiated and a control group (n =

17 per group). Gamma radiation sterilization was performed by a commercial provider

(Steris Isomedix, Morton Grove, IL, USA). Specimens in the gamma radiated group were shipped overnight on dry ice to Steris Isomedix, where gamma radiation sterilization was performed. The target radiation dosage was 30 kGy as this is the standard dosage for allograft sterilization. Specimens received an average dosage of 12

29.32 kGy and were exposed for an average time of 194.33 minutes. Specimens were immediately returned to our laboratory following sterilization. Total transport time was forty-five hours. Specimens in the control group were packed in dry ice and remained in our facility while the gamma radiated specimens were in transport. When the irradiated specimens arrived at the lab all the specimens were returned to the freezer at 20 C .

Mechanical Testing

Prior to testing, specimens were thawed and stained with 0.5mM xylenol orange in distilled water to label pre-existing microscopic tissue damage (damage present naturally or caused by specimen preparation) (O'Brien, Taylor et al. 2002; Bigley, Singh et al. 2008). Specimens were stained for eight hours at 7 C (Nagaraja, Couse et al. 2005)

(Appendix VIII). After staining the specimens were rinsed thoroughly in distilled water and stored in saline prior to mechanical testing.

Specimens were tested in compression beyond yield. Mechanical testing with brass endcaps is the standard technique for mechanical testing of cancellous bone specimens because this technique reduces random systematic error associated with end- artifacts during mechanical testing (Keaveny, Pinilla et al. 1997). This technique is effective for compression testing of cancellous bone specimens with low density (bone volume fraction < 0.30) (Keaveny, Wachtel et al. 1994; Morgan and Keaveny 2001;

Bevill, Eswaran et al. 2009). When testing high density cancellous bone, the endcap approach is often not feasible as the loads experienced during testing surpass what can be supported by the glue used to secure the specimens within the endcaps. To evaluate high density cancellous bone (bone volume fraction > 0.30) others have either reduced the 13

specimen cross section on a lathe (Keaveny, Wachtel et al. 1994) and/or used platens testing (Morgan and Keaveny 2001). Platens testing was selected for the current study as the technique requires less handling and has been shown to produce only slightly more variability in elastic modulus measures than the endcap technique applied to dense bone specimens (Keaveny, Pinilla et al. 1997; Morgan and Keaveny 2001).

Specimens were placed between stainless steel platens without lubricant in a servo-hydraulic materials testing machine (Instron 8501M, Norwood, MA, USA). A pivoting joint was present on the lower platen (Figure 1). A 10mm extensometer attached to the midsection of the specimen with rubber bands (Miniature Model 3442-

010M-010-ST, Epsilon Technology Corp, Jackson, WY, USA). The specimen was held between the platens with a 10N compressive load, the extensometer was set to zero, and a compressive preload of 50N was applied. Ten preconditioning cycles to 0.2% at a strain rate of 0.5% strain per second were applied (Morgan and Keaveny 2001). Following the preconditioning cycles, the specimen underwent a final ramp to 1.3% compressive strain in displacement control. This maximum strain magnitude was chosen to ensure the specimen would exceed yield. The specimens were unloaded immediately after the ramp loading.

After compression testing the specimens were stained with 0.5mM calcein in distilled water (O'Brien, Taylor et al. 2002; Bigley, Singh et al. 2008) for eight hours at

7 C (Nagaraja, Couse et al. 2005) to identify microscopic tissue damage created by mechanical loading (Appendix VIII). After staining, the samples were thoroughly rinsed of the calcein stain with distilled water and then fixed in 70% ethanol.

Stress-strain curves were generated to determine elastic modulus, yield stress, yield strain, post-yield strain and residual strain. Since compression testing with platens requires a preload, none of the stress-strain graphs initiated at zero strain. The zero strain point was determined by extending a tangent line from the steepest slope on the stress – strain graph (ASTM 2004) (Figure 2). Occasionally a toe region was observed due to the accumulation of damage and deflection at the platens interface (Keaveny, Guo et al.

1994). Toe regions were removed from the stress-strain curve prior to calculating elastic properties (ASTM 2004) (Appendix XI).

The initial elastic modulus was defined as the slope from a quadratic fit to the first

0.2% of the stress-strain curve (Morgan, Yeh et al. 2001). Yield was defined using the

0.2% offset (Morgan, Yeh et al. 2001; ASTM 2004) (Figure 2). Total post-yield strain applied to a specimen was calculated as the difference between the maximum applied strain and the yield strain. Residual strain was defined as the strain following unloading to zero force.

Histological Preparation

Specimens were dehydrated in increasing concentrations of ethyl alcohol solutions and embedded undecalcified in methyl methacrylate (see Appendix XII).

Longitudinal slices with a thickness of 250m were made using a low speed saw (Isomet

Low Speed Saw, Buehler, Evanston, IL, USA). Two sections from the middle of each specimen were hand polished to 100m using increasingly fine sandpaper. The polished section was then rinsed in fresh distilled water for one minute under ultrasonic stimulation three times to remove polishing particles. The section was mounted on a glass slide under a cover slip. 17

Histomorphometry

Each slide was examined with transmitted light microscopy (Nikon Optishot-2,

Melville, NY, USA) at 100x magnification under ultraviolet excitation (Chroma 11000

V2, Rockingham, VT, USA). The region of interest was defined as a central rectangular area 7mm in the longitudinal direction and 6mm wide. This ensured that the region was within the extensometer gage length and at least 1 mm from the edges cut during specimen preparation (to avoid damage associated with preparation and side artifacts).

The region of interest was marked directly on the microscope slide (Appendix XIII).

Microscopic tissue damage was quantified histologically using the fluorochrome stains applied to the specimen before and after mechanical testing. In vitro staining of cancellous bone with xylenol orange or calcein covers all bone surfaces. Regions stained with calcein (post-testing stain) but not xylenol orange (pre-testing stain) therefore represent bone surfaces that were not present prior to mechanical loading (i.e. cracks and other damage caused by loading). Four distinct kinds of microscopic tissue damage were quantified: diffuse damage, cross hatched regions, microcracks, and trabecular microfracture.

Diffuse damage is defined as areas of brightly colored post stain (calcein, green under UV excitation, Figure 3). Diffuse damage can extend to the surface of a trabecula, but must also be present in the internal region of a trabecula so as not to be confused with out-of-plane trabecular surfaces (see below). A region of diffuse damage does not have distinct edges, but have edges that gradually fade out to regions of no stain (Fazzalari,

Forwood et al. 1998; Wang, Guyette et al. 2005; Kummari, Davis et al. 2009).

Figure 3. Diffuse damage is the area of green stain in the center of the image indicated with the arrow. The blue areas represent bone and the black areas represent porous space.

Cross hatched regions are an area with numerous small crossing cracks (Fazzalari,

Forwood et al. 1998). The region of cross hatching caused by loading also shows a general amount of staining that can be characterized as diffuse damage (calcein stain under UV excitation, Figure 4). As a result, cross hatching was quantified both through point counting (as a form of diffuse damage) and by quantifying each crack within a cross hatched region (Arthur Moore and Gibson 2002). Microcracks in cross hatched regions must be linear, thin, sharp, and prominent enough to be easily distinguished from the other surrounding microcracks.

Figure 4. A region of cross hatched damage is present in this trabecula and indicated with the arrow. The blue areas represent bone and the black areas represent porous space.

An individual microcrack (not part of a cross hatched region) is defined as an area of brightly colored calcein stain that is a linear, thin, sharp line, and displays a halo of damage stain (Figure 5). A microfracture is a large microcrack that extends completely through a trabecula (Figure 6).

Figure 5. A microcrack is seen above as the thin, sharp, linear region of green stain in the center of the image indicated with the arrow. The blue areas represent bone and the black areas represent porous space.

An eyepiece grid reticule with a grid spacing of 133.33m was used to determine bone volume fraction and diffuse damage using point counting. Bone volume fraction and diffuse damage area (both in and separate from cross hatched regions) were calculated through point counting. Bone volume fraction (BV/TV) was calculated by dividing the total number of points intersecting with bone (BV) by the total number of points in the region of interest (TV). Total microcrack number, crack density, total microfracture number, and average microcrack length were also determined. All microscopic tissue damage quantification was performed on blinded specimens by a single observer (SJD). Microscopic tissue damage was observed directly under a

microscope rather than using digital images of the slide to prevent errors associated with vessels and oblique trabecular surfaces. Vessels within cancellous bone are easily mistaken for cracks, but can be distinguished by thickness (vessels are thicker), the presence of two distinct walls and/or rounded edges. Trabecular bone surfaces at an oblique angle within a 100 micron histological section may therefore appear as microdamage (diffuse damage, cross hatched regions, or microcracks). To prevent counts of oblique surfaces as microdamage, the optical plane was varied to confirm that an area of microdamage (diffuse damage, cross hatched regions, or microcrack) was not part of the trabecular surface viewed in cross-section (Appendix XV).

Statistical analysis

Mechanical properties (elastic modulus, yield strain, yield stress, post-yield strain, and residual strain), bone volume fraction, and microdamage (diffuse damage, crack density, total microcrack number, total microfracture number, and average crack length) were compared between the control and irradiated groups using ANOVA (Minitab,

Version 15, Minitab Inc, State College, PA, USA). Linear regression analysis was also used to identify any relationships between the mechanical properties and bone volume fraction, microscopic tissue damage data and bone volume fraction, microscopic tissue damage data and post-yield strain, and microscopic tissue damage data and residual strain. Because the number of microfractures and the proportion of diffuse damage in cross hatched regions did not have a normal distribution, the Kruskal-Wallis non- parametric test was used for group comparisons. Additionally, the incidence rate of

specimens reaching ultimate strain or having microfractures was tested with the Fisher exact test.

Results

Mechanical Properties

Of the thirty-four specimens tested, five exceeded ultimate load in regions outside of the extensometer region and were excluded from the study (2 control and 3 irradiated) and two specimens were lost due to operator error (2 control). This left twenty-seven specimens in the study (thirteen control and fourteen gamma irradiated). Four specimens

(1 control and 3 irradiated) exceeded ultimate strain as indicated by the force and extensometer data, but were not excluded from this study because they underwent the same total applied strain as the other specimens.

No significant differences in elastic and yield properties (elastic modulus, yield strain and yield stress) were observed between irradiated and control specimens. The proportion of specimens exceeding ultimate strain in the irradiated group was not different from that in the control group (p = 0.60). Gamma irradiated specimens showed increased residual strain (p < 0.01). No significant differences in bone volume fraction were observed between the study groups (Table 1). Elastic modulus and yield stress were both significantly related to bone volume fraction (p < 0.01 each) and the elastic modulus-bone volume fraction and yield stress-bone volume fraction relationships did not differ between the control and irradiated specimens (p > 0.73, Figure 7). Yield strain was independent of bone volume fraction in the control group, but was negatively related to bone volume fraction in the irradiated group (p < 0.01, Figure 8). Post-yield strain was independent of bone volume fraction in the control group, but significantly related to bone volume fraction in the irradiated group (p = 0.04). Residual strain was not related to bone volume fraction (Table 2).

Control Irradiated p-value Bone Volume Fraction (BV/TV, 0.40 (0.10) 0.39 (0.07) 0.62 mm22/ mm ) Modulus 2541 (1513) 2217 (1201) 0.54 (MPa) Yield Strain 0.90 (0.11) 0.93 (0.11) 0.44 (%) Yield Stress 17.3 (9.7) 15.5 (7.4) 0.59 (MPa) Post-Yield 0.43 (0.13) 0.33 (0.18) 0.11 Strain (%) Residual Strain 0.19 (0.03) 0.26 (0.08) 0.01 (%)

Table 1. The mean values for bone volume fraction and mechanical properties are shown for both study groups along with the results of ANOVA comparisons.

Control Irradiated 6000 Linear Fit

y = -2686+12789x 4000 r2=0.65

2000

Elastic Modulus (MPa) 0.0 0.1 0.2 0.3 0.4 0.5 0.6 Bone Volume Fraction (BV/TV)

1.2

1.0

0.8

0.6 y = 1.345-1.062x r2=0.47 0.4

0.2 Control Irradiated

Yield Strain (%) Linear Fit for Irradiated 0.0 0.0 0.1 0.2 0.3 0.4 0.5 0.6 Bone Volume Fraction (BV/TV)

Figure 8. Yield strains for the irradiated group decrease with increasing bone volume fraction, but are independent of bone volume fraction for the control group.

Control Irradiated p- a b 2 a b p-value 2 value r r Elastic -2433 12306 -3035 13551 Modulus <0.01 0.68 <0.01 0.61 (1056) (2540) (1238) (3149) (MPa) Yield 1.34 -1.06 N.S. N.S. 0.92 N.S. <0.01 0.47 Strain (%) (0.13) (0.33) Yield -15.69 81.68 -15.99 81.29 Stress <0.01 0.73 <0.01 0.58 (6.24) (15.01) (7.82) (19.87) (MPa) Post-Yield -0.217 1.41 N.S. N.S. 0.64 N.S. 0.04 0.30 Strain (%) (0.244) (0.62) Residual N.S. N.S. 0.11 N.S. N.S. N.S. 0.37 N.S. Strain (%)

Table 2. Results of linear regression models between mechanical properties and bone volume fraction (BV/TV) are shown (y = a + b(BV/TV)). Standard error for each coefficient is listed in parentheses.

Microscopic Tissue Damage

The control specimens showed a greater amount of diffuse damage in the form of cross hatched bone area (as determined through point counting) than the irradiated specimens (p < 0.01). Trabecular microfracture was more likely to occur in the gamma irradiated specimens (p = 0.03) and irradiated specimens showed an increased number of microfractures (p = 0.02). No differences in the amount of diffuse damage, crack density, or crack length were observed between the two groups (Table 3). The diffuse damage area (diffuse damage in cross hatched regions + diffuse damage separate from cross hatched regions) did not differ between groups (p = 0.62, Figure 9). However, the control group had a higher ratio of cross hatching damage points per total diffuse damage points than the irradiated group (p = 0.045). Qualitatively, the majority of microcracks occurred within regions of cross hatching with few microcracks located outside of these regions. In regions where microfractures were found, other kinds of microscopic tissue damage (diffuse damage, cross hatched regions, microcracks, or more microfractures) were frequently observed nearby.

Control Irradiated p-value Diffuse Damage 0.00425 (0.00313) 0.00361 (0.00345) 0.62 ( mm22/ mm ) Crack Density 0.433 (0.290) 0.282 (0.257) 0.17 ( mm2 ) Crack Length ( m ) 72.16 (24.75) 77.10 (30.42) 0.65 Number of 0.154 (0.555) 1.643 (3.225) 0.02 Microfractures Table 3. Average amounts of microscopic tissue damage in the region of interest are shown (standard deviation in parentheses) along with the results of ANOVA comparisons between the two groups.

Diffuse Damage Separate from Cross Hatching Diffuse Damage in Cross Hatching Regions 100

0 Control Irradiated Percent of Diffuse Damage (%) Damage Diffuse of Percent

Figure 9. This graphically represents the percent of diffuse damage found in cross hatched regions and found in regions separate from cross hatched regions. The control group has a higher ratio of diffuse damage found in cross hatched regions than the irradiated group (p = 0.045). Error bars represent standard error.

Crack density and number of microcracks were found to increase with bone volume fraction in the irradiated group (crack density: pr0.02,2 0.38 , Figure 10, number of microcracks: pr0.01,2 0.43 ), but these relationships were not found in the control group. No other measures of microscopic tissue damage were related to bone

volume fraction. The number of microfractures was significantly related to residual strain in the irradiated group (p = 0.01, r 2  0.43 ). No relationship between the number of microfractures and residual strain was observed in the control group. All other microscopic tissue damage measures (diffuse damage, crack density, and crack length) were independent of residual strain in both groups.

Control Irradiated Linear Fit of Irradiated

) 0.8 -2

mm 0.6 (

0.4 y = -0.0675+2.295x 0.2 r2=0.38

0.0 Crack Density 0.0 0.1 0.2 0.3 0.4 0.5 0.6 Bone Volume Fraction (BV/TV)

Figure 10. Crack density is significantly related to bone volume fraction for the irradiated group, but is independent of bone volume fraction for the control group.

Discussion

The goal of the current study was to determine the effects of gamma radiation sterilization on the yield properties of dense cancellous bone and identify any differences in microdamage accumulation/type associated with gamma radiation sterilization. We found that gamma radiation sterilization does not affect the elastic or yield properties

(yield stress and yield strain) of bovine cancellous bone. This result is consistent with the idea that gamma radiation sterilization affects primarily the post-yield characteristics of bone tissue (Hamer, Stockley et al. 1999; Akkus, Belaney et al. 2005). The current study had a power of 0.7 to detect a difference in yield strain of 0.11% strain, suggesting that if gamma radiation sterilization influences cancellous bone yield strain, the difference is less than 0.11% strain. In addition, we found that gamma radiated cancellous bone had higher residual strain than the control group. Residual strain is the amount of permanent deformation in the specimen after loading and can be interpreted as a measure of the amount of damage caused by loading. As a result, increased residual strain in the irradiated specimens suggests that irradiated cancellous bone accumulates more damage and different types of damage than the control group and undergoes more permanent deformation than non-irradiated bone. In terms of microscopic tissue damage, the control specimens had a higher ratio of diffuse damage in cross hatched regions than the irradiated specimens and the irradiated specimens had more microfractures than the control specimens. In addition, irradiated specimens had a higher incidence rate of microfractures. These results support the idea that gamma radiation sterilization modifies the damage processes within dense cancellous bone. Irradiated bone has more crack propagation and brittle fracture of individual trabecula as compared to untreated

cancellous bone which shows increased accumulation of microscopic tissue damage

(cross hatching).

There are a number of strengths in our approach that lend support to our conclusions. We used a staining technique that differentiated between microscopic tissue damage created in vivo or through specimen preparation and microscopic tissue damage created through mechanical testing (O'Brien, Taylor et al. 2002). In addition, we quantified microscopic tissue damage by examining the microscope slide and not digital images. Previous studies relating microdamage to applied loading in cancellous bone performed stereological counts on digital images of the specimen cross-sections rather than through direct stereological counts under a microscope (Arthur Moore and Gibson

2002; Nagaraja, Couse et al. 2005; Wang, Guyette et al. 2005). The use of collected images rather than direct observation has the potential to cause over-counting of microcracks in cancellous bone specimens due to oblique bone surfaces within the relatively thick sections (100m ). Trabecular bone surfaces at an oblique angle within a

100 micron histological section may appear as microdamage. When quantifying microdamage directly under a microscope the observer is able to raise and lower the optical plane to distinguish between oblique or out-of-plane bone surfaces and microdamage. As stated above this helps prevent oblique surfaces being counted as microdamage.

Our study also had some limitations that must be considered. We are ultimately interested in the effect that gamma radiation sterilization has on dense human cancellous bone and therefore a limitation of our study is that we performed our experiment on bovine cancellous bone. However, bovine cancellous bone is similar to dense human 32

cancellous bone (Keaveny, Wachtel et al. 1994; Morgan and Keaveny 2001). Also, it was found that all measures of damage (reductions in elastic modulus, strength, and presence of residual strain) in cancellous bone were similar for bovine and human cancellous bone (Keaveny, Wachtel et al. 1994; Keaveny, Wachtel et al. 1999).

Therefore our results should provide a good estimate of how gamma radiation sterilization affects dense human cancellous bone.

Another limitation to our study was that we used platens for testing our specimens in compression. It is known that end-artifacts are generated when using platens testing and can generate errors in mechanical properties that can be as much as 40% (Keaveny,

Pinilla et al. 1997). End-artifact errors can be addressed by modifying the boundary conditions of the cancellous bone specimen or, if possible, attaching an extensometer directly to the specimen (Keaveny, Pinilla et al. 1997; Morgan and Keaveny 2001; Bevill,

Eswaran et al. 2009). In our study the strain data was collected from an extensometer directly attached to the specimen which reduces errors associated with platens (Keaveny,

Pinilla et al. 1997). Additionally, the fact that our results are consistent with prior work in bovine cancellous bone using the endcapping technique (see below) suggests that errors associated with the platens approach were small.

The applied loading of 1.3% strain was expected to exceed yield but not reach ultimate in bovine cancellous bone. However, four of the specimens (1 control, 3 irradiated) exceeded ultimate strain. These specimens remained in the analysis because prior work had indicated that damage processes were similar among cancellous bone specimens receiving the same total applied strain (Wachtel and Keaveny 1997).

Removing the specimens that exceeded ultimate strain from the study would not modify

the results regarding differences in the ratio of diffuse damage in cross hatched regions (p

= 0.02) and residual strain between groups (p < 0.01). Differences in number of microfractures between groups, however, would lose significance (p = 0.06). However, the conclusions of the current study regarding yield properties and differences in damage processes associated with gamma radiation sterilization would not be modified by excluding specimens loaded beyond yield.

Our results are consistent with prior examinations of the compressive properties of bovine cancellous bone. Our data for elastic modulus and bone volume fraction closely agrees with the relationship that Wang et al. found (Figure 11). Residual strain in the current study was consistent with that reported previously for similar total applied strains (Keaveny, Wachtel et al. 1999; Arthur Moore and Gibson 2002).

Control Irradiated Wang et al. 2005 6000

4000

2000

0 0.0 0.1 0.2 0.3 0.4 0.5 0.6 Elastic Modulus (MPa) Bone Volume Fraction (BV/TV)

Diffuse damage, microfracture number, and the average crack length for the control group are consistent with values reported in other studies with similar loading histories (Arthur Moore and Gibson 2002; Wang, Guyette et al. 2005) (Table 4). Our reported values of diffuse damage for both the irradiated and control groups were not significantly different from other studies with similar loading (Arthur Moore and Gibson

2002; Wang, Guyette et al. 2005) (p  0.26 ). We found numerous microfractures in the irradiated group, but only 2 in the control group which is consistent with the rates of microfracture observed in non-irradiated cancellous bone by Moore and Gibson.

Additionally, our mean crack length data for both groups agrees well with other studies.

Other studies have found that crack length does not increase with increasing strain and remains constant at about 60m (Arthur Moore and Gibson 2002; Wang, Guyette et al.

2005). It has therefore been hypothesized that the mean crack length is similar to the mean distance between lacunae and therefore may act as crack arrestors (Arthur Moore and Gibson 2002).

Moore & Wang et al., Control Irradiated Gibson, 2002 2005 (1.3% strain) (1.3% strain) (1.3% strain) (2% strain) Diffuse 0.00425 0.00361 0.006 0.0045 Damage (0.00313) (0.00345) (0.005) (0.0014) Crack Density 0.433 (0.290) 0.282 (0.257) 1.2 (1.1) 4.34 (1.97) ( mm2 ) Crack Length ( m ) 72.16 (24.75) 77.10 (30.42) 65.3 57 (29) Microfractures 0.154 (0.555) 1.643 (3.225) 0 N.R. Table 4. Average amounts of microscopic tissue damage are shown for our data along with data reported in literature (standard deviation in parentheses). Moore and Gibson loaded bovine proximal tibia to 1.3% strain and Wang et al. loaded bovine proximal tibia to 2% strain. (N.R. = not reported)

Crack density measured in our study was less than that reported in previous studies of damage in cancellous bone (Arthur Moore and Gibson 2002; Wang, Guyette et al. 2005) . However, our values do fall in the range of crack density that Moore and

Gibson reported. One explanation for the differences in microcrack measurements between our study and prior studies may be related to the methods used in quantifying the microdamage. Previous studies relating microdamage to applied loading in cancellous bone perform stereological counts on digital images of the specimen cross-sections rather than through direct stereological counts under a microscope (Arthur Moore and Gibson

2002; Nagaraja, Couse et al. 2005; Wang, Guyette et al. 2005). Because the optical plane cannot be modified in digital images, microcrack counts performed in digital images are more likely to mistake oblique cancellous bone surfaces and vessels for microcracks potentially explaining the differences in microcrack counts between our results and prior studies. Oblique bone surfaces and vessels often appear to be thin, linear, and brightly stained with the post stain and this could explain why our results for diffuse damage were similar to other studies, but our results for microcracks were much less than other studies.

Although there were no differences in yield strain, microcrack number or crack density between the study groups, the irradiated group displayed significant but subtle relationships indicating that increases in bone volume fraction were associated with reduced yield strain (Figure 8) and increased crack number and crack density (Figure 10).

These trends are consistent with prior investigations that have suggested that when cancellous bone reaches apparent yield, the amount of bone tissue that has exceeded yield within the cancellous bone structure is positively correlated with bone volume fraction

(Morgan, Bayraktar et al. 2004). In our irradiated specimens tissue yielding is more

likely to lead to microcrack formation, potentially explaining why crack number and crack density were positively related to bone volume fraction. A reduction in yield strain is also consistent with this accumulation of microscopic tissue damage.

Our results indicate that gamma radiation sterilization at a standard dosage has no effect on the yield properties of cancellous bone. However, gamma radiation sterilization affects the damage processes of dense cancellous bone as indicated by increased microfracture, reduced amounts of cross hatching type of diffuse damage, and increased residual strain. This suggests that irradiated cancellous bone has a more brittle failure and is consistent with what has been found in cortical bone. Although previous studies suggest that gamma radiation sterilization at a standard dosage does not affect the failure properties of cancellous bone (Anderson, Keyak et al. 1992; Vastel, Meunier et al. 2004).

Based on their work there is no reason to believe the structurally intact cancellous bone allograft becomes more brittle during standard clinical radiation doses. Our study demonstrates that irradiation of cancellous bone will be more prone to permanent deformation after an overloading event. Our results indicate that crack propagation and brittle fracture of individual trabecula is more prevalent in irradiated (allograft) bone.

Therefore, it is important to not cause damage to the allograft prior to transplantation and try to minimize overloading events in vivo to reduce the risk of allograft failure.

Additional studies are required to determine how these alterations in damage formation influence ultimate strength and fatigue properties.

Appendices

Appendix I: SOP – Techniques Using the Butcher Saw (The Biro Model 11)

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Techniques Using the Butcher Saw (The Biro Model 11)

Version #2 05/29/08

Author: Kathy Ehlert Principle Investigator: Christopher Hernandez

Summary: This protocol describes the handling of the butcher saw (the Biro Model 11) located in the specimen preparation room (Glennan 623). The different techniques described below include sawing, cleaning and reassembling of the saw.

Key Words: Biro Model 11, Specimen preparation, cleaning, reassembling, handling techniques

Materials: Butcher Saw (Biro Model 11), Chlorine Bleach (McMaster-Carr, # 63645T21), Sponge, Spray Bottle, Plastic Containers, Metal Guides (optional)

I. INTRODUCTION

The following procedures are to be used on the preparation of bone specimens using the Biro Model 11 saw. This also contains saw maintenance and clean-up.

SAFETY REQUIREMENTS: The use of the saw requires the user to wear protective eye gear, have their hair tied back, no hand or wrist jewelry and rolled sleeves (if they go beyond the forearm). The handling of bone specimens and chlorine bleach require the user to wear nitrile gloves and a lab coat.

II. METHODS

SAWING TECHNIQUE SAFETY NOTICE: When operating the bandsaw, DO NOT reach around the blade while the machine is on. Make sure that the blade has stopped completely before manipulating the specimen on the table.

1. Be sure to following the safety guidelines. 2. Unplug machine. 3. Check tension of saw blade and the basic assembly for loose parts. 4. Plug machine back in. 5. With protective goggles on, stand to the left of the machine and turn on. Observe the blade and the rest of the machine to ensure that the equipment is working correctly. 6. Turn off machine. 7. Take the specimen that is to be cut and place it on the table leaning against the raised edge of the sliding table. 8. Move the table (with the specimen) towards the stationary blade and align it with the blade. 9. Move the table back and turn on the machine. 10. Slowly push the table towards the blade, being careful to keep the specimen stationary. It may be useful to use metal guides to restrict the specimen from moving. Keep hands and fingers far away from the blade. The sliding table and metal guides can help with this. 11. Complete cut and turn off the bandsaw. 12. Wait until the blade is stationary to remove the specimen. 13. Repeat steps 7-12 if another cut is necessary.

CLEANING TECHNIQUE NOTE: ALL surfaces must be sanitized with 10% chlorine bleach solution, allowed to soak for at least 10 minutes with the bleach, and rinsed.

1. Unplug machine. 2. Sanitize a large, flat surface with 10% chlorine bleach solution. 3. Release the tension in the blade by squeezing silver lever (on the right hand side of the machine) and pushing the lever up. 4. Remove all parts that can be, placing the blade guard, and lower wheel guard in a plastic container and the large parts (the lower console door, sliding table, upper console door, upper wheel, and stationary table) on the surface. 5. Spray each of the large parts with the 10% chlorine bleach solution. 6. Fill container with 10% chlorine bleach solution. Make sure that all the parts are submerged in the bleach. Also, fill plastic container that is in the bandsaw with 10% chlorine bleach solution. 7. Spray stand. Both inside and outside surfaces must be coated with bleach. 8. Let everything sit for at least 10 minutes. 9. Wash all the parts in the sink with a sponge and 10% bleach solution. Rinse and set them back on the flat surface. 10. Scrub stand, making sure that all of the specimen debris has been removed from the inside surfaces. 11. Let everything dry overnight. 12. The next day reassemble (see below) and sanitize the surface.

RE-ASSEMBLING TECHNIQUE

1. Take plastic container and slide onto shelf located on the inside of the stand 2. Take upper wheel and hook into place. The two hooks are located on the back wall of the upper console. 3. Place lower wheel guard just above lower wheel. Take silver mounting cap and screw the guard to the interior wall. This is done by placing the screw attached to the guard through the large hole near the blade entrance to the stand and screw the silver cap on the other side of the wall. 4. Wrap saw blade around the top and bottom wheels. Make sure the blade faces the operator. Drop plastic guide attached to the table down so that the saw blade is in the notch. 5. Mount blade guard to its rectangular rod. The rod is attached to the upper console, coming down near the cutting area. 6. Squeeze silver lever (on the right hand side of the machine) and push the lever down. This raises the top wheel creating tension on the blade. 7. Check the blade tension and contact. The blade needs to be touching as much of the top and bottom halves of the top and bottom wheels, respectively. Readjust if necessary which is done by squeezing the silver

lever again and raising it slightly, loosening the tension on the blade. Retighten when satisfactory. 8. Place inner table on right of the saw blade making sure that the raised side is on the far right. 9. Push the two eye bolts (located on the top of the right side of the stand) in, securing the table. 10. Slide outer table onto the track on the left side of the saw blade with the raised edge nearest to the operator. 11. Slide table to the far back position and swing stopper counterclockwise to the 12 o’clock position. 12. Slide upper console door into hinges and lock shut. 13. Slide stand door into hinges which are located just inside the wall of the stand and lock shut. 14. Plug bandsaw back into outlet. 15. With protective goggles on, stand to the left of the machine and turn on. Observe the blade and the rest of the machine to ensure that the equipment is working correctly.

Appendix II: SOP – Preparing and Slabbing Bovine Long Bones

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Preparing and Slabbing Bovine Long Bones

Version #2 Date: 09/21/09

Author: Stephanie Dux Principal Investigator: Christopher Hernandez

——————————————————————————————— Summary: This protocol describes the removal of tissue from bovine long bones and the subsequent production of trabecular bone slabs. The slabs are used to prepare trabecular bone parallelepipeds which can be cored for mechanical testing or permeability testing. It is based on SOPs written by Eric A. Nauman for UCBerkely and T.P. Pinilla and M.J. Cutler for the Orthopaedic Biomechanics Laboratory at Beth Israel Hospital in Boston, Massachusetts. Note: This protocol does not apply to preparation and slabbing of human bone or the preparation of cortical bone samples.

Key Words: Sample preparation, trabecular bone, slabs, soft tissue removal, mechanical testing, bovine ——————————————————————————————— Materials:

1. Permanent, waterproof markers 2. Surgical mask 3. Scalpel blades 4. Scalpel handles 4. Safety glasses 5. Face shield 6. Butcher saw 7. Bovine long bones (tibia, humerus, femur) 8. Large ziploc bags 9. Gauze 10. Large (10”x12”) plastic container (to thaw specimens) 11. Chucks 12. Forceps 13. Biohazard bags 14. Plastic Wrap 42

15. Saline

———————————————————————————————

I. GENERAL TISSUE HANDLING GUIDELINES

Improperly handling or storing bone tissue will adversely affect its material properties (yielding inaccurate data). Thus, following these guidelines should reduce any effects of tissue processing on the subsequent mechanical properties.

1. Never let bone tissue dry out. When working with a large number of specimens, minimize the time that the bones are not submerged. Whenever possible, soak bones in a cold water bath or keep them in sealed bags/containers.

2. Never leave bones at room temperature overnight. At the end of each tissue processing session, gather the specimens together in a well-labeled (bone type, bone number, project, investigator’s initials, and date) sturdy box or ziploc bags and place them in a freezer. Prevent specimens from drying out during storage (keep the epiphyses of the long bones in sealed ziploc bags, wrap small specimens in wet gauze and place in sealed containers, etc.).

3. Minimize the number of freeze/thaw cycles the specimens must undergo. Each freeze/thaw cycle affects a bone’s material properties, so an excessive number of cycles can significantly affect the quality of the resulting test data. Never freeze specimens completely submerged in water.

4. Thoroughly thaw the bones to be handled prior to each processing session. Thawing can be achieved either by simply removing the bones from the freezer or by soaking the specimens (within bags/containers) in a cool (~21C) water bath. Depending upon specimen size, thawing can take from 30-90 minutes.

5. Never allow the specimen to overheat when cutting, coring, or performing other machining operations on them. Always insure that there is ample cooling fluid and lubrication.

6. Plan and schedule processing time in advance. Before any bones are removed from the freezer, make sure that the specimen prep lab is available, all necessary equipment is available and functional, and the session’s procedures are well understood by those involved. Only remove specimens which will be processed, and, if possible, remove the specimens well before the scheduled processing time to allow them to thaw. Thorough organization of a specimen harvest will yield more specimens, better quality specimens, and take less time.

II. BONE PREPARATION

Note: Read through the butcher saw techniques SOP to learn how to operate and clean the butcher saw.

1. Obtain bovine bones from meat packer or butcher.

2. Remove gauze from bone (this can be done by soaking the bone in water). Remove the soft tissue from the bone. Locate the site where bone is exposed (not covered by muscle). Using a #22 scalpel blade, carefully slice between the muscle and the bone to separate them. The muscle, along with a considerable amount of connective tissue can be removed in one continuous piece. Remove any remaining large pieces of muscle or connective tissue from the diaphysis, metaphysis, and epiphysis. Care should be taken to remove as much soft tissue as possible. Note: all the soft tissue (including any fresh meat) must be assumed to be biologically contaminated and disposed of in a red biohazard bag.

3. Bone can be frozen when being cut. Separate the proximal head from the diaphysis. Using a bandsaw, cut the diaphyses at least 2 inches from the beginning of the metaphysis of the proximal head (the longer the diaphysis is the easier the bone is to handle during cutting). Firmly grasp both ends of bone. Do not remove the marrow at this point. Place the unused diaphyses in a red biohazard bag and dispose of them properly or if needed for another study wrap in saline soaked gauze, label, and place in freezer.

IMPORTANT: When using the bandsaw, always wear a surgical mask (to protect against bone dust inhalation), safety glasses (to protect eyes from bone shards), and a face shield (to protect entire face).

4. Prepare the proximal head for slicing. Carefully remove all remaining soft tissue from the proximal head using #22 scalpel and forceps. Change blades frequently as the blades dull quickly. Dispose of used blades in the red sharps container. Do not put any sharps in the regular trash. Pay special attention to removing as much connective tissue as possible, for it can get caught in the bandsaw blade during slab cutting, with unpredictable consequences.

III. SLAB CUTTING

IMPORTANT: Use extreme caution when cutting slabs. Even well cleaned heads have the potential to "grab" (jam) the blade and tear out of the operator's hands. This is not a rare occurrence. Make sure the area surrounding the saw and the operator is clear of any potential obstructions or safety hazards.

KEY: Always maintain full concentration and grasp the bone firmly. Never allow your hands to be closer than 2” to the cutting blade. Use push-sticks whenever possible. In the

event of a specimen grabbing the blade, do not panic, and move hands away from the saw as quickly as is safely possible.

1. Choose sagittal (lateral), coronal (frontal), or transverse orientation of cut. For tibial specimens, coronally cut slabs have a higher yield for transversely oriented specimens, while sagittally cut slabs appear to be better suited for longitudinally oriented specimens. Additionally, heads which are oriented for sagittal slices tend to be easier to hold and control during slicing.

2. Using a micrometer or calipers, mark the bone at 15mm intervals with the scalpel in order to obtain 15 mm thick slabs.

3. Turn on bandsaw.

4. Place proximal head, resting on remaining sliced section of diaphysis and facing proper direction (sagittal, coronal, or transverse), on bandsaw cutting surface.

5. Carefully make first pass, removing outer "edge" of head. Make this cut as straight as possible while maintaining correct alignment (sagittal, coronal, or transverse). Remove as much of the outer cortical shell as possible, while avoiding regions of trabecular bone, on the first pass. This will be easier if the fence does not interfere with the condyles (i.e. the condyles are higher than the top edge of the fence).

6. Continue slicing head. Use push sticks whenever possible and use long diaphysis to aid in cutting. Discard any leftover pieces of the head (those containing no trabecular bone) into a red biohazard bag and dispose of properly.

7. Wrap each slab in saline soaked gauze and plastic wrap. Number each slab in the order that they were cut. Place in large Ziploc bag that is labeled with bone type, bone number, investigator’s name, and date.

8. Clean butcher saw according to the butcher saw techniques SOP.

Figure 12. The above figure shows 15mm thick sagittal slices of bovine distal femur. Each slice has been wrapped in saline soaked gauze and plastic wrap and labelled in order.

Appendix III: SOP – Templating and Cutting Trabecular Bone Parallelepipeds

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Templating and Cutting Trabecular Bone Parallelepipeds

Version #2 Date: 09/21/09

Author: Stephanie Dux Principal Investigator: Christopher Hernandez

——————————————————————————————— Summary: This protocol describes the templating and subsequent production of parallelepipeds to be used for mechanical testing. It is based on one written by Eric A. Nauman from UCBerkeley and T.P. Pinilla and M.J. Cutler for the Orthopaedic Biomechanics Laboratory at Beth Israel Hospital in Boston Massachusetts. Note: This protocol does not apply to the templating and specimen preparation of human bone or to the preparation of cortical bone cylinders.

Key Words: Sample preparation, trabecular bone, templating, parallelepipeds, mechanical testing, bovine ——————————————————————————————— Materials: Tracing paper Pencil and eraser Posterboard (or other heavy paper) Tape White paper Light box Centimeter ruler T-square Scissors Waxed based marker Ziploc bags (small and large) Small plastic vials Labeling tape

———————————————————————————————

I. TEMPLATING/SPECIMEN SELECTION

1. Collect the following materials: radiographs of slabs to be analyzed, a light box, tracing paper, white paper, and a pencil.

2. If the specimens are to be used for mechanical testing with endcaps, draw a 40 mm x 15 mm box on the heavy paper using a ruler and a T-square. If the specimens are to be used for mechanical testing with platens, the box should be 30 mm x15 mm. Cut out the box with scissors. Measure the resulting template using the digital calipers to insure that the internal dimensions are at least the requisite length and width. Also, for mechanical testing specimens, mark the central 10 mm of the length in order to highlight the gage length region of the template.

3. Make a template of the radiograph. Place the radiograph on the light box and place a piece of tracing paper over it. Record the specimen number and slab number on the tracing paper. Trace outlines of each slab on the tracing paper. Important: Pay attention to the alignment of the tracing paper relative to the radiograph (i.e. do not move the tracing paper while tracing) and pay special attention to accurately tracing the bottom of each slab (to allow accurate placement on bone slabs later).

4. Analyze each slice. For each slice, look for regions where trabeculae are oriented along a single axis. Place the parallelepiped template over these regions to investigate more closely. Check first that a 40 mm specimen will be obtainable which contains a minimum amount of cortical shell or growth plate. The specimen can extend up to the growth plate, but should not traverse it. Regions containing significant amounts of cortical bone have a tendency to break coring tools and should be avoided. Cortical bone resists the coring tool more than the trabecular bone and the uneven forces tend to bend the tool and cause fracture. Next, investigate the region within and immediately surrounding the gage length to check for uniform density and orientation.

5. Make sure that the tracing paper is properly aligned over the radiograph. Place the parallelepiped template over the location of the desired specimen. Using a pencil, trace the parallelepiped onto the tracing paper.

6. After all radiographs have been templated and analyzed, have an experienced lab member review each template. Though a fairly subjective process, the templating and specimen selection is perhaps the most important step in obtaining quality specimens for testing. Therefore, a thorough review by at least one other member of the lab is essential to maximize the specimen yield.

7. Using the light box trace the outlines of each slab and the parallelepipeds onto white paper. Cut out the slabs and the parallelepipeds using scissors. This creates a stencil for each slab. Be sure to label each stencil with the bone number and slab number. Also, label each parallelepiped on the stencils and tracing paper. Keep labeled tracing paper in your lab notebook.

II. CUTTING THE PARALLELEPIPEDS

1. Remove the desired slabs from the freezer and match them with the stencils.

2. Accurately position the slab beneath the appropriate outline on the template. Trace the rectangular parallelepiped cutout directly onto the slab with a wax-based marker (crayons are fine - blue, black, and green seem to show up best). Water based markers will not clearly mark wet bone specimens.

2.1 Accuracy is essential to this particular step. Make sure that the templates are exactly positioned on top of the bone slabs.

2.2 Repeat this step for all of the samples before moving on to step 3.

3. Cut out the parallelepiped with the bandsaw. The bone samples do not need to be thawed for this step. In fact, they will be easier to cut if the bone slabs are still frozen. Cut all edges as straight as possible, paying special attention to the ends. Be careful to cut along the outside edge of the marked parallelepiped region to maximize the volume of the resulting sample. Put all the remaining pieces of each slab into a red biohazard bag and dispose of properly.

4. Wrap each specimen in saline soaked gauze, wrap in plastic wrap, place in small Ziploc bag, and place in small plastic vial. Label the Ziploc bag with permanent marker and vial with labeling tape with bone and parallelepiped number. Example of label: Bovine PT5-3 12/3/08 (proximal tibia, bone #5, parallelepiped #3).

5. Place all specimen containers together in a labeled, dated, and initialed plastic bag. Store the specimens in the freezer.

6. Clean bandsaw according to SOP.

Figure 13. Radiograph of bovine distal femur.

A). B). C).

D). E). Figure 15. The image labelled A). shows a stencil of a 15mm thick sagittal slice from a bovine proximal tibia. Areas with aligned trabeculae are cut out. The image labelled B). shows a 15mm thick sagittal slice from a bovine proximal tibia. The image labelled C). depicts the sagittal slice with the stencil overlayed on top of it. The image labelled D). shows the sagittal slice with the regions of aligned trabeculae traced on to it. The image labelled E). shows the parallelpipeds with aligned trabecule cut from the sagittal slice.

Appendix IV: SOP – Coring Trabecular Bone Specimens

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Coring Trabecular Bone Specimens

Version #4 Date: 09/21/09

Author: Stephanie Dux Principal Investigator: Christopher J. Hernandez

______Summary: This protocol describes the production of cylindrical specimens of bovine trabecular bone for mechanical testing. It is based on a SOP originally written by Eric A. Nauman for the University of California Berkley. Note: This protocol does not apply to specimen preparation of human bone or the preparation of cortical bone cylinders.

Key Words: Sample preparation, trabecular bone, coring, mechanical testing, bovine ______Materials:

1. Drill Press 2. Mounted Coring Tool STARLITE 1111 Lancaster Avenue, P.O. Box 990 Rosemont, PA 19010-0911 Phone: (800) 727-1022, Fax: (610) 527-4463 Part number: 102095 for 8.3 mm diameter cores 3. 2 Portable Hand Vices 4. Plastic Container (15” Long, 9” Deep, 8” Tall) 5. Scrap Plexiglass 6. Dressing Stick 7. Wax marker 8. Isomet saw with cover 9. 4” to 6” piece of 5/16” Brass Barstock ______

I. INTRODUCTION

The following procedure was written to create cylindrical specimens of bovine trabecular bone for mechanical testing.

II. METHODS

DRILL PRESS SET-UP

1. Determine the proper coring tool diameter. For mechanical testing or permeability studies, the coring tool should have an inside diameter of approximately 8.3 mm. Smaller diameters are generally not recommended.

2. Determine the appropriate RPM for coring. The faster the setting, the greater the chance that trabeculae will be broken. It is recommended that the drill press be set to 420 RPM (although any speed up to and including 760 RPM may be acceptable).

3. Open the top cover of the drill press to check the current setting. The chart on the inside cover provides the RPM corresponding to each belt position.

4. To adjust the speed, first make sure that the drill press is unplugged. 4.1 Loosen the red levers on each side of the case (they are located in the back, near the motor) so the spindle is free to move. 4.2 Use the small, silver handle (on the right hand side, near the motor) to relax the tension in the V-belt. 4.3 Rearrange to V-belt so that it provides the appropriate RPM. 4.4 Tension the V-belt using the silver handle and tighten the two red levers without releasing the tension on the V-belt. Make sure that there is tension in the V-belt or else the coring tool may slip if you drill too quickly.

5. Mount the coring tool into the chuck and tighten the chuck with the chuck key. Be careful not to crush the coring tool when tightening the chuck

6. Turn on the drill press. If the coring tool wobbles, turn the drill press off and remove the coring tool. If reinserting the tool and retightening does not help, try another coring tool.

VICE AND SPECIMEN SET-UP

1. Fill the plastic container about 1/2-2/3 full of tap water.

2. Apply a coat of rust inhibitor to the portable hand vice.

3. Place the vice into the container and add water until the water level is at least 1 inch above the top of the vice.

4. With the drill press turned off, check that the drill press handles do not hit the plastic container when the spindle is lowered. If they do, reposition the plastic container. This will insure that nothing interferes with the travel of the coring tool.

5. Before coring any samples, clamp the dressing stick in the vice. Core into the dressing stick to help remove the metal matrix and expose the diamond particles in the bit (the diamond particles actually do the cutting).

6. Place the parallelepiped bone specimen into the vice. It is best to core the bone when it is still frozen. Make sure that there is a piece of scrap plexiglass between the specimen and the vice on both sides and that the parallelepiped is not positioned above a metal part of the vice. This will insure that the coring tool does not come in contact with the metal vice.

7. Make sure that the specimen is completely covered by water to a depth of at least 1 inch. The water serves two purposes. First it cools the specimen and the coring tool and second, it helps remove marrow and debris.

8. Situate the vice under the coring tool and adjust the height of the table so that, at maximum travel, the coring tool almost reaches the bottom of the specimen. This will insure that the specimen is almost cored out without the possibility of damaging the coring tool by drilling into the vice.

CORING

1. For safety purposes, the user must wear goggles, face shield, and nitrile gloves. In addition, the user should wear lab coat and hair net.

2. Make sure that the coring tool is clean and has been recently dressed (sharpened).

3. Position the parallelepiped under the coring tool.

4. Turn on the drill press and begin drilling. 4.1 Slowly and with very light pressure push down on the specimen. Usually, just the weight of your arm is sufficient. 4.2 Remove the coring tool completely every 5-7 seconds. Make sure that you are holding the vice in place with your free hand so that the specimen does not move. 4.3 Change the water every 5-6 specimens or when it becomes difficult to see the specimen.

5. Mark the side of the parallelepiped with a wax marker to indicate the distance that the coring tool travelled within the parallelepiped.

REMOVING CORES

1. Secure parallelepiped in Isomet brace that has two supports.

2. Attach brace to Isomet arm such that the uncored end of the parallelepiped is closest to the cutting blade.

3. Move the Isomet arm so that the wax marker line on the parallelepiped is lined up with the blade

4. Lower cover and cut off end of parallelepiped.

5. It should now be easy to slide the core out of the parallelepiped. If the core cannot be slide out move the Isomet arm over a few millimeters and cut again. Repeat until the core can be removed.

MAINTENANCE OF CORING TOOLS

1. Once drilling cores is completed drill the coring tool into the dressing stick. This will clean off any pieces of bone or marrow that are stuck inside the coring tool.

2. Clean all the tools and drill press with a 10% bleach solution.

3. Thoroughly dry the vice and drill press and spray rust inhibitor on each.

4. When disposing of the water, put a small amount of bleach in it, let it sit for a few minutes, and then dump it in the sink.

MAINTENANCE OF ISOMET

1. Completely disassemble Isomet and clean all parts and Isomet with 10% bleach solution.

2. Rinse the Isomet and all parts with water.

3. Completely dry Isomet and all parts.

ALTERNATE CORING METHOD

1. For this method an unmounted coring tool must be used (part number 101095 for 8.3 mm diameter cores).

2. Situate the vice under the coring tool and adjust the height of the table so that, at maximum travel, the coring tool just extends beyond the parallelepiped. This will insure that the specimen is completely cored out without the possibility of damaging the coring tool by drilling into the vice.

3. Follow coring steps 1-4. The only difference is that the coring tool will core entirely through the parallelepiped.

4. Once the drilling is complete remove the coring tool from the drill press. 4.1 Secure coring tool in second vice between two pieces of plexiglass with the cutting end down and below the vice jaws. Be careful not to crush the coring tool with the vice. 4.2 Use the brass barstock to push the bone specimen out. Be careful not to damage the cored specimen or the cutting end of the coring tool.

Figure 17. The above image shows a cylidrical bovine cancellous bone specimen that has been removed from its parallelpiped using the Isomet.

Appendix V: SOP – Polishing Trabecular Bone Specimens

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Polishing Trabecular Bone Specimens

Version #2 Date: 09/21/09

Author: Stephanie Dux Principal Investigator: Christopher J. Hernandez

______Summary: This protocol describes the polishing of cylindrical specimens ("cores") of bovine trabecular bone for mechanical testing. This protocol is necessary for mechanical testing using platens, but is not needed for mechanical testing using end caps. Note: This protocol does not apply to specimen preparation of human bone or the preparation of cortical bone cylinders.

Key Words: Sample preparation, trabecular bone, polishing, mechanical testing, bovine ______Materials:

1. Buehler Ecomet 6: Variable speed grinder-polisher 2. Polishing paper: Carbimet Paper Discs 240 Grit (No. 30-5108-240-100) 400 Grit (No. 30-5108-400-100) 3. Custom polishing jigs Inner diameters: 8.23mm, 8.27mm, 8.29mm 4. 4” to 6” piece of 5/16” Brass Barstock 5. Calipers ______

I. INTRODUCTION

The following procedure was written to create flat and parallel ends on cylindrical specimens ("cores") of bovine trabecular bone for mechanical testing using platens.

II. METHODS

POLISHER-GRINDER SET-UP

1. Remove covers from polisher and turn machine on.

2. Remove metal ring from each polishing wheel. Place a piece of 240 grit paper on one wheel and a piece of 400 grit paper on the other. Replace metal ring on each polishing wheel.

3. Lift up water faucets and position over each polishing wheel. Turn water at tap and turn on water on polishing machine.

4. Turn polishing wheel on and set at 40 RPM.

POLISHING

1. For safety purposes, the user must face shield, lab coat, and nitrile gloves.

2. Insert core into polishing jig. 2.1 Each polishing jig has a different inner diameter so insert the core into the polishing jig that it best fits in. 2.2 Insert core into polishing jig so that a few millimeters of the core extends beyond the polishing jig.

3. Place the core extended beyond the jig on the polishing wheel with the 240 grit paper. 3.1 Ensure that you are always holding the jig perpendicular to the wheel. 3.2 Do not put downward pressure on the jig; let the weight of the jig provide the pressure needed to polish the core.

4. When the core is flush with the polishing jig, place the jig with core on the 400 grit paper to finely polish the end polished with the 240 paper. 4.1 Ensure that you are always holding the jig perpendicular to the wheel. 4.2 Do not put downward pressure on the jig; let the weight of the jig provide the pressure needed to polish the core.

5. Remove the core from the polishing jig using the piece of barstock. 5.1 Replace the core in the jig with the other end extending slightly beyond the jig. 5.2 Polish this end following steps 3 and 4. 5.3 Again, remove core from jig.

6. Record the length three times and the diameter 5 times in different places along the specimen.

7. Replace polishing paper as it wears down.

MAINTENANCE OF ECOMET

1. Dispose of polishing paper in the biohazard trash located in 623.

2. Turn off water at facet and turn off polisher.

3. Completely disassemble polisher and clean all parts and polisher with 10% bleach solution and let sit 10 minutes. Clean surrounding counter with 10% bleach solution and let sit 10 minutes

4. Rinse the polisher, all parts, and surrounding counter with water.

5. Completely dry polisher, all parts, and counter.

Figure 18. The above image shows the custom polishing tool.

Appendix VI: SOP – Marrow Removal for Trabecular Bone Specimens Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Marrow Removal for Trabecular Bone Specimens

Version #1 Date: 01/26/09

Author: Stephanie Dux Principal Investigator: Christopher J. Hernandez

______Summary: This protocol describes the marrow removal of cylindrical specimens ("cores") of bovine trabecular bone. This protocol is necessary for damage staining and imaging. Note: This protocol does not apply to specimen preparation of human bone or the preparation of cortical bone cylinders.

Key Words: Sample preparation, trabecular bone, marrow removal, damage staining, imaging, bovine ______Materials:

1. WaterPik (2-3) 2. Saline (prepared) 3. Large plastic tub that fits in sink 4. Beakers 1 600mL beaker, 1 2L beaker 5. Chucks 6. Paper towels ______

I. INTRODUCTION

The following procedure was written to remove the marrow from bovine trabecular cores. This will make it easier for stain to infiltrate the bone for damage staining and will create cleaner images when imaging after mechanical testing.

II. METHODS 61

Note: For safety purposes, the user must face shield, lab coat, and nitrile gloves.

MARROW REMOVAL

1. Lay chucks around sink

2. Soak core in saline to thaw specimen.

3. Place large tub in sink and fill about halfway with water.

4. Assemble waterpik by connecting the water container to the top of it and attaching pick attachment. Fill water container with water, plug in, and set at highest pressure setting.

5. Submerge core and pick attachment under water in the tub. Turn on waterpik.

6. Point waterpik perpendicular to core and move in circular motion to remove marrow from all surfaces of the core.

7. Continue marrow removal process until all marrow has been removed or until there has been no change in marrow for 5 minutes.

8. Fill up waterpik water container as needed.

Note: The waterpik easily overheats and can break. It is best to have multiple waterpiks when doing marrow removal. Waterpiks should only be used for about 30 minutes at a time and then they should be allowed to cool off for many hours.

CLEAN UP

1. Put bone cores back in freezer

2. Fill water container with water and add bleach so that solution is 10% bleach. Submerge pick attachment in tub and turn on so that 10% bleach solution goes throughout waterpik.

3. Add bleach to large water tub such that the solution is 10% bleach and add beakers into tub.

4. Dispose of chucks in biohazard trash in room 625.

5. Spray surrounding counters, sink, water facet, and the external parts of the waterpik with 10% bleach solution.

6. Let all parts sit with 10% bleach for 10 minutes.

7. Fill up the waterpik's water container with water and turn on so that clean water rinses out the waterpik.

8. Rinse large tub, beakers, surrounding counters, sink, water facet, and external parts of waterpik.

9. Completely dry all pieces.

Appendix VII: SOP – Gamma Radiation Sterilization of Trabecular Bone Specimens Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Gamma Radiation Sterilization of Trabecular Bone Specimens

Version #1 Date: 01/27/09

Author: Stephanie Dux Principal Investigator: Christopher J. Hernandez

______Summary: This protocol describes how to get a quote from Steris Isomedix for gamma radiation sterilization of specimens, how to set up gamma radiation sterilization processing, and how to ship specimens on dry ice. This protocol is necessary for experiments studying the effects of gamma radiation sterilization on trabecular bone. Note: This protocol does not apply to sterilization of human bone.

Key Words: gamma radiation sterilization, trabecular bone, bovine ______Materials:

1. Radiation request form 2. Dry Ice Shipper's Training packet 3. Hazard class 9 stickers 4. Dry ice shipping container 5. Dry ice ______

I. INTRODUCTION

Steris Isomedix provides gamma radiation sterilization to boxes shipped to them. The following procedure details how to receive a quote, set up processing, and ship to Steris Isomedix.

II. METHODS

RADIATION REQUEST FORM

1. Locate the radiation request form on Steris Isomedix's website at http://www.isomedix.com/Gamma/Default.aspx?id=564.

2. Read the instructions on the website and read through the form.

3. Fill out the radiation request form according to the needs of your experiment. If you have any questions about filling out the form call Steris Isomedix at 1-847-966-1160.

4. Submit the form.

5. If you are a new customer to Steris Isomedix call them at 1-847-966-1160. There are forms that new customers need to fill out. They will check to see if you need to fill out any forms and if they do they will email them to you for you to fill out and fax back.

6. Steris Isomedix will email back a quote in a few days. The quote needs to be signed and dated and then faxed to Steris Isomedix at 847-965-2855.

SETTING UP PROCESSING

1. Ask Angelika Szakacs to call Steris Isomedix to give them the credit card information. At the time of writing, Latrice Sutherland was the financial contact at Steris Isomedix. Also, Angelika needs to mention the PI and document ID number of quote so that the credit card information is applied to the right account.

2. Submit a purchase order to Angelika.

3. Call Steris Isomedix to ensure that they have all the forms and information needed.

4. If priority or same day processing is being used, call Steris Isomedix to schedule. Note: If same day processing is being used it needs to be shipped First Overnight with FedEx.

SHIPPING SPECIMENS TO STERIS ISOMEDIX

1. Specimens need to be shipped on dry ice to ensure that they remain frozen. Dry ice is considered a hazardous material and therefore Dry Ice Shipper's Training needs to be completed through Case's DOES. Contact DOES and request the training materials, complete the training materials, return to DOES, and wait to receive certification.

2. After training has successfully been completed get a hazard class 9 sticker from DOES.

3. Talk with Sheila Campbell early in the morning the day that the package is being shipped. Tell Sheila what time the package needs to be shipped out and ask her to schedule a pickup with FedEx. Give Sheila the shipping address and account number to be charged.

4. Locate a dry ice shipping container in Glennan 623 or Wearn 511.

5. Get dry ice from Greenfield's lab on the 3rd floor of the BRB. Go in the morning to ensure that you get dry ice.

6. Pack specimens in dry ice container according to the guidelines in the training packet. Drop package off with Sheila on the 4th floor.

7. Attach FedEx airway bill and include a copy of the radiation request form in the same envelope.

Appendix VIII: SOP – Staining Techniques for Trabecular Bone Cores Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Staining Techniques for Trabecular Bone Cores

Version #3 5/28/09

Author: Daniel Ramsey/Kathy Ehlert Principal Investigator: Christopher Hernandez

SUMMARY: This protocol describes the staining process for trabecular bone cores. The stains indicate damage in the cores before and after load testing. This is an adaptation of the sterile Xylenol orange and Calcein staining preparation SOPs written by Cameron Patthanacharoephon and Evgeniy Tkachenko and the staining protocol used by Seetha Ramudu Kummari (Calcein Staining_SOP_version 1) and Kathy Ehlert. This is also based off the protocol described in the article written by R.F. Bigley called “Validity of serial milling-based imaging system for micro-damage quantification.” Revisions have been made based on the article by O’Brien called “An improved labeling technique for monitoring microcrack growth in compact bone”. KEY WORDS: Fluorescent bone labeling, trabecular bone cores, Xylenol orange solution, Calcein solution MATERIALS: Sodium Chloride (S9888, Sigma), Sodium Bicarbonate (S5761, Sigma), Calcein (190167, MP Biomedicals), Xylenol Orange (152269, MP Biomedicals), Distilled Water, 1N Hydrochloric Acid Solution, 1N Sodium Hydroxide Solution EQUIPMENT: Scale, hood, stir plate, graduated cylinder, stirring bar, aluminum foil, two brown bottles for storage, Electronic pH meter, vacuum, glass containers (with lids), shaker

INTRODUCTION This protocol works with Xylenol Orange, Calcein, Sodium Chloride, and Sodium Bicarbonate solutions. Both the Xylenol Orange and Calcein solutions have a molarity of 0.5 mM.

METHODS Note: When handling Xylenol Orange or Calcein solutions, the solutions MUST be stored either in a beaker wrapped in aluminum foil or a brown glass bottle. 1. Prepare the 2% Sodium Bicarbonate physiological saline solution. 2. Place an appropriate sized beaker on the stir plate. 3. Wrap beaker in aluminum foil. 4. Mix 900 mg NaCl 100 mL distilled water 68

with a stir bar in the beaker, preparing a 5. 0.9% (w/v) NaCl solution. 6. Note: Make sure that you have prepared enough NaCl solution for both the xylenol orange solution and the Calcein solution. It is recommended that you prepare at least 20 times the total volume of the bones for each solution. 7. Slowly add 20mg NaHCO 3

1 mL 0.9% NaCl Solution to the 0.9% (w/v) NaCl solution to make a 2% Sodium Bicarbonate saline solution. 8. Prepare the Xylenol Orange solution. 9. Slowly add 0.3830 mg Xylenol Orange

1 mL 2% NaHCO solution and stir with stir bar until completely3 dissolved. This may take several hours to complete. 10. Check the pH of the solution and make sure that it still is 7.4 11. Take PH meter and calibrate 12. Turn on PH meter 13. Hit “1” on the side of the meter, and drop a dot of the yellow solution (pH=7) at the tip of the meter. The reading should flash and then stabilize 14. Rinse tip in dital water 15. Hit “2” on the side of the meter, and drop a dot of the blue solution (pH=10) on the tip of the meter 16. Rinse tip in the distal water 17. Now the pH meter is ready to use 18. Dip the tip of the pH meter into the solution on the stir plate. It will be around 7.8 19. 2.3. Add HCL 1N solution by drops until the pH meter reads 7.4 20. 2.4.Store in brown glass bottle (if necessary). 21. Prepare Calcein staining solution. 22. Slowly add 0.3113 mg Calcein

1 mL 2% NaHCO3 solution and stir with stir bar until completely dissolved. This should only take a few minutes to complete. 23. Check the pH of the solution and make sure that it still is 7.4 24. Repeat all the steps of 2.2 to get the pH meter ready 25. Add HCL 1N solution by drops until the pH meter reads 7.4 26. Store in brown glass bottle (if necessary). 27. Gently place trabecular bone cores into beaker with Xylenol Orange solution (use at least 20x total bone volume). 28. Vacuum samples. 29. Place samples in vacuum chamber. 30. Allow the vacuum settle at 30mmHg (note: the vacuum pump, the gauge is offset by 5mmHg so you must go to approximately 35 mmHg). 31. Let specimens sit for 60 minutes.

32. Slowly release vacuum. 33. Repeat steps 5.1-5.4 three times. 34. Remove samples from vacuum chamber. 35. Rinse samples. 36. Place samples in a glass container (with lid) filled with distilled water and place on the shaker for 2 minutes. 37. Dispose of distilled water in Xylenol Orange waste container. 38. Repeat 6.1-6.2 three more times. 39. Place specimens in .9%NaCl solution for at least 30 minutes prior to testing to allow remaining stain to wash out. 40. Place specimens in a new container of .9%NaCl solution. 41. Dispose of Xylenol Orange staining solution in Xylenol Orange waste container by bleaching for 10 minutes then pouring waste down the sink. 42. Test bone cores on material testing device. 43. Gently place trabecular bone cores into beaker with Calcein solution (use at least 20x total bone volume). 44. Vacuum samples. 45. Place samples in vacuum chamber. 46. Allow the vacuum settle at 30mmHg (note: the vacuum pump, the gauge is offset by 5mmHg so you must go to approximately 35 mmHg). 47. Let specimens sit for 60 minutes. 48. Slowly release vacuum. 49. Repeat steps 11.1-11.4 three times 50. Remove samples from vacuum chamber. 51. Rinse samples. 52. Place samples in a glass container (with lid) filled with distilled water and place on the shaker for 2 minutes. 53. Dispose of distilled water in Calcein waste container. 54. Repeat 6.1-6.2 three more times. 55. Dispose of Calcein staining solution and distilled water by bleaching for 10 minutes then pouring waste down the sink.

Appendix IX: SOP – Mechanical Testing of Trabecular Bone Using Platens

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Mechanical Testing of Trabecular Bone Using Platens

Version #4 Date: 06/02/09

Author: Stephanie Dux, Eileen Chu Principal Investigator: Christopher J. Hernandez ______Summary: This protocol describes how to operate the Instron machine located in Wearn 505 and how to perform compression testing on trabecular bone using platens. Note: This protocol does not apply to the mechanical testing of human bone, cortical bone cylinders, or the testing of trabecular bone using endcaps.

Key Words: Mechanical testing, trabecular bone, mechanical testing, platens, bovine ______Materials:

1. Instron materials testing machine 2. Saline (prepared) 3. Pivoting platens 4. 10mm extensometer 5. Camel rubber bands 6. Crochet hook 7. Beaker, 600mL beaker 8. Chucks 9. Paper towels 10. Kimwipes 11. Grease ______

I. INTRODUCTION

The following procedure was written to the mechanical testing of bovine trabecular bone cylindrical specimens (cores). This protocol uses pivoting platens to test the bone cores.

Therefore, it is necessary that the ends of the bone cores have been polished so that they are flat and parallel to each other.

II. METHODS

Note: For safety purposes, the user must use goggles, lab coat, and nitrile gloves.

SET-UP

1. Lay chucks on counter space.

2. Allow cores to thaw for 30 minutes before testing. Soaking the cores in saline can speed up the thawing process.

TURNING ON AND SETTING UP INSTRON

1. Turn on pump by turning on new Instron machine 2. Turn on Instron machine 3. Move actuator at midpoint a. Set point, set at zero. This step zeros the load cell. b. Be very careful when doing this 4. Lower crosshead a. Release crosshead clamp b. Use “raise” and “lower” knobs to move it up and down c. Clamp crosshead 5. Position safety clamp as a safety precaution 6. Attach pivoting platens to Instron. a. The “pivoting” platen needs to be on the bottom 7. Plug in 10mm extensometer into output 1. 8. Ensure that the platens are aligned.

9. Ensure that the pivoting platen is pivoting. If the pivoting platen seems sticky or does not pivot easily, take the pivoting platen apart, use a kimwipe to remove the old grease, and then put new grease on the pivoting platen. 10. Make sure that the PID values for load are correct. Check with Jay Bensusan on the correct PID values, they will vary for each load cell.

MECHANICAL TESTING

1. Attach 10mm extensometer to specimen with camel rubber bands. The crochet hook can help with this. Center the specimen in the extensometer. When handling the extensometer, remember to hold it by the sides (do not compress the top and bottom). 2. Put machine in displacement control 3. Place specimen in machine between two platens a. center specimen between the platens b. lower platens onto the specimen placing about -10N of preload c. make sure there is enough clearance between the extensometer and platens so the extensometer will not be damaged during the test. (usually the limit is 16 mm on a 10 mm extensometer) 4. Set displacement minimum and maximum limits. a. Minimum and maximum limits should be set to +/- 1mm of the current position b. Action=transfer and hold 5. Balance the extensometer 6. Switch machine into load control 7. Set point to -50N (-25 if human trabecular bone) a. Ensure strain goes negative when compressive load is applied b. If strain goes positive, place machine in displacement control, remove specimen, reposition extensometer, and replace specimen and apply -50N (-25 if human trabecular bone) load and check that strain goes negative. 8. Switch machine into displacement control 9. Set waveform a. Double ramp to -2mm (-5% calculated strain) b. Set rate to calculated 0.5% strain/sec. Make sure the units are correct. 10. Set strain1 minimum limits a. Set minimum limit to -0.2% strain b. Action = reset c. Turn limits on 11. Set time for test in data acquisition test = 4 sec 12. Run data acquisition 73

13. Run test 14. Save data 15. Record the load at which strain1=-0.2% strain 16. Switch machine to load control 17. Set waveform a. Waveform = haversine b. Amplitude = load at which strain1 is -0.2% c. Rate = 1.25Hz 18. Set event detection on to 10 cycles 19. Set data acquisition time = 10 sec 20. Run data acquisition and waveform 21. Save data 22. Switch machine to displacement control 23. Set strain1minimum limit a. Set minimum limit to -1.3% strain b. Action=reset c. Turn limit on 24. Set data acquisition time 25. Run data acquisition and waveform 26. Save data 27. Unload specimen 28. Remove specimen from machine

TURNING OFF INSTRON

1. Raise crosshead 2. Shut down Instron machine (hit 0 on the controller) 3. Shut off pump (ask Jay about this step) 4. Clean extensometer and platens with alcohol

Appendix X: SOP – Calculating Modulus from Instron Data

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Calculating Modulus from Instron Data

Version #1 Date: 01/27/09

Author: Stephanie J. Dux Principal Investigator: Christopher J. Hernandez

______Summary: This document outlines the protocol for calculating sample modulus, yield strain and strength and ultimate strain and strength automatically from Instron raw data. It is based on a SOP originally written by Christopher J. Hernandez for the University of California Berkley ______Key Words: Instron, modulus, stress, strain ______Materials: data files from Instron, Matlab. ______

I. FORMATTING DATA FILES

1. The data file in Excel should show the data in columns in the following order: Time (seconds) Load (Newtons) Displacement (mm) Strain (%) Displacement from extensometer (if used) (mm)

2. Transfer file to the workstation you will be using Matlab at. 3. Place files in the same folder as xltension.m, xltension1.m, xltenion2.m, xltenstion3.m, and xltensionTC. I find it easiest to create a folder for every day of testing and place files from that day and copies of the m files in the folder. 4. In Matlab change the files from text files to data files. In the current directory window, right click on the file, click rename, and delete the .txt.

II. CALCULATING MODULUS, STRAIN AND STRENGTH

1. Edit the file xltensionTC. The file is made up of three columns of data and an end of file command (END). Input the name of the data file, the average diameter and the average effective length in order separated by tabs (if endcaps are used the effective length is one half the sum of the average sample length and the average length between endcaps, if platens are used the effective length is the length of the specimen). Only include rows for the data files you are analyzing. 2. Open matlab. From the directory where your data files and xltension.m and xltensionTC are located, run the program xltension, xltension1, xltension2, xltension3 (type xltension). a. xltension.m is the original m-file from UC Berkley b. xltension1.m can be used to graph the data. Stress vs. strain, stress vs. time, or any other parameters can be graphed against each other. c. xltension2.m calculates modulus and yield strain using the method outlined in Morgan, E.F, et al, 2001, “Nonlinear Behavior of Trabecular Bone at Small Strains,” J of Biomech. Eng., 123, pp 1-9 and plots. d. xltension3.m calculates modulus between 0.1 and 0.4% strain and plots. 3. The output for each file you’ve listed in xltensionTC appears on the screen in the following order: Modulus, Yield Strain, Yield Stress, Ultimate Strain, Ultimate Stress 4. The same data is recorded in new files named for each of your data files (namedat) and in a summary file (outmm-dd-yy where mm, dd, yy are month day and year).

Note: If a data file does not reach yield it will cause an error in xltension.m, xltension2.m, and xltension3.m.

Appendix XI: SOP – Cutting Toe Region from Raw Data

Case Western Reserve University Biomechanics Laboratory Standard Operating Procedure

Cutting Toe Region from Raw Data Version #1 Date: 07/14/09

Author: Stephanie J. Dux Principal Investigator: Christopher J. Hernandez

______Summary: This document outlines the protocol for cutting out the toe region out of the raw data from a compression test. Not all data files have a toe region so this SOP is only needs to be used for specimens that have a toe region ______Key Words: toe region, strain ______Materials: data files from Instron, Matlab. ______

1. Delete the first few data points from the specimen’s data file because this is noise. Delete the data points before the strain value begins to increase. This is the point where the test actually begins. 2. Graph data with xltension1 to produce a stress strain graph. 3. Inspect graph to determine if there is a toe region. See ASTM Standard Test Method for Compressive Properties of Rigid Cellular Plastics D 1621 – 04a for a description and examples of toe regions. 4. Visually examine the graph to determine about where the toe region ends. Use the data cursor function in Matlab to click on the point where you think the toe region ends and record the corresponding strain. It is important to underestimate the toe region at this stage. 5. Open the data file and delete all data points before the data point with the strain that you recorded. 6. Run xltension2 7. Examine the graph and see if the entire toe region has been cut off. Also examine the red line. This line uses the data to graph a line with the slope that Morgan's method has determined. This line and the data (blue line) should very closely

match (they should have the same slope and pretty much be on top of each other when the toe region has been eliminated). 8. If the toe region is visually still there or if the lines don't match delete some more data points. Delete about 5 at a time so as not to delete data that is outside the toe region. 9. Run xltension2 again, and repeat if necessary.

Note: See data files raw_data and trimmed_data for examples of this method.

Appendix XII: SOP – Clear Embedding Protocol for Cylindrical Bovine Cancellous Bone Specimens

CASE Western Reserve UNIVERSITY Musculoskeletal Mechanics Laboratory Standard Operating Procedure

Clear Embedding Protocol for Cylindrical Bovine Cancellous Bone Specimens

Version 2

06/03/09

Author: Stephanie Dux Principal Investigator: Christopher J. Hernandez

Summary This standard operating procedure (SOP) describes the embedding protocol for cylindrical bovine cancellous bone specimens. This SOP is adapted from the Calcein Staining and Clear Embedding Protocol for Rat Tail Caudal Vertebra Samples to Label Micro Cracks by Seetha R. Kummari. Key Words: n-butyl phthalate, methyl methacrylate, methyl salicylate

Materials: cassettes (Fisher Scientific # 15182706), Ethanol, Methyl Methacrylate (MMA, Aldrich Co, #M55909-2L), Methyl salicylate (Fisher Scientific, #S80082), 2oz glass bottles (Qorpak GLC-01611), distilled water, vacuum pump, beakers, 2L- amber bottles (we can use cleaned empty MMA bottle), Benzoyl Peroxide (Fisher Scientific, #B274-1LB), n-butyl phthalate (Fisher Scientific # D30-500), deionized water, Vacuum dessicator.

SAFETY: Handling of chemicals requires the user to wear splash resistant goggles, a lab coat and disposable gloves.

NOTE: When a bottle is opened label the bottle with the date opened.

1. Dehydration:

 Change the solution to 70% ethanol solution (70% ethanol, 30% distilled water) for 12 hours and store on shaker.  Change the solution to 70% ethanol solution for 12 hours and store on shaker.  Change the solution to 80% ethanol solution for 12 hours and store on shaker.  Change the solution to 80% ethanol solution for 12 hours and store on shaker.  Change the solution to 95% ethanol solution for 12 hours and store on shaker.  Change the solution to 95% ethanol solution for 12 hours and store on shaker.  Change the solution to 100% ethanol solution for 24 hours and store on shaker.  Change the solution to fresh 100% ethanol solution for 24 hours and store on shaker.  Change the solution to fresh 100% ethanol solution for 24 hours and store on shaker. Note: When the 100% ethanol solution is used, a new bottle of ethanol must be opened every time for every solution. Use the pint size bottles for this.

2. Clearing:

(Note: Handling of any type of MMA and Methyl Salicylate, must be in the Vacuum hood.)

1. Put the cassettes into bottle with 50:50, Methyl Salicylate:100% ethanol 2. Keep the bottle on the shaker for 24 hours. 3. Repeat steps 1 – 2 twice (2X24 hour washes) with 100% Methyl Salicylate.

3. preparation of Dry Benzoyl Peroxide and MMA solutions:

6.1 Preparation of dry Benzoyl Peroxide: Fill indicating desiccants (Drierite, #23005) into a desiccator. Put wet benzoyl Peroxide into a dish, and then put the dish into the desiccator. Change desiccants if they turn to pink (Originally they are light blue). When desiccants don’t change color any more, it means benzoyl peroxide is completely dry. Dry Benzoyl Peroxide is stored in the mini‐ desiccator in the hood.

(Note: Removing the inhibitor from the original MMA solution seems to make little difference in the rate of polymerization once the catalyst, benzoyl peroxide, is added.)

6.2 Preparation of MMA solutions:

1. MMA‐I: Follow the steps below to make 100 ml of MMA‐I solution.

 Take 85 ml of original MMA solution in a beaker.  Add 15 ml of n‐butyl phthalate to the solution.  Cover the beaker with plastic wrap to prevent evaporation.  Mix the solution of stir‐plate with heat off.  You can store this solution at room temperature. 2. MMA‐II: Follow the steps below to make 100 ml of MMA‐II solution.

 Take 100 ml of MMA‐I solution in a beaker.  Add 1 g dry benzoyl peroxide to the solution.  Cover the beaker with plastic wrap to prevent evaporation.  Mix on stir‐plate with heat off.  Store in a dark bottle in the refrigerator to prevent polymerization. 3. MMA‐III Follow the steps below to make 100 ml of MMA‐III solution.

 Take 100 ml of MMA‐I solution in a beaker.  Add 1.85 g dry benzoyl peroxide to the solution.  Cover the beaker with plastic wrap to prevent evaporation.  Mix on stir‐plate with heat off.  Store in a dark bottle in the refrigerator to prevent polymerization.

4. Infiltration with MMA solutions:

7.1 Infiltrate with MMA I:

1. Put Cassettes into the bottle with MMA I solution 2. Store the bottle under the hood for 48 hours.

7.2 Infiltrate with MMA II:

1. Put Cassettes into the bottle with freshly prepared MMA II solution (should be 20Xbones volume) 2. Store the bottle with cassettes in the refrigerator for 48 hours. 3. Drain MMA I into appropriate disposal vessel.

7.3 Infiltrate with MMA III:

1. Put cassettes into the bottle with freshly prepared MMA III solution (should be at least 20Xbones volume). 81

2. Store this bottle in the refrigerator for 48 hours. 3. Drain MMA II solution into appropriate aging containers

7.4 Embed in MMA III (detailed in Section 8)

(Note: Clear MMA will take about 7‐10 days to polymerize.)

5. Embedding process:

8.1. Preparation of Bases: 1. Make MMA III solution sufficient enough (20mL X # of bases) for the number of bases you are planning to make (Refer section 5.2 for the preparation of MMA solution) 2. Pour 20mL MMA III solution into 2oz Qorpak jar. 3. Keep these vials without caps with in vacuum bell jar. 4. Turn the vacuum on and allow vacuum to stabilize at 30 in-Hg. 5. Close valve on bell jar lid, turn vacuum off and seal off hose. 6. Let these bases sit under vacuum for 90 minutes. After each 30 min open the valve slowly and let air in to bring the system to normal pressure then reapply the vacuum at 30 in-Hg. Close the valve and seal off the hose until the next 30 min. 7. Put the caps on the vials and seal them as tightly as possible. Note: do no tighten too much the caps can crack. 8. Seal the caps with tape. 9. Keep these vials inside Isotemp with as much as gap possible between them. 10. Switch on the Isotemp and set the temperature to 320 C 11. To check on the sample to see if it has hardened, the vial is taken out of the Isotemp. Carefully, it is placed on a level surface and slowly open the cap. Once the cap is removed, the wooden end of a large sample Q‐tip is used to test the hardness of the MMA. If it has not hardened (the Q‐tip leaves a mark on the surface of the MMA) keep them back in the Isotemp with caps on them. 12. Remove vials from the Isotemp and switch it off once MMA is hardened. Clear MMA will take about 7 to 10 days to completely polymerize.(note: polymerization results in a slight ~10% shrinking of the base so you must take this into account with pouring MMA)

8.2 Embedding

1. Pour 2 to 5 mL of MMA I into the hardened bases. It should form a very thin layer of liquid on the top surface.

(This has to be done at least 2 hours prior to the plan of embedding process. Top surface of the bases should become gluey before we start the embedding process. Check the bases every hour to see if they are gluey. When they are gluey embedding can occur.)

2. Place the bone cylinder horizontally on the base. 3. Make sure bone is sticking to the surface. Otherwise move the bone sample little bit so that bone sticks to the gluey surface. 4. Fill out the bottle with MMA III up to the neck 5. Follow section 8.1 for vacuuming and rest of the embedding process. Note: Be very careful when transferring the jar from the vacuum chamber to the water bath. Jostling the jar could create air bubbles or knock the bone out of place.

Appendix XIII: SOP – Staining, Polishing and Mounting of Embedded Cylindrical Bovine Cancellous Bone Specimens

CASE WESTERN RESERVE UNIVERSITY Musculoskeletal Mechanics Laboratory Standard Operating Procedure

Sectioning, Polishing and Mounting of Embedded Cylindrical Bovine Cancellous Bone Specimens

Version 2 09/21/2009

Author: Stephanie Dux, Eileen Chu Principal Investigator: Christopher J. Hernandez

Summary: This standard operating procedure (SOP) describes the sectioning, polishing and mounting of embedded cylindrical bovine cancellous bone specimens. This SOP is based off the SOP Sectioning, Polishing, and Mounting of embedded rat tail caudal vertebra segment (C7+C8+C9) by Seetha R. Kummari.

Key Words: Sectioning, polishing, mounting, microscope

Materials: Eukitt Mounting Medium, Slides (Fisher Scientific 12‐544‐7), Cover slips (Fisher Scientific 12‐543‐D), Kimwipes, Small pipettes, popsicle sticks, Isomet, Bandsaw, Distilled water, 600 grit polishing paper (Buehler 30‐5118‐600‐102), 1200 grit polishing paper (Buehler 30‐5528‐012‐100), tweezers, ruler, sharpie, post it notes

SAFETY: Handling of chemicals requires the user to wear splash resistant goggles, a lab coat and disposable gloves.

I. Trimming the specimen:

1. Breaking the glass vial:  Wear glass resistant goggles and gloves. These are located in the Histology lab. 84

 Put the glass vial in absorbent underpad and wrap it such a that broken glass particles stay inside the underpad  Take a hammer and break the glass vial by slowly hitting on the underpad. You can choose to do this on the floor.  Once the glass is broken, take out the specimen and remove all the glass particles sticking on it.  Throw the broken glass pieces into the waste cardboard box which has a label "GLASS ONLY"

2. Trimming off the extra plastic:  Draw straight lines on the plastic where you want to trim the specimen. Draw it as shown in the figure below

Figure 20. The schematic shows how to trim off the excess methyl methacrylate so that thin sections can be made using the Isomet.

 Trim off the extra plastic by using vertical band saw machine. (Be careful when using the machine. When specimen is too thin, you might use two wooden pieces to guide the specimen. Wear safely goggles while operating the machine)

 The trimmed specimen should look like figure below

Figure 21. The above schematic shows how to mark the first cut on the trimmed embedded specimen.

 Using 600 grit polishing and distilled water, quickly polish the sides of the trimmed specimen.  Label the specimen  Draw a line parallel to the side of the bone as seen in the figure above.

II. Sectioning by using the Isomet

1. Set the blade on the Isomet. Thumb tight is enough for the final screw. 2. Pour water in the chuck such a way that isomet blade is about 5 to 10 mm inside the water 3. Fit the specimen in the jig provided for holding the specimen. Don't the tighten the screws 4. With a flat edge, see where your initial cut (rough cut) going to be. Make sure it is not well inside the bone. Make sure you are cutting the specimen parallel to the long axis of the bone. This can be accomplished by making sure that the drawn Sharpie line is parallel with the jig. 5. Check whether you can get through cuts with the current position. If you are not getting through cuts then adjust the inclination of the specimen so that you can get through cuts 6. Adjust the 'Autostop', so that you can get through. You can adjust this 'Autostop' even in between the cutting process. 7. Set the speed of the blade to the maximum and switch on the machine. 8. Put a weight on the top of the jig. (weights should be chosen according the speed of the cut we wanted). 9. Hold the jig with specimen and slowly drop it on to the blade. (Don't do it other way: i.e don't drop the specimen first and then start the blade. Always start the blade first and slowly drop the specimen onto the blade). 10. Once after getting initial/rough cut, don't disturb the alignment of the job and the blade. 86

11. To get next section, rotate the micrometer attached the machine, one complete turn +15 to get sections of about 250 microns a. Note: If you are using the large Isomet, use one of the blades that are either 400 or 450 microns thick. Rotate the wheel 0.8mm between sections. Also, the large Isomet cuts sections quicker than the smaller one. 12. Collect the section, label the section, and put it in the small ziploc bag. Label the bag properly with the specimen’s name. Sections should be labeled sequentially (1 through however many sections are cut for a specimen). 13. Once you are done sectioning the specimens, remove the blade and rinse with tap water. 14. Clean all the chucks on the machine.

III. Polishing

1. Adhere 600 grit sandpaper on the big granite block. (Make the granite surface little wet so that it would be easier to remove sand paper later.) 2. Adhere another piece of 600 grit sandpaper on the flat end of the metal block or plastic block (I like using the metal block for this step). 3. Place the sample on larger sandpaper with some distilled water. Samples must be wet all the time during polishing 4. Polish the sample between two sandpapers in smooth circular motion. 5. Regularly check the thickness of the sample with the special scale. (Scale which is mounted on small granite block) 6. Do the polishing till you reach about 110 microns 7. Repeat the steps 1‐5 with 1200 grit sandpaper till you reach 100 microns (I like to use the plastic block for this step). 8. When you have proper thickness the samples need 3 rinses (1 minute long each) in fresh distilled water within the ultrasonicator. Ultrasonicator must be filled with tap water, you then place a vessel within the ultrasonicator that is full of distilled water. Change distilled water for every wash. Hold the section with tweezers in the distilled water for each rinse. 9. After 3 rinses, place the sample on kimwipe paper to make the sample dry. Note: During all steps make sure each section is clearly labeled with the specimen and slide number. I write this information on a kimwipe and keep the section on the labeled kimwipe.

IV. Mounting:

NOTE: Mounting must occur under the hood. Gloves and splash proof goggles need to be worn.

1. Place the sample on micro‐slide facing letters up. 2. Label slide with specimen name and slide number. 87

3. Put many drops of EUKITT on the slide. There should be a puddle of EUKITT on the slide about the size of the polished section. 4. Using tweezers, place the polished section in the puddle of EUKITT on the slide. 5. Use a popsicle stick to gently push the EUKITT on top of the section until the entire section is covered with EUKITT. 6. Take a cover slip and try to put it on the sample starting from edge. Place the cover slip on section in one smooth motion. 7. Press the cover slip gently and wipe out extra EUKITT. Do this by dabbing the excess off the sides with a kimwipe or the chuck that is in the hood. If you see air bubbles in your specimen, press a bit firmer on the cover slip to rid of these excess bubbles and dab the sides of the slide again on the chuck. In the event that EUKITT gets on top of the cover slide, do NOT try to wipe it off with a kimwipe as this will leave streaks. Simply let the EUKITT dry clear. 8. Let the slide dry for 3 to 4 hours 9. Place slide in a slide case and label the slide case.

V. Making Eukitt Mounting Medium

1. Mix 15 ml of xylene and 85 ml Eukitt from the bottle. 2. Xylene evaporates over the time so be sure to put the cap on tightly after every use.

NOTE: The consistency should be thin enough so that if you stick a pipette or popsicle stick into the mixture and take it out, liquid drop should form quickly and drop down (within fraction of seconds ).

VI. Marking the Region of Interest

1. With a ruler and sharpie draw two lines that are parallel to the longitudinal axis and is 1mm in from both sides of the specimen.

2. With a ruler and sharpie mark the middle 7mm of the specimen.

3. Cut small strips of post it notes and cover each sharpie line with one.

Figure 22. A microscope slide without the region of interest marked is depicted in the left image. The orange area in the left image is the polished histological section of bovine cancellous bone. The region of interest was defined as a central rectangular area 7mm in the longitudinal direction and 6mm wide and is pictured in the image on the right.

Appendix XV: SOP – Microdamage Counting of Cancellous Bone

CASE WESTERN RESERVE UNIVERSITY Musculoskeletal Mechanics Laboratory Standard Operating Procedure

Microdamage Counting of Cancellous Bone

Version 1 09/22/2009

Author: Stephanie Dux Principal Investigator: Christopher J. Hernandez

Summary: This standard operating procedure (SOP) describes how to count microscopic tissue damage (or microdamage) in cancellous bone histological sections.

Key Words: microdamage, diffuse damage, cross hatching, microcracks, microfractures, bone volume fraction

Materials: Prepared microscope slides, microscope (Nikon Optishot‐2), UV filter (Chroma 11000 V2), eyepiece grid

I. Setting up the Microscope:

 Turn on the microscope.  Ensure that the eyepiece grid is in the eyepiece. If it is not in the eyepiece put it in the eyepiece.  Place microscope slide under the microscope.  Set to 100x magnification, turn the magnification to 10x (10x in eyepiece).  Move the microscope table and eyepiece so that the eyepiece grid is aligned with a corner of the region of interest on the microscope slide.

II. Examining the Region of Interest

 Examine the first area in the corner of the region of interest on the microscope slide. Bone volume fraction, diffuse damage, cross hatching, microcracks, and microfractures should be examined in the middle 8x8 points of the eyepiece grid (see Figure 23).  Once the region under the eyepiece grid is examined move the table (up, down, right, or left) so that the eyepiece grid overlaps where the previous eyepiece grid lay by one row

(or column depending on how you are moving the table). Before moving the table, pick a point on the microscope slide to line the eyepiece grid up with so that it overlaps where the previous eyepiece grid lay by one row. See Figure 24 for an example.  Continue to examine the area under the eyepiece grid and move the eyepiece grid until the entire region of interest on the microscope slide has been examined.

should then be picked so that the eyepiece grid can be moved down to the area surrounded by the blue outline. The blue box overlaps the red box by one row. The gray area inside the blue box should be examined. This process should be repeated and the eyepiece should be moved to the green box and then the purple box.

III. Point Counting

 Bone volume fraction, diffuse damage separate from cross hatching regions, and diffuse damage in cross hatching regions should all be calculated using point counting.  Examine the slide under the grid eyepiece and within the middle 8x8 points. Look for whatever you are looking to measure (bone volume fraction, diffuse damage separate from cross hatching regions, or diffuse damage in cross hatching regions). Anytime this intersects with one of the middle 8x8 points count that point (see Figure 25).  To calculate bone volume fraction all of the points that intersect with bone must be counted and all the points that intersect porous space must be counted. Bone volume fraction = # of bone points / (# of bone points + # of porous points).  To calculate diffuse damage separate from cross hatching regions all the points that intersect with diffuse damage separate from cross hatching regions must be counted. Diffuse damage separate from cross hatching regions = # of points of diffuse damage separate from cross hatching regions / # of bone points.  To calculate diffuse damage in cross hatching regions all the points that intersect with diffuse damage in cross hatching regions must be counted. Diffuse damage in cross hatching regions = # of points of diffuse damage in cross hatching regions / # of bone points.

Figure 25. The schematic above represents an example of point counting. The grid represents that eyepiece grid and the blue shape represents what we are interested in

counting. The red dots represent the points that we should count because they are in the internal 8x8 grid of the eyepiece and intersect with the blue shape.

IV. Diffuse Damage:

Diffuse damage is defined as areas of brightly colored post stain (calcein, green under UV excitation). Diffuse damage can extend to the surface of a trabecula, but must also be present in the internal region of a trabecula so as not to be confused with out‐of‐plane trabecular surfaces (see below). A region of diffuse damage does not have distinct edges, but have edges that gradually fade out to regions of no stain (Fazzalari, Forwood et al. 1998; Wang, Guyette et al. 2005; Kummari, Davis et al. 2009).

V. Cross Hatching

Cross hatched regions are an area with numerous small crossing cracks (Fazzalari, Forwood et al. 1998). The region of cross hatching caused by loading also shows a general amount of staining that can be characterized as diffuse damage (calcein stain under UV excitation). As a result, cross hatching was quantified both through point counting (as a form of diffuse damage) and by quantifying each crack within a cross hatched region (Arthur Moore and Gibson 2002). Microcracks in cross hatched regions must be linear, thin, sharp, and prominent enough to be easily distinguished from the other surrounding microcracks. The microcracks must be counted and the length of each microcrack must be measured.

VI. Microcracks and Microfractures

An individual microcrack (not part of a cross hatched region) is defined as an area of brightly colored calcein stain that is linear, thin, sharp line, and displays a halo of damage stain. The microcracks must be counted and the length of each microcrack must be measured. A microfracture is a large microcrack that extends completely through a trabecula.

VII. Distinguishing Microdamage from Out of Plane Surfaces and Vessels

Microscopic tissue damage was observed directly under a microscope rather than using digital images of the slide to prevent errors associated with vessels and oblique trabecular surfaces. Vessels within cancellous bone are easily mistaken for cracks, but can be distinguished by thickness (vessels are thicker), the presence of two distinct walls and/or rounded edges. Trabecular bone surfaces at an oblique angle within a 100 micron histological section may therefore appear as microdamage (diffuse damage, cross hatched regions, or microcracks). To prevent counts of oblique surfaces as microdamage, the optical plane was varied to confirm that an area of microdamage (diffuse damage, cross hatched regions, or microcrack) was not part of the trabecular surface viewed in cross‐section. When going in and out of focus, if the area of “microdamage” is always present than this is microdamage, if it disappears and reappears then this is an oblique surface.

Examples

Figure 30. The image above shows an example of a vessel (indicated by the arrow). We can determine that it is a vessel because there are two distinct walls.

Appendix XVI: Matlab Code for Plotting Stress – Strain Curves and Calculating Mechanical Properties (xltension2.m)

%modified from modycfmod.m copied from emorgan/hvb 9/01. GJK % %mody.m modified for HVB files of EFM from 8/97 %strain (estar) and load need to be in the columns listed in lines 20,21 %change signs in lines 20 and 21 for tension vs. compression %file opened in line 1 contains specimen data file names in column 1 %diameters in column 2 and lengths in column 3

%This code calculates mechanical properties from a compression test using %methods described in Morgan's 2001 paper. %This code is written for data where load is in Newtons and strain is in % clear all clc fid=fopen('xltensionTCa','r'); %cd X:\Personal Folders\Stephanie\Modulus\July152008 bigflag=0; k=1; while bigflag==0, in=fscanf(fid,'%s',1); %load in file and specimen dimensions di=fscanf(fid,'%s',1); length=fscanf(fid,'%s',1); diam=str2num(di); length=str2num(length); if in(1)=='E' | di(1)=='E', %when it reaches the end of the array bigflag=-1; end if bigflag==0, f=load (in); %f=eval(in) out=[in 'dat']; fid1=fopen(out,'w'); %will contain results csa=.25*pi*diam^2; %a=f(:,4)*25/length %strain=a-a(1) strain=-f(:,4); b=f(:,2)/csa;%divide load by cross-sectional area stress=-b; %i=1; %segment=f(:,5); %while segment(i)<6 % i=i+1; % %end % i=i-1 % strain(1:i)=[] % stress(1:i)=[]

%*********** INITIAL MODULUS *********** 96

ct=1; %lim=.2; lim=.2 + strain(1); %because our strain may not start at zero while strain(ct)

%***************** FINDING ZERO POINT FROM INITIAL MODULUS ********** % b=stress(1)-modinit*strain(1); % x=-b/modinit; % xoffset=x+.02;

%********** FINDING ZERO POINT FROM MAX LINEAR SLOPE ************* begin=1; increment=10; finishlim=.95; % [P,S]=polyfit(strain(begin:increment),stress(begin:increment),1); while strain(increment)

%************ LOOKING AT FIT ******** %w=1 %while w

% %********** YIELD, ELASTIC, AND ULTIMATE STRESSES AND STRAINS *********

ystrain=0; ystress=0; ct=1; while ystrain==0 if ystrain==0, if offset(ct)>stress(ct), ystress=stress(ct-1); ystrain=strain(ct-1);

end end

ct=ct+1; end % ct=1; % while stress(ct)>offset(ct) % ct=ct+1; % end % ct % ystrain=strain(ct); % ystress=stress(ct);

ultstress=0; ultstrain=0; [i,j]=max(stress); if strain(j)

postyieldstrain=max(strain)-ystrain; [r,s]=max(strain); Esecant=(stress(s)/(r-(2*xintercept)))*100

modulus=[modinit*100, ystrain-xintercept, ystress, ultstrain, ultstress, postyieldstrain];

fprintf(fid1,'%4.2f %4.6f %4.2f %4.6f %4.2f \n',modulus'); disp(modulus) status=fclose(fid1);

% %********** PLOTS *********

%subplot(3,2,k),plot(strain,stress, strain,(strain- xoffset)*modinit,strain,line) %plot(strain,stress, strain,(strain-xoffset)*modinit, ystrain, ystress,'o', xintercept, 0, 'o') subplot(3,2,k),plot(strain,stress,ystrain, ystress, 'o') %subplot(3,2,k),plot(strainplot,stressplot,strainplot,stressfit) k=k+1; axis([0 1.5 0 45]) 98

xlabel('Compressive Strain (%)') ylabel('Compressive Stress (MPa)') %legend(in,'offset','line', 'yield point') %legend(in,'offset') legend(in, 'yield point') end clear T end %end while loop

Appendix XVII: Stress – Strain Curves for Control Specimens

100

101

Appendix XVIII: Stress – Strain Curves for Irradiated Specimens

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Appendix XIX: Mechanical Properties Data

Control Specimens

Elastic Yield Yield Ultimate Ultimate Residual Post Yield Modulus Strain Stress Strain Stress Strain Strain (%) (MPa) (%) (MPa) (%) (MPa) (%)

PT7-9 3731.37 1.00 30.11 0.31 0.20

PT7-5 845.58 0.97 6.54 0.43 0.15

PT6-14 1500.44 0.97 11.55 0.38 0.16

PT16-8 1740.54 0.92 12.56 0.41 0.17

PT16-24 1444.06 0.90 10.16 0.44 0.18

PT15-7 2202.24 0.88 15.01 1.15 15.68 0.45 0.23

PT15-5A 3868.35 0.88 26.41 0.43 0.21

PT15-1A 6139.45 0.76 34.46 0.57 0.22

PT14-7 1188.26 0.90 8.32 0.46 0.17

PT14-4 2328.11 0.71 11.93 0.62 0.24

PT14-11 3559.17 0.73 18.86 0.60 0.19

PT18-5 1161.85 0.95 8.76 0.36 0.21

PT19-6 3325.62 1.12 30.58 0.14 0.20

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Irradiated Specimens

Elastic Yield Yield Ultimate Ultimate Post Yield Residual Modulus Strain Stress Strain Stress Strain Strain (MPa) (%) (MPa) (%) (MPa) (%) (%)

PT7-17 2956.80 0.79 17.58 0.43 0.18

PT6-11 2444.97 0.77 13.85 0.57 0.30

PT5-8 1242.42 0.91 8.83 0.48 0.20

PT16-9 2707.30 0.97 20.81 1.06 21.32 0.34 0.24

PT16-16 423.49 1.09 3.79 0.05 0.26

PT15-5B 2155.23 0.93 15.73 0.38 0.29

PT15-4 2254.50 0.85 14.65 0.48 0.26

PT15-10 2333.04 0.87 15.73 0.92 15.86 0.45 0.35

PT14-9 3286.23 0.94 24.50 0.36 0.20

PT14-3 5227.28 0.81 32.01 0.97 33.48 0.51 0.48

PT15-8 2134.57 1.04 18.00 0.11 0.16

PT17-7 920.39 0.99 7.34 0.30 0.28

PT19-15 871.75 1.10 7.89 0.08 0.23

PT19-16 2079.53 0.99 16.47 0.09 0.24

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Appendix XX: Microscopic Tissue Damage Data

Control Specimens

Diffuse Diffuse Damage Average Damage in Crack Separate # of # of Crack Cross Density from Cross Microfractures Microcracks Length Hatching ( mm2 ) Hatching ( m ) Regions Regions

PT7-9 0.00679 0.00533 0 29 0.79150 90.80

PT7-5 0.00385 0.00128 0 5 0.18049 66.67

PT6-14 0.00000 0.00118 0 4 0.13297 83.33

PT16-8 0.00315 0.00378 0 17 0.60247 74.51

PT16-24 0.00168 0.00337 0 26 0.82112 64.10

PT15-7 0.00000 0.00203 0 12 0.34264 61.11

PT15-5A 0.00125 0.00166 0 8 0.18705 83.33

PT15-1A 0.00000 0.00207 0 12 0.23271 91.66

PT14-7 0.00065 0.00259 0 12 0.43655 77.78

PT14-4 0.00093 0.00139 2 13 0.33918 64.10

PT14-11 0.00042 0.00501 0 31 0.72905 80.64

PT18-5 0.00118 0.00529 0 25 0.82762 100.00

PT19-6 0.00034 0.00000 0 0 0.00000 0.00

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Irradiated Specimens

Diffuse Diffuse Damage Average Damage in Crack Separate # of # of Crack Cross Density from Cross Microfractures Microcracks Length Hatching ( mm2 ) Hatching ( m ) Regions Regions

PT7-17 0.00585 0.00045 3 18 0.45549 92.29

PT6-11 0.00046 0.00046 0 3 0.07827 44.44

PT5-8 0.00058 0.00000 0 2 0.06571 33.33

PT16-9 0.00520 0.00142 1 23 0.61230 79.71

PT16-16 0.00000 0.00000 0 2 0.08425 83.33

PT15-5B 0.00050 0.00000 0 3 0.08366 111.11

PT15-4 0.00000 0.00118 1 9 0.19941 100.00

PT15-10 0.00000 0.00055 0 2 0.06212 83.33

PT14-9 0.00046 0.00325 1 20 0.52303 93.33

PT14-3 0.00953 0.00217 12 33 0.80432 91.92

PT15-8 0.00110 0.00275 0 10 0.31007 86.66

PT17-7 0.00468 0.00000 1 0 0.00000 0.00

PT19-15 0.00156 0.00078 0 3 0.13211 100.00

PT19-16 0.00668 0.00095 4 20 0.53701 80.00

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