Murine Metapodophalangeal Sesamoid Bone Mineralization: a Light and Electron Microscopy Study

MURINE METAPODOPHALANGEAL SESAMOID BONE MINERALIZATION: A LIGHT AND ELECTRON MICROSCOPY STUDY

A thesis submitted to Kent State University in partial fulfillment of the requirements for the degree of Master of Arts

Alison R. H. Doherty

December, 2007 Thesis written by Alison R. H. Doherty B.A., University of Wyoming, 2003 M.A., Kent State University, 2007

Approved by

______William J. Landis, Ph.D, Advisor

______C. Owen Lovejoy, Ph.D, Co-Advisor

______Richard S. Meindl, Ph.D, Chair, Department of Anthropology

______John R. Stalvey, Ph.D, Dean, College of Arts and Sciences

Table of Contents

List of Figures ...... iv List of Tables...... v Acknowledgments ...... vi Introduction ...... 1 Literature Review ...... 2 Vesicle-Mediated Mineralization ...... 3 Collagen-Mediated Mineralization ...... 5 Sesamoid Bones...... 7 Fibrocartilage...... 9 Materials and Methods ...... 12 Sample Acquisition ...... 12 Radiography ...... 13 Histology ...... 13 Whole Mount Staining ...... 20 Transmission Electron Microscopy Sample Determination ...... 23 Transmission Electron Microscopy ...... 23 Results ...... 27 Radiography ...... 27 Histology ...... 29 Whole Mount Staining ...... 38 Transmission Electron Microscopy Sample Determination ...... 45 Transmission Electron Microscopy ...... 46 Discussion ...... 54 Conclusion ...... 60 Future Research ...... 61 Literature Cited ...... 64 Appendix ...... 69

iii List of Figures

Figure 1: Murine Front and Hind Paw Radiographs ...... 28 Figure 2: Murine Metatarsophalangeal Joint Anatomy ...... 30 Figure 3: Two-Week-Old Metatarsophalangeal Sesamoid ...... 31 Figure 4: Two-Week-Old Metatarsophalangeal Sesamoid Under Polarized Light ...... 32 Figure 5: Metatarsophalangeal Sesamoid Fibrocartilaginous Insertion Site ...... 33 Figure 6: Proteoglycan Content of the One-Week Metatarsophalangeal Sesamoid ...... 34 Figure 7: One-Week-Old Metacarpophalangeal Sesamoid ...... 35 Figure 8: One-Week-Old Metatarsophalangeal Sesamoid ...... 35 Figure 9: Two-Week-Old Metacarpophalangeal Sesamoid ...... 36 Figure 10: Three-Week-Old Metacarpophalangeal Sesamoid ...... 37 Figure 11: Three-Week-Old Metatarsophalangeal Sesamoid ...... 37 Figure 12: Whole Mount Double-Stained One-Week-Old Mouse ...... 39 Figure 13: One-Week-Old Alizarin Red S Stained Front and Hind Paws...... 40 Figure 14: Double-Stained One-Week-Old Hind Paw ...... 41 Figure 15: The First Mineralized Sesamoid ...... 42 Figure 16: Secondary Ossification Centers in the One-Week-Old Hind Paw ...... 43 Figure 17: Three-Week-Old Alizarin Red S Stained Front and Hind Paws ...... 44 Figure 18: Thick Section of the Two-Week-Old Metacarpophalangeal Sesamoid...... 47 Figure 19: Matrix Vesicles in the Extracellular Space between Cells ...... 48 Figure 20: Matrix Vesicles near Chondrocytes ...... 49 Figure 21: Chondrocytes within Calcifying Cartilage ...... 50 Figure 22: Perpendicular Collagen Fiber Orientation ...... 51 Figure 23: Areas of Parallel Collagen Fiber Orientation ...... 51 Figure 24: Thick Section of the Three-Week-Old Metacarpophalangeal Sesamoid ...... 52

iv List of Tables

Table 1: Calcium Oxalate Endpoint Test ...... 14 14 Table 2: Tissue Processing ...... 15 15 Table 3: Toluidine Blue Stain ...... 17 17 Table 4: Picrosirius Red Stain ...... 18 18 Table 5: Safranin-O Red Stain ...... 19 19 Table 6: Whole Mount Staining ...... 22 22 Table 7: TEM Processing and Embedding ...... 24 24 Table 8: Formvar Coated Grids ...... 25 25 Table 9: Positive-Contrast Staining Grids ...... 26 26

v Acknowledgments

I express my deepest gratitude to my loving and supporting husband, Adam

Doherty, for standing behind me all the way in this endeavor. I could not have made it without his encouragement and willingness to edit. I am also very appreciative to

Elizabeth Lowder and the training she gave me in all aspects of histology, radiography, and general laboratory procedures. When I ran into roadblocks, Beth was always my brainstorming partner and backup plan expert.

Special thanks go to my advisor, Dr. William Landis, for his guidance and suggestions in this research project, as well as to Dr. Owen Lovejoy and Dr. Richard

Meindl for their support. In addition, this project would not have been feasible without the training and expertise of Jeanette Killius, the NEOUCOM Electron Microscopy

Laboratory Coordinator.

Finally, I thank Dr. Hans Thewissen, Dr. Chris Vinyard, Robin Jacquet, Janet

Hamilton, my fellow graduate colleagues, and my parents for their role in supporting this thesis project. Without funding from the NEOUCOM Skeletal Biology Focus Area and the Graduate Student Senate this project would not have been possible.

Introduction

Extracellular matrix vesicles have been documented as initiating mineralization of many vertebrate calcified structures, including calcifying cartilage, tendon, fibrocartilage, and bone (Ali et al., 1970; Hsu and Anderson, 1978; Landis, 1986; Lowenstam and

Weiner, 1989; Yamada, 1976). In calcifying cartilage and bone, mineral crystal growth first occurs in matrix vesicles and then quickly engulfs the surrounding organic matrix.

However, vesicles in normally mineralized turkey tendons do not appear to be the only site of mineral formation. In these tendons, collagen initiates mineral growth in areas isolated from matrix vesicles, but these nucleation events always follow vesicle presence in the tissue. Therefore, the role of matrix vesicles in initiating mineralization is still under investigation in vertebrate tissues (Boskey, 1998; Landis et al., 1996; Lowenstam and Weiner, 1989).

Sesamoid bones may provide a unique opportunity to observe matrix vesicles.

These small structures are highly conserved in vertebrates, develop within and are intimately associated with tendon, and yet are present as cartilaginous models in the fetus. The focus of this research is to test the hypothesis that matrix vesicles are responsible for initiating calcification in murine metapodophalangeal sesamoid bones

(found at the joint between the metapodia and proximal phalanges). Matrix vesicle

presence, their potential role in initial mineral formation, and their association with collagen fibers will be investigated using transmission electron microscopy (TEM).

Normally developing CD-1 mice will provide a readily available model to examine the development and mineralization of metapodophalangeal sesamoids to test the hypothesis of this study. In order to determine matrix vesicle presence, it is first necessary to obtain documentation of the development of murine sesamoid bones using radiography, light microscopy, and whole mount staining. These methods can provide initial data regarding the basic anatomy, morphology, cellular arrangement, and mineralization timeline of the sesamoids for further investigation. The developmental stages determined most likely to contain matrix vesicles will be examined by TEM for vesicle presence and function in sesamoid mineralization. All data collected using light and electron microscopy will be compared to the developmental process of calcifying tendon, fibrocartilage, and cartilage already documented in the literature.

Literature Review

The source and location of initial mineral formation in vertebrate calcified tissues are currently being investigated. Extracellular matrix vesicles have been suggested as initiating mineral formation before any other structure in the organic matrix of calcifying cartilage (Ali et al., 1970; Gerstenfeld et al., 1998; Hsu and Anderson, 1978; Kirsch et al., 1997; Plate et al., 1996), fibrocartilage (Yamada, 1976), predentin (Pecile and de

Bernard, 1990), fish scales (Lowenstam and Weiner, 1989), antler (Szuwartz et al.,

1998), and other vertebrate mineralizing tissues. However, recent evidence suggests that collagen molecules spatially isolated from matrix vesicles can initiate mineralization particularly in calcifying tendon (Landis and Song, 1991; Landis et al., 1993; Landis et al., 1996; Landis and Silver, 2002; Siperko and Landis, 2001). Therefore, a new and important question in the field of biomineralization is whether vesicle-mediated mineralization always precedes collagen-mediated mineralization or if the two processes can occur concurrently (Boskey, 1998; Landis et al., 1996; Lowenstam and Weiner,

1989).

Vesicle-Mediated Mineralization

Found within vacant spaces of the extracellular matrix of many vertebrate mineralizing tissues, matrix vesicles are globular, membrane-bound structures with diameters ranging from 60-250 nm. Matrix vesicles can only be identified using transmission electron microscopy (Ali et al., 1970; Landis, 1986, 1989). Analyses conducted on the trilaminar membrane of matrix vesicles within calcifying cartilage indicate that they are derived from the surrounding chondrocytes, yet they are distinct from intracellular organelles (such as lysosomes) in their extracellular location and enzyme composition (Ali et al., 1970; Anderson, 1969; Lowenstam and Weiner, 1989).

Containing multimolecular complexes thought to be transmembrane ion transporters, the insoluble membrane of matrix vesicles appears to facilitate mineralization (Kirsch et al.,

1997; Lowenstam and Weiner, 1989).

The membrane of matrix vesicles has several important functions. It defines an enclosed space within the vesicle suggested to confine and protect the initial mineral nucleus in a designated volume away from the surrounding extracellular fluid (Ali et al.,

1970). The insoluble membrane, in association with the embedded transport molecules, also provides a means of controlling the composition of the ions, such as calcium and

+2 -3 phosphate (Ca /PO4 ), transported and/or diffused into the vesicle (Lowenstam and

Weiner, 1989). In addition, alkaline phosphatase, an enzyme within the vesicle, may trigger the accumulation, supersaturation, and growth of mineral ions (Ali et al., 1970).

Nucleation events leading to initial mineral formation have been observed within the matrix vesicles as well as on the surface of their membranes (Hsu and Anderson,

1978). The first detectable mineral in most vertebrate calcified tissues is hydroxyapatite

(Ca10 (PO4)6 (OH)2), which composes 80% of the mineral phase in initial cartilage

+2 -3 calcification (Ali et al., 1970; Hsu and Anderson, 1978). In addition, Ca and PO4 , the two principal ions in vertebrate biomineralization, are stored and concentrated in matrix vesicles (Hsu and Anderson, 1978; Lowenstam and Weiner, 1989). Nucleation events mediated by matrix vesicles result in the growth of mineral as small platelets measuring

40-170 nm in length, 30-45 nm in width, and 4-6 nm in thickness (Landis and Silver,

2002).

In calcifying cartilage, numerous matrix vesicles are present in the longitudinal spaces between the columnar arrangement of chondrocytes within the epiphyseal growth plates (Anderson, 1969). As mineral crystals grow within the matrix vesicles, they are thought eventually to rupture the limiting trilaminar membranes (Kirsch et al., 1997;

Lowenstam and Weiner, 1989). The calcification of the surrounding extracellular matrix proceeds as many mineralization centers (the ruptured matrix vesicles) coalesce. The vesicles and other extracellular molecules (such as collagen) appear to be obliterated by mineral following vesicle membrane rupture.

It is believed that once the minerals escape the vesicles, some ions (such as Ca+2

-3 and PO4 ) resolubilize and diffuse to initiate calcification at nearby sites, such as on collagen fibers (Hsu and Anderson, 1978; Lowenstam and Weiner, 1989). However, it is very difficult to determine the origin of such diffusible ions, and it is unknown if matrix vesicles are solely responsible for initiating collagen-mediated mineralization. This is the heart of the current discussion surrounding the initial process of calcification in vertebrate tissues (Boskey, 1998; Lowenstam and Weiner, 1989).

Collagen-Mediated Mineralization

Recent evidence suggests that collagen fibers in the calcifying matrix of turkey tendons initiate mineralization in the absence of matrix vesicles (Gupta et al., 2003;

Landis and Song, 1991; Landis et al., 1993; Landis et al., 1996). Normally calcifying turkey tendon is unique in that it provides an animal model in which the progression of mineralization by both vesicles and collagen may be detected and assessed in a spatial and temporal sequence (Landis and Song, 1991). Linear arrays of numerous matrix vesicles, located between the longitudinal columns of spindle-shaped fibroblast cells within the tendon, temporally precede collagen nucleation events (Landis, 1986).

Subsequently, collagen fibers spatially isolated from matrix vesicles appear to form

mineral independently (Landis, 1986; Landis and Song, 1991; Landis et al., 1993).

Therefore, vesicle- and collagen-mediated mineralization can persist simultaneously in normally calcified tendon in the turkey leg.

There are many types of collagen, but mineralized vertebrate tissues are predominately composed of type I (bone, tendon, dentin, and cementum) and type II

(calcifying cartilage and fibrocartilage) collagen (Alberts et al., 2002). These fibril- forming proteins are synthesized and secreted from cells composing the respective tissues. Unlike vesicles, collagen is a triple helical, rod-shaped protein that requires further modification at the plasma membrane of the cell before it is functional (Alberts et al., 2002). The mature collagen fiber has a characteristic periodic banding pattern (64-70 nm), representing the hole and overlap zones in the arrangement of collagen molecules and fibrils (Hodge and Petruska, 1963; Landis and Song, 1991).

Two apparently independent nucleation events associated with collagen fibers have been described from studies of calcifying tendon (Landis and Song, 1991; Landis et al., 1993; Landis et al., 1996). One defines mineral crystal nucleation on the surface of fibers. These crystals develop into thin plates that eventually coalesce. In addition, mineral formation occurs within the periodic hole and overlap zones of the fibers. The mineral crystals that form associated with collagen, of similar initial dimensions of those reported for matrix vesicles, orient themselves parallel to each other and to the long axis of the fibers themselves. The surface mineral crystals and those that are growing from inside the fibers eventually form a continuous mineralized structure with their associated

collagen units, thereby calcifying the extracellular matrix of tendon (Landis, 1986;

Landis and Song, 1991; Landis et al., 1993; Landis et al., 1996).

Sesamoid Bones

Sesamoid bones are “skeletal elements that develop within a continuous band of regular dense connective tissue (tendon or ligament) adjacent to an articulation or joint”

(Vickaryous and Olson, 2007). Often, and incorrectly, referred to as accessory bones, sesamoids develop from a cartilaginous anlage like much of the rest of the skeleton

(Bland and Ashhurst, 1997; Bizarro, 1921; Gray, 1918; Joseph, 1951; Le Minor, 1988;

Sarin and Carter, 2000). In addition, sesamoids, like other bones of the skeleton, are genetically inherited and are found in most extinct and extant vertebrates, although in variant arrangements/numbers.

In addition to being considered accessory bones in much of the literature, sesamoids are often overlooked in research. Few studies have been conducted on the development of sesamoids, excluding that of the patella (Bland and Ashhurst, 1997;

Reese et al., 2001; Sarin and Carter, 2000; Walmsley, 1940). In early patellar development, a cluster of cells begins to develop within the patellar tendon shortly after the epiphyses of the femur and tibia are established. The cell cluster grows from the deep surface of the tendon into a hyaline mass that forms a cartilaginous model from which bone develops (Vickaryous and Olson, 2007). Centers of ossification quickly replace the cartilage, and the sesamoid grades through a transition of fibrocartilage where the tendon inserts on the bone (Clark and Stechschulte, 1998).

The placement of the sesamoid (e.g. patella) embedded within the deep surface of the tendon and in close association with the femur and tibia at the joint is the defining trait of the majority of sesamoids (Bizarro, 1921). In addition, sesamoids are usually found within the tendon or ligament near a muscle insertion and where forces are considered the greatest (Bizarro, 1921; Vickaryous and Olson, 2007). These bones are thought to modify stresses, reduce friction, and alter the direction of the pull of a muscle at a joint (Bizarro, 1921; Gray, 1918; Sarin et al., 1999). Although several studies have suggested that mechanical forces dictate sesamoid development (Benjamin and Ralphs,

1997; Sarin et al., 1999; Vickaryous and Olson, 2007), the normal occurrence of these small bones is ultimately reliant on genetically inherited pattern formation and positional information of cells directing the fetal development of the skeleton (Karaplis, 2002).

The oldest examples of sesamoids date to 295-250 m.y.a. and are associated with captorhinids, an extinct form of lizard-like reptiles, found at the joints between the metacarpophalangeal and interphalangeal joints (Vickaryous and Olson, 2007). The metapodophalangeal sesamoids are also some of the most common in mammals, like the patella (Le Minor, 1988; Vickaryous and Olson, 2007). These bones are found in pairs at the joints of the metapodia and proximal phalanges closely associated with the flexor digiti tendons along the palmar/plantar surface of the autopod. The microscopic and ultrastructural anatomy and development have not been thoroughly documented like those features of the patella. The assumption that the patella represents all sesamoid development may be erroneous (Lewis, 1977). It is the specific focus of this research to

investigate the mineralization and the role of the structures involved in metapodophalangeal sesamoid bone mineralization.

Fibrocartilage

The metapodophalangeal sesamoids, similar to other bones, have a large fibrocartilaginous insertion site (also called an enthesis) in the transition from the flexor digiti tendons to bone (Vickaryous and Olson, 2007). The four zones composing an enthesis (tendon, fibrocartilage, calcified fibrocartilage, and bone) make this an ideal candidate for the study of mineralization. Fibrocartilage has many similarities with both tendon and cartilage, and it mineralizes normally and pathologically (Benjamin and

Ralphs, 1998). It is possible that the fibrocartilaginous region associated with the sesamoid could contribute to the mineralization process of the bone (Benjamin and

Evans, 1990).

Fibrocartilage, like tendon, contains fibroblast cells. However, the cells in this tissue can be ovoid/round, have additional cell extensions, and often are associated with distinct lacunae (Bancroft and Gamble, 2002; Ham, 1969; Leeson et al., 1985). These fibroblasts transition from those found in tendon into the chondrocytes composing the hyaline cartilage of the patella in a fashion similar to that described for columnar chondrocytes in the epiphyseal growth plates. It is not possible to differentiate between the cells of tendon, fibrocartilage, and cartilage near the transition zones. However, where distinction can be made between the two types of fibroblasts, the flat, spindle- shaped fibroblasts associated with tendon will be referred to as tenocytes (Landis, 1986),

and the ovoid/round cells in fibrocartilage will be identified as fibrocartilage cells

(Benjamin et al., 1995) in order to differentiate between the cells of these two tissues.

Composed of about 90% type I and 5% type II collagen, fibrocartilage is found in areas where tensile strength and tough support are required (Benjamin and Evans, 1990;

Leeson et al., 1985). Collagen fibers are generally oriented parallel to the direction of the loading force, like those seen in tendon (Vickaryous and Olson, 2007; Bancroft and

Gamble, 2002). However, collagen arrangement near the bone within a fibrocartilaginous insertion has been described as an interweaving or basket-weave network similar to that found in cartilage (Benjamin and Evans, 1990; Rufai et al., 1996).

Not as stiff as hyaline cartilage and having less tensile strength than tendon, fibrocartilage displays mechanical properties of both tissues to maintain the integrity of the joint during loading at a transitional attachment site.

Although intensively studied at the microscopic level, little work has been done to document the ultrastructure of fibrocartilage (Rufai et al., 1996), and only two studies have explicitly documented the structures involved in the mineralization of this tissue

(Cooper and Misol, 1970; Yamada, 1976). Cooper and Misol (1970) reported smooth membrane-bound vesicles near and within the fibrocartilage cells. However, no recognition was given to any mineral nucleation events associated with the vesicles. Two types of collagen mineralization events were documented proceeding from the fibrocartilage zone into the calcified cartilage zone. The first mineral formed between the collagen fibers as electron dense regions. The second phase of mineral was reported to occur on the surface of and within the collagen, parallel to fiber orientation.

In a study of the rat Achilles tendon enthesis, Yamada (1976) described a mineralization process resembling that seen in the calcifying turkey tendon. Matrix vesicles were reported as the initial mediator of mineral formation in the spaces between the longitudinally arranged fibrocartilage cells. The closer to the tidemark, or zone of calcifying fibrocartilage (Benjamin et al., 1986), the number of matrix vesicles increased.

Soon after the rupture of the trilaminar vesicle membrane, mineralization of the surfaces of collagen fibers began. However, no mention was made concerning isolated collagen nucleation events distanced from the matrix vesicles.

It is thus apparent that the fibrocartilaginous insertion sites require additional ultrastructural analyses in order to understand fully the mineralization process in this tissue. The enthesis associated with the flexor digiti tendons and metapodophalangeal sesamoid bones is a prime area for research. The investigation of the relationship between fibrocartilage and sesamoid mineralization may provide a deeper understanding of the structures involved in the calcification process of vertebrate tissues in general.

Materials and Methods

Sample Acquisition

Charles River Laboratories CD-1 mice were bred and their litters raised for examining metapodophalangeal sesamoid bones (Wilmington, MA). To examine the process of mineralization in these bones, the sesamoids were observed before mineral growth began, as mineral developed, and after complete calcification. According to cleared and Alizarin red-stained samples reported by Wirtschafter (1960), the metapodophalangeal sesamoids begin to mineralize in C3H mice at one week of age.

However, Patton and Kaufman (1995) found no appearance of such sesamoid mineralization in cleared and double stained (Alcian blue/Alizarin red S) two-week-old hybrid (C57BL x CBA) mice. The onset and presence of mineralization may be strain- dependent in laboratory animals, which may account for the reported source of variation in the calcification process of mice (Patton and Kaufman, 1995; Spark and Dawson,

1928). In addition to strain, varying results have been published pertaining to factors such as sex and litter size that may affect skeletal development (Atchley and Hall, 1991;

Spark and Dawson, 1928). Information is absent in the literature regarding most sesamoid bones, including that of the metapodophalangeal sesamoid bones. The CD-1 mice used for this study were examined from one to six weeks of age, regardless of sex or litter size, to ensure an accurate examination of the mineralization of these bones.

To investigate sesamoid development each week, three mouse pups were euthanized in a carbon dioxide chamber, and their front and hind feet were dissected.

The feet from one mouse were fixed and prepared for radiography and subsequently for whole mount staining. The left feet from the other two mice were fixed and prepared for histology, and the right feet of the same mice were fixed for transmission electron microscopy.

Radiography

The feet dissected from each age group of mice for radiography were fixed in

95% ethyl alcohol (EtOH) for a period of two to four days, depending on the age of the mouse. Each front and hind paw was placed palm down with the digits spread gently on a paraffin block. Radiographs were taken using a mammography radiation unit (GE

Lorad M-II, Bedford, MA), operated at 25 kvp for 0.2 seconds at 2.4 mA. Scientific imaging film was used to obtain radiographs (Kodak X-OMAT LS, Rochester, NY), which were developed using an automatic x-ray film processor (Fischer Industries Futura

3000 S, Geneva, IL). The radiographs were then scanned at 1600 dpi in order to magnify and examine the metapodophalangeal joints (Epson Expression 1600, Long Beach, CA).

Histology

The left front and hind paws dissected from two mice of each age group were fixed in 10% neutral buffered formalin for 24 hours (the corresponding right paws from

these mice were fixed for electron microscopy as described below). After fixation, the paws were decalcified two to eight weeks in a solution of 5.5% EDTA in 10% neutral formalin (Appendix, Solution1) and continually agitated at room temperature (Bancroft and Gamble, 2002; Brown, 1978). Although removing the mineral content of the samples, decalcification was necessary in order to section the digits for histological study.

The samples were tested weekly using a calcium oxalate endpoint test to determine the remaining calcium content (Table 1). Decalcification was considered complete when no mineral precipitation formed 30 minutes after adding 5% ammonium oxalate (Appendix,

Solution 2) to 5mL of the used sample EDTA/formalin solution.

Table 1: Calcium Oxalate End Point Test (Bancroft & Gamble, 2002; Brown, 1978) 5 mL of the used sample EDTA fluid 5 mL 3% ammonium oxalate pH the fluid using litmus paper Add 6 N HCl until the solution is between 2 and 3 Add 5 mL of 3% ammonium oxalate Swirl the solution to mix Check the solution after 30 minutes If clear: Decalcification is complete Rinse samples in PBS x2 for 15 minutes Store in 70% EtOH until processed Cloudy precipitate: Insufficiently decalcified Rinse in PBS x2 for 15 minutes Put back into EDTA/Formalin for another week *Check samples every week to ensure proper decalcification

After decalcification, the samples were processed in an Autotechnician Mono

Processor (Tarrytown, NY) (Table 2). The paws were then infiltrated with paraffin using a vacuum infiltration oven at 60º C and 15 kPa of Hg for 30 minutes (National Appliance,

Skokie, IL), and they were then embedded in paraffin (Paraplast Xtra, Houston, TX).

The fifth digit of each paw was positioned medial side down in the embedding mold in order to obtain the proper orientation of the sample for sectioning on a microtome

(American Optical 820 rotary microtome, Buffalo, NY).

Table 2: Tissue Processing Set the Autotechnicon Mono tissue processor for a 12 hour cycle. Tissues will pass through the following series of solutions: Two 75% EtOH, 1 hour each Two 95% EtOH, 1 hour each Two 100% EtOH, 1 hour each 100% EtOH/xylene solution, 1 hour Two 100% xylene, 1 hour each Two Paraplast PLUS paraffins, 1.5 hours each At the end of the cycle, remove the samples to a vaccum infiltration oven Infiltrate the samples for 30 minutes or longer Embed samples upon removal from the infiltration oven

The fifth digit of each paw prepared for light microscopy was sectioned in a medial to lateral fashion at 7μm thickness. This orientation during sectioning provided the best view of the sesamoid in relation to its associated tendon, metapodia, and proximal phalanx. The sections were floated on a bath of distilled water and agar (Fisher

Tissue Prep, Pittsburgh, PA). Distilled essential oil was used as necessary to allow the sections to flatten and release the tissue (Histoclear, Atlanta, GA). The sections were then collected onto slides and allowed to dry.

The paraffinized thick sections were stained using toluidine blue and Picrosirius red (Appendix, Solution 3 and 4). Toluidine blue stains connective tissue mucins and ground substances of cartilage (Clark, 1981), accentuating the general morphology of a sample (Table 3). Picrosirius red stains collagen fibers, which are birefringent under polarized light (Table 4). Birefringence is of specific interest in illustrating the relation

of the sesamoids to their surrounding tendons, ligaments, and other bones. In addition, the one-week-old samples were stained with a Safranin-O red stain (Appendix, Solution

5), in order to view the proteoglycan content of the sesamoid (Table 5).

These stains made it possible to examine the metapodophalangeal sesamoids in relation to their surrounding bones and tendons using a light microscope and magnification of 40, 100, and 400x (Olympus Optical Ix70-S1F2, Japan). In addition, development of the sesamoids was compared in the corresponding front and hind paws of the same mouse, and to those of the other age groups of mice. Finally, each histological sample was examined for the presence of a mineralizing front in the sesamoids, indicative of matrix vesicles similar to that seen in calcifying cartilage, fibrocartilage, or tendon.

Table 3: Toluidine Blue Stain (Clark, 1973) Place slides in oven at 58º C for at least 30 minutes to deparaffinize the sections Dewax the slides: Three changes of 100% xylene, 5 minutes each 100% EtOH, 2 minutes 95% EtOH, 2 minutes Two changes of 75% EtOH, 2 minutes each

Distilled water rinse (dH2O), 2 minutes Stain: Place in freshly filtered 0.1% toluidine blue solution for 1 minute Dip slides 5 times in dH2O Dehydrate: Two changes of 95% EtOH, 2 minutes each Two changes of 100% EtOH, 2 minutes each Two changes of 100% xylene, 5 minutes each Mount the slides in Permount The stain can be reused Results: Acidic carbohydrates: Metachromatic (purple) Nuclei and cytoplasm: Orthochromatic (blue)

Table 4: Picrosirius Red Stain (Puchtler et al, 1973; Junqueira et al, 1979) Place slides in oven at 58º C for at least 30 minutes to deparaffinize the sections Dewax the slides: Three changes of 100% xylene, 5 minutes each 100% EtOH, 2 minutes 95% EtOH, 2 minutes Two changes of 75% EtOH, 2 minutes each dH2O rinse, 2 minutes Stain: Place in Picrosirius red for 1 hour Wash in two changes of acidified water, 2 minutes each Physically remove most of the water from the slides by vigorously shaking Dehydrate: Three changes of 100% EtOH, 2 minutes each Three changes of 100% xylene, 5 minutes each Mount the slides in Permount The stain can be reused Results: Collagen: Red Nuclei: Black, grey, or brown Background: Yellow

Table 5: Safranin-O Red Stain Place slides in oven at 58º C for at least 30 minutes to deparaffinize the sections Dewax the slides: Three changes of 100% xylene, 5 minutes each 100% EtOH, 2 minutes 95% EtOH, 2 minutes 75% EtOH, 2 minutes Distilled water rinse, 2 minutes Stain: Stain the nuclei with Weigert's iron hematoxylin, 7 minutes

Rinse in dH2O, 5 minutes Stain with 1% light green working solution, 3 minutes Rinse in 1% acetic acid, 1 minute

Rinse in dH2O, 10 dips Stain with 0.1% Safranin-O, 6 minutes Rinse in 1% acetic acid, 1 minute Dehydrate: Dip two times in two different 75% EtOH rinses Two changes of 95% EtOH, 1 minute each Two changes of 100% EtOH, 1 minute each Two changes of 100% xylene, 5 minutes each Mount the slides in Permount Results: Proteoglycans: Red

Whole Mount Staining

Three types of clearing and staining methods were performed on the fore- and hind-paws of the mice (Table 6). After several trial runs using various protocols, a modified clear and double-stain method (Wassersug, 1976), using Alizarin red S to stain calcium deposits and Alcian blue to stain glycosaminoglycans (Bancroft and Gamble,

2002), was used on the one- and two-week-old samples (Appendix, Solution 6). These stains were used to identify the presence of the cartilaginous sesamoids and the epiphyseal plates of their associated metapodia and phalanges. However, a modified clear and single-stain protocol using Alizarin red S proved to be a more reliable and efficient method to view the mineralizing bone of all age groups (Clark, 1981). The method of final clearing and glycerine infiltration of the tissues was adapted from

Kaufman (1992) and applied to the two protocols above.

After dissection from the mouse, the samples were skinned as much as possible and fixed according to the clear and stain protocol (Table 6). The one- and two-week-old paws selected for double staining using the Wassersug methodology (1976) were fixed in

10% neutral buffered formalin for 24 hours. Samples of this protocol were rinsed first in water and then in a 40% acetic acid/alcohol solution in preparation for the staining process. A few drops of 0.1% Alcian blue were added to a fresh acetic acid/alcohol solution for a period of 12 to 48 hours to stain the cartilaginous regions in the samples.

When the desired level of staining was established in the cartilaginous regions, the

samples were rinsed in 100% EtOH and placed in 1% potassium hydroxide (KOH) for about 12 hours (or until the bones were clearly visible through the soft tissue).

Paws from all sample age groups were fixed and prepared for single-staining following the protocol described by Clark (1981). Fixation in 95% EtOH for two to four days was immediately followed by clearing in 1% KOH as described above for the double-stained samples. When bones were visible through the soft tissues, staining proceeded for all samples utilizing the Clark methodology (1981). One drop of 0.1%

Alizarin red S was added to the fresh 1% KOH solution of each sample. Once the bones were stained sufficiently, they were put into fresh KOH for another 24 to 48 hours to continue the clearing process.

Complete clearing and glycerine infiltration of the samples was conducted utilizing the clear and stain protocol established by Kaufman (1992) to achieve the best results without over-maceration of the tissues. The feet were put into successive concentrations of 1% KOH and glycerine (20%, 50%, 80%) to complete the clearing procedure. This process took up to two weeks for some samples. Under a dissecting microscope, any remaining soft tissue (such as skin or muscle) was removed as necessary at each clearing stage. However, some tissue (especially that associated with the younger samples) was not dissected in order to preserve the integrity of the sample. Finally, the samples were infiltrated in 100% glycerine for 24 hours and stored in fresh 100% glycerine. The metapodophalangeal sesamoids were best viewed under a dissection microscope between 2 and 6.6x (Zeiss Stemi SV II, West Germany) and photographed in glycerine thinned with a few drops of 3% KOH.

Table 6: Whole Mount Staining Part A: Alcian blue stain, 1 & 2 week-old samples (adapted from Wassersug, 1976) Skin and fix in 10% neutral buffered formalin

Wash in dH2O for 24-48 hours, changing water 4-5 times Place in acetic acid/alcohol mix for 2-3 days, change solution periodically Add drops of 0.1% Alcian blue to fresh acetic acid/alcohol solution Allow to stain 12-24 hours, check frequently and remove when cartilage is blue Place in 100% EtOH for 3-4 days, changing solution periodically Proceed to Step 2 of Part B Part B: Alizarin red S stain for all samples (adapted from Clark, 1981) Step 1: For single stain samples: Skin and fix in 95% EtOH for 2-4 days Proceed to Step 2 of Part B Step 2: For all single-stained/initial Alcian blue double-stained samples: Place in 1% KOH until the bones are clearly visible through the soft tissue Add one to two drops of 0.1% Alizarin red S to fresh KOH When sufficiently stained, place samples in fresh KOH for 24-48 hours Part C: Clearing and infiltration (adapted from Kaufman, 1992) Place samples through a series of graded glycerine/KOH solutions 20% glycerine in 1% KOH 50% glycerine in 1% KOH 80% glycerine in 1% KOH The timing of each series varies for each sample: Clear the samples as much as possible without over-maceration 100% glycerine for a period of 24 hours Store in fresh 100% glycerine

Transmission Electron Microscopy Sample Determination

The three methodologies described previously complimented each other to avoid the limitations of each procedure and ensure accuracy in sample selection. All data collected from the radiographs, histology slides, and whole mount samples were compared in order to identify the age at which matrix vesicles would most likely be present in the mineralizing metapodophalangeal sesamoid bones. The samples associated with the age group(s) that were determined probable to have matrix vesicles were then selected for examination using the transmission electron microscope.

Transmission Electron Microscopy

The right front and hind paws were immediately dissected from the mice after euthanasia and prepared for transmission electron microscopy (the corresponding left paws were fixed according to the histological specifications described above). The fixation solution of 2.5% glutaraldehyde and 0.1M cacodylic buffer (pH 7.4) was combined with a chelating agent (7.5% EDTA) to facilitate sectioning the samples during ultramicrotomy (Appendix, Solution 7 and 8). The samples were fixed/decalcified with continual agitation and refrigeration (4º C) for a period of seven days. The samples were then rinsed and stored in 0.1M cacodylic buffer until all information obtained from radiography, histology, and whole mount staining was compiled and the appropriate samples were selected.

After sample selection, the second digits were dissected from the two- and three- week-old fore-paws and secondarily fixed in a solution of osmium tetroxide (OsO4) and cacodylic buffer (Appendix, Solution 9). The samples were then dehydrated in a series of solutions of EtOH and propylene oxide, infiltrated with propylene oxide and Embed 812

(Appendix, Solution 10), and embedded in Embed 812 resin for sectioning (Table 7).

Table 7: TEM Processing and Embedding Secondary Fix: Place samples in osmium tetroxide/cacodylate for 1 hour Rinse three times in cacodylate for 5 minutes each Dehydrate: 15 min in 50% EtOH 20 min in 70% EtOH 30 min in 95% EtOH 30 min in 100% EtOH, x2 30 min in 1:1 100% EtOH:Propylene Oxide (P.Oxide) 30 min in 100% P. Oxide, x2 Infiltrate: 30 min in 1:3 Embed 812:P.Oxide 1 hour in 1:1 Embed 812:P.Oxide 1 hour in 3:1 Embed 812:P.Oxide 1 hour in 100% Embed Embed: Place Embed 812 in plastic, labeled molds Place samples in epon tip in correct orientation Top molds off to obtain a mirror image in the Embed Place molds in 70º C oven overnight Allow to cool, extract resin blocks from molds for sectioning

The digits were rough trimmed and then thick sectioned at 1 μm using an industrial grade histo-quality diamond knife on an ultramicrotome (Reichert-Jung

Ultracut E, Deerfield, IL). These sections were collected on glass slides and stained using toluidine blue to determine the area of interest for thin sectioning and ultimately for

TEM examination. Thin sectioning of the resin block (at 92 nm), using a gem quality 45º

or 55º (depending on the difficulty of cutting) diamond knife at an angle of 5º, followed identification of the area of interest containing the sesamoid, distal portion of the metacarpal, proximal part of the phalanx, and the associated tendons. Chloroform was wafted over the floating thin sections in the knife trough to stretch the resin and eliminate folds. These sections were collected on charged, formvar-coated 200 mesh grids (Table

8; Appendix, Solution 11).

Table 8: Formvar Coated Grids In hood, filter Formvar solution into glass slide holder Dip cleaned slides into Formvar, x2 Allow slide to dry in an empty glass slide holder Score around the outer edge of the slide with Formvar coating Breathe heavily on the slide to allow the Formvar to lift from slide

Gently dip slide in ddH2O bath Place grids shiney side down on the floating Formvar film Press down and lift the Formvar film out of the water using a parafilm section Allow to dry, remove grids from parafilm

The grids were then positive contrast-stained using 2% uranyl acetate and lead citrate (Table 9; Appendix, Solution 12 and 13). Positive-contrast staining increases the density of the biological material as a result of the attachment of heavy metal salts to the molecules in the specimen, allowing for better viewing of the samples (Bozzola and

Russell, 1992). The prepared and stained grids were examined under a transmission electron microscope (JEOL JEM-100S, Peabody, MA) at 80 to 100 kV at magnifications of 500 to 20,000x. Kodak 35mm Panatomic-x Rapid Process Copy Film (Rochester, NY) was used to photograph areas of interest. The images were developed in a dark room in a

1:2 solution of Kodak Dektol developer (Rochester, NY) for 4 minutes, followed by a water bath rinse for 1.5 minutes. The micrographs were then fixed in Kodak Rapid Fixer

(Rochester, NY) for 5 minutes, rinsed in water for 20 minutes, and were then allowed to dry for 12 to 24 hours before being examined and scanned at 1200 dpi (Epson Expression

1600, Long Beach, CA).

Table 9: Positive-Contrast Staining Grids *Use hood, Uranyl Acetate and Lead Citrate are hazardous Filter uranyl acetate and lead citrate into two different scintillation vials Centrifuge vials for 5 minutes Line two Petri dishes with parafilm, paper side facing up

Filter double distilled water (ddH2O) and place in two plastic disposable beakers In one Petri dish, ring the outer edge with sodium hydroxide (NaOH) pellets Place drops of uranyl acetate in one Petri dish

Place drops of the filtered ddH2O beneath each uranyl acetate droplet Set grids, section side down, on the uranyl acetate droplets for 15 minutes Remove grids to the droplets of water to prevent overstaining Proceed to rinse grids gently in each beaker of water 30x Place drops of lead citrate and water in the NaOH-ringed Petri dish Place grids on the droplets of lead citrate for 2 minutes Remove grids to the droplets of water to prevent overstaining Let grids dry before examining with the TEM

All data collected on mouse sesamoids (including age of development, matrix vesicle presence, role, and association with collagen) were compiled and analyzed to provide a complete understanding of sesamoid mineralization. These data were compared to the mineralization processes reported in the literature to determine if the metapodophalangeal sesamoids develop like that of calcifying cartilage, tendon, fibrocartilage, or in a unique pattern different from either tissue.

Results

Radiography

The radiographs of the murine front and hind paws were of low resolution, especially for the one- and two-week-old mice (Fig. 1). The one-week-old samples appeared to contain little or no mineral, since no skeletal distinction could be seen in either paw (Fig. 1, a). The radiograph of the two-week-old mouse paw contained outlines of the metapodia and some amorphous condensations of bone at the joints, but a clear indication of mineralized sesamoids was not present (Fig. 1, b). In addition, the paws of each sample appeared to be in the same developmental stage as their front or hind counterpart.

Skeletal distinction in the radiographs of the three-week and older samples was more pronounced compared to the one- and two-week-old samples (Fig. 1, c). In fact, metacarpophalangeal sesamoid bones were visible in the front three-week-old paw. Two sesamoids at the metacarpophalangeal joints of digits two, three, and four can be seen

(Fig. 1, c). The poor resolution of the radiograph in the first and fifth digit of the three- week front paw obscured the presence or absence of sesamoids at these joints. The bones of the hind paw of the three-week sample were clearly defined in the radiograph, yet it was difficult to determine if the two light condensations at the second digit were the early forms of sesamoids (Fig. 1, c). The obvious presence of sesamoids at several

metacarpophalangeal joints and their near absence in the hind paw suggested that the front and hind paws of the CD-1 mouse developed at different ages.

Fig. 1 Murine front and hind paw radiographs, a) one-week, no visible bone structure, b) two-week, some distinct bone, c) clear formation of the sesamoids in the three-week front paw (arrow), d) the first evidence of a sesamoid in the four-week hind paw (arrow), e) five-week sample, f) six-week sample. Scale bar = 1 cm, magnification: 2.1x.

In the four-week-old sample, there was still too little resolution to determine if the metacarpophalangeal sesamoids of the first and fifth digit had truly developed (Fig.1, d).

However, the metatarsophalangeal sesamoids of the second, third, and fourth digit were visible as they were in the front paw. Finally, in both paws of the five- and six-week samples, all metapodophalangeal sesamoids, excluding the fifth digits of each, were clearly visible in the radiographs as dark condensations flanking either side of the distal end of the metapodia (Fig. 1, e and f).

It is believed here that the sesamoids at the fifth digit develop in a similar time frame as the other digits. However, it is not clear from the radiographs with what age group their initial presence can be identified. This is likely a result of the misalignment of the placement of the fifth digit during radiography.

Histology

The decalcification process eliminated all possibilities of testing for calcium deposits in the developing metapodophalangeal sesamoid bones of the fifth digit. The general morphology, cell appearance, and any apparent evidence of decalcified bone, such as the presence of trabeculae and marrow cavities like that in the epiphyseal plates, were examined using light microscopy. In addition, the birefringence of both the toluidine blue and Picrosirius red stained sections was important in identifying the sesamoid, fibrocartilage, and tendon relationship.

A metatarsophalangeal sesamoid bone and its associated tissues were clearly illustrated in the toluidine blue stained section depicted in Fig. 2 (only one sesamoid is

visible at a time in the medial-lateral sectioning). The sesamoids of each joint were found on either side of the distal end of the metapodia. These small bones articulated with the metapodia and proximal phalanx on the ventral surface of the joint.

Fig. 2 The metatarsophalangeal sesamoid bone of the left hind fifth digit of a one-week- old mouse. The sesamoid depicted here is less developed than the sesamoids observed in other age groups. MT metatarsal; PP proximal phalanx; MP middle phalanx; DP distal phalanx; Ses sesamoid. Toluidine blue stain. Magnification: 15.5x.

Articular cartilage was apparent along the distal and dorsal surfaces of the sesamoid at higher magnification of an older sample (Fig. 3). In addition, the flexor digiti tendons were attached to, and ran along the length of, the ventral surface of the

sesamoids. These tendons terminated along the ventral surface of the proximal phalanx at a fibrocartilaginous insertion and were especially visible under polarized light microscopy of both the toluidine blue and Picrosirius red stained sections (Fig. 4). The distinct polarization at the sesamoid insertion site in these slides was indicative of abundant parallel collagen fibers, like that of tendon.

Fig. 3 Two-week-old metatarsophalangeal sesamoid. Note the articular cartilage on the dorsal and distal surfaces that contact the metatarsal and proximal phalanx (arrowheads). Hypertrophic chondrocytes are numerous within the core of the sesamoid (asterisk). The flexor digitorum tendon (arrow) attached to the sesamoid along the ventral and distal surface, and it ended at the proximal phalanx. The area of tendon attachment between both the sesamoid and the phalanx is characteristic of fibrocartilage (F). Toluidine blue stain. Scale bar = 200 μm, magnification: 90x.

Fig. 4 The corresponding toluidine blue (left) and Picrosirius red (right) stained two- week metatarsophalangeal sesamoid. Note the abundant birefringent collagen fibers of tendon/fibrocartilage under polarized light. Scale bars = 200 μm, magnification: 45x.

Three major cell types were associated with the sesamoids in the transition from tendon, fibrocartilage, cartilage, and bone. The dorsal and distal surfaces of the sesamoids contained chondrocytes that maintained the articular cartilage in contact with the metapodia and proximal phalanges. In unmineralized sesamoids, hypertrophic chondrocytes dominated the interior region of the sesamoid resembling those seen in the epiphyseal plates. These cells, and those at the articular surfaces, were round, metachromatic, and associated with distinct lacunae typical of chondrocytes.

The two types of fibroblasts, tenocytes and fibrocartilage cells, were also abundant in the samples examined. Spindle-shaped tenocytes were documented in

longitudinal columns parallel to the collagen fibers within the tendon anterior to attachment with the sesamoid. The fibrocartilage cells transitioned from tenocyte-like cells into more ovoid cells approaching the cartilage of the sesamoid, and they were arranged in linear isogenous groups at the fibrocartilaginous insertions (Fig. 5). These cells surrounded the proximal, ventral, and distal borders of the sesamoids. Closer to the cartilaginous region of the sesamoid, the fibrocartilage cells became more round, were associated with distinct lacunae, and were chondrocyte-like. In the conversion from tenocytes into fibrocartilage cells and fibrocartilage cells into chondrocytes, it was not possible to differentiate in the transition zones.

Fig. 5 An enthesis of the sesamoid and proximal phalanx (PP) of the two-week-old mouse. Fibrocartilage cells transform into chondrocytes in linear isogenous groups (arrowhead). F fibrocartilage, MT metatarsal, PC proliferating chondrocytes, * hypertrophic chondrocytes. Toluidine blue, scale bar = 100 μm, magnification: 210x.

The one-week metapodophalangeal sesamoids were devoid of evidence of mineralization. It is possible that mineral formation had started in the front paw of these one-week-old samples, but any evidence was lost in the decalcification process. On the other hand, the sesamoids of the front paw were clearly more developed than those of the hind when considering the size of the sesamoid, the number of mature chondrocytes within it, and the development of the epiphyseal plates of the metacarpal and proximal phalanx. Proteoglycans were abundant in the developing sesamoid, metapodia, and proximal phalanges (Fig. 6). Chondrocytes in the interior of the sesamoid of the front paw were comparable to those in the hypertrophic zone of calcifying cartilage in the epiphyseal growth plate (Fig. 7). The chondrocytes in the hind paw appeared to contain proliferating cartilage cells and were more metachromatic than those observed in the front (Fig. 8).

Fig. 6 Proteoglycans (red) in the metatarsophalangeal sesamoid of the one-week mouse (asterisk). Safranin-O, scale bar = 200 μm, magnification: 85x.

Fig. 7 Metacarpophalangeal sesamoid in the one-week-old mouse. Note the presence of hypertrophic chondrocytes in the interior of the sesamoid (asterisk). MC metacarpal, PP proximal phalanx. Toluidine blue stain. Scale bar = 200 μm, magnification: 125x.

Fig. 8 Metatarsophalangeal sesamoid in the one-week-old mouse. Note the abundance of proliferating chondrocytes composing the sesamoid (asterisk). MT metatarsal, PP proximal phalanx. Toluidine blue stain. Scale bar = 200 μm, magnification: 65x.

The two-week samples, both the front and hind paws, contained hypertrophic chondrocytes in the metapodophalangeal sesamoids (Fig. 3 and Fig. 9). The degree of hypertrophy was greater in the front than the hind. In addition, these chondrocytes were more prevalent in the two-week samples than those documented in both one-week-old paws. No clear indication of sesamoid mineralization was observed in these samples.

Fig. 9 Two-week-old metacarpophalangeal sesamoid bone of the mouse. Note the abundance of hypertrophic chondrocytes in the interior of the sesamoid (asterisk), but there is no evidence of mineralization. MC metacarpal, PP proximal phalanx. Toluidine blue stain. Scale bar = 200 μm, magnification: 55x.

The first evidence of calcification was observed in the three-week-old sesamoids of the front paw (Fig. 10). Although decalcified, the appearance of irregular trabeculae and marrow cavities, signatures of spongy bone, was indicative of mineralization in these sesamoids. In the corresponding hind paw, there was still an abundant number of hypertrophic chondrocytes but no evidence of calcification (Fig. 11).

Fig. 10 Three-week metacarpophalangeal sesamoid bone (asterisk). Mineralization can be identified by the irregular pattern of trabeculae/marrow. MC metacarpal, PP proximal phalanx. Toluidine blue, scale bar = 200 μm, magnification: 75x.

Fig. 11 Metatarsophalangeal sesamoid bone of the three-week mouse. The interior of the sesamoid is primarily composed of hypertrophic chondrocytes (asterisk). MT metatarsal. Toluidine blue, scale bar = 200 μm, magnification: 75x.

No further histology was conducted on the older samples. It was apparent that the metacarpophalangeal sesamoids were osseous in the three-week-old samples. According to the literature, extracellular matrix vesicles should be present in the sesamoid as it is undergoing calcification and, by the time the structure ossifies, they could possibly be obliterated by rapidly increasing mineral deposition over the whole extracellular tissue matrix (Anderson, 1969). In this situation, the level of mineralization seen in the three- week-old metacarpophalangeal sesamoids was beyond useful examination by transmission electron microscopy. Unfortunately, the decalcification process reduced the histological data to morphological analyses alone. Thus, it was unnecessary to continue sectioning the older samples for anatomical documentation that had already been obtained in the one-, two-, and three-week samples.

Whole Mount Staining

Unlike the histology specimens, the cleared and stained samples were complete and undecalcified. This method allowed observation of the anatomical structure without any alteration to the boney tissues (such as tearing and separation of the bones at the joint often seen in the histological samples) and early formation of calcium deposits (Fig. 12).

The cleared and stained specimens provided the most compelling data regarding the mineralization of the sesamoids.

Fig. 12 Whole mount double-stained one-week-old mouse. Alcian blue = cartilage (blue), Alizarin red S = bone (red). Magnification: 2.3x.

Coinciding with radiology and histology, the one-week-old samples contained the least amount of mineralized tissue (Fig. 13). The sesamoids and epiphyseal plates of both the proximal phalanges and the metapodia were apparent as cartilaginous models at the metapodophalangeal joints in all one-week-old samples (Fig. 14). The sesamoid placement at the joint was consistent with the findings of radiography and histology.

Fig. 13 Alizarin red S single-stained, one-week-old front (left) and hind (right) paws from the same mouse. The epiphyseal plates of the metacarpals and proximal phalanges have begun to mineralize in the front paw (arrowhead). The cartilaginous models, devoid of secondary ossification centers, of the metatarsals and proximal phalanges of the hind paw are clearly visible without a double-stain (arrow). D1 first, D2 second, D3 third, D4 fourth, and D5 fifth digit. Magnification of front paw: 15x, magnification of hind paw: 13x.

Fig. 14 Medial view of an Alcian blue and Alizarin red S double-stained hind paw of a one-week-old mouse. Note the blue cartilaginous sesamoids located at the distal ends of the metatarsals (arrows). The epiphyses of the metatarsals and proximal phalanges have not yet begun to mineralize (asterisks). Note the outline of the cleared soft tissue and the remaining strands of hair on digit four and five. Magnification: 27x.

In two of the five one-week-old samples, calcium nodules were observed at the central core of the lateral-most sesamoids at the metacarpophalangeal joints of the second digits (Fig. 15). No other sesamoid mineralization was visible at any of the other joints in this age group. These results suggest that sesamoids of the second digit develop earlier than any other sesamoid in the front paw.

Fig. 15 The first evidence of mineralization at the metacarpophalangeal joint of the second digit from a one-week-old fore-paw (arrow). The secondary centers of ossification of the proximal phalanx (PP) and the metacarpal (MC) are clearly visible (asterisk and square, respectively). Alizarin red S single stain. Magnification: 47x.

In addition, as seen in the histology, the front one-week paws were more developed than the hind samples. However, the hind paws, coinciding with the early sesamoid mineral deposition, were more developed than the other one-week samples in that the distal ends of the epiphyseal plates of the metatarsals had begun to mineralize

(Fig. 16). No sesamoid mineral formation was recorded in these hind paws, suggesting that mineral formation in the epiphyseal growth plates precedes sesamoid development.

The presence of mineralization in only some one-week samples indicates that individual

variation (possibly due to sex or litter size) plays a role in murine metapodophalangeal sesamoid development.

Fig. 16 Alizarin red S single-stained, one-week-old hind paw counterpart to the front paw with a mineralizing sesamoid (Fig. 15). Note that the secondary centers of ossification of the second, third, and fourth metatarsals have begun to mineralize in this sample (arrows). The cartilaginous models of the proximal phalanges and metatarsals are clearly outlined at the metapodophalangeal joints. Faint sesamoids can be made out as soft tissue ridges flanking either side of the secondary ossification centers. Magnification: 30x.

The sesamoids of the two-week-old samples were present at all metapodophalangeal joints in both the front and hind paws. Those of the hind paws were smaller than the sesamoids in the front. All two-week sesamoids, compared to those of

the three-week samples, were smaller both in length and width. The sesamoids of the two-week hind paw were small and round, similar in shape to that of the second digit of the one-week-old samples, but opposed to large and oblate in the three-week samples

(Fig. 17). Thus, mineralization continued between two and three weeks of age.

Fig. 17 Three-week-old sesamoids (asterisks) of the front (left) and hind (right) paws of the same mouse. The sesamoids, with a characteristic oblate shape, are greater in size in the hind paw compared to those of the front. Note the degree of epiphyseal closure in the proximal phalanges and metapodia dorsal to the sesamoids in each paw. Alizarin red S single stain. Magnification: 17x.

The epiphyseal plates of both the two- and three-week-old hind paws were open according to the whole mount staining (Fig. 17). However, the three-week-old front proximal phalanges appear to be nearly fused, followed closely by the metacarpals, compared to the two-week-old plates of the phalanges and metacarpals. It was still possible in the two-week samples to distinguish the remnants of the cartilaginous models of these bones, similar to that seen in the one-week-old paws (Fig. 16).

The four-week-old epiphyseal plates of the metacarpals were nearly fused, but those of the metatarsals were still open. The sesamoids appeared to be slightly larger

than those observed in the three-week-old samples (Fig. 17), a result suggesting that final mineralization occurred at or between the ages of three and four weeks. Except for the overall larger size of the hind sesamoids in comparison to those of the front, there was no apparent difference in the morphology of the metapodophalangeal sesamoids of four-, five-, and six-week-old samples. In addition, beyond three weeks of age, the sesamoids of both the front and hind paws were large and oblate with no apparent signs of calcified fibrocartilage at the insertion sites of the tendons.

These data suggest that, except for early cases of mineralization in the one-week lateral sesamoid of the second digit of the forepaw, most sesamoid mineralization occurs between one and two weeks of age. Growth continues through three or four weeks of age until the full size of the sesamoid is reached. The sesamoids can be considered to be mature at four weeks of age. Calcifying fibrocartilage at the entheses may develop, but this apparently must occur sometime after six weeks of age.

Transmission Electron Microscopy Sample Determination

Samples were chosen for TEM examination based on the data compiled from the radiography, histology, and cleared and stained specimens. According to the results from histology and radiography, most mineralization took place between two and at least three weeks of age. However, the cleared and stained samples indicated that initial mineral formation occurred first in the lateral sesamoid at the metacarpophalangeal joint of the second digit of some one-week-old samples. Sesamoids continued to mineralize through

at least the third week of age. Finally, all procedures revealed that the front paws developed in advance of the hind.

Since the two-week-old samples in all methodologies appeared to be in intermediate stages of mineralization, these samples were selected for TEM analysis. In addition, the second digit from the front paw was considered the most likely candidate for obtaining information regarding the presence and role of matrix vesicles in the mineralization process. Thus, the two-week-old second metacarpophalangeal joint was processed, embedded, sectioned, stained, and analyzed for electron microscopy by methods described previously. The corresponding three-week-old samples were also chosen as TEM samples for comparison.

Transmission Electron Microscopy

The toluidine blue stained thick section of the two-week-old sample illustrated in

Fig. 18 will serve as a guide to the thin sections described below. The sesamoid was located ventral to the metacarpal and proximal to the phalanx. Areas of interest included the apparent tendon connecting the sesamoid to the proximal phalanx and the dense connective tissue at the opposite end of the bone on the dorsal surface. Both of these areas were considered to be composed of fibrocartilage. The borders and interior of the sesamoid, the ligament, and the dense connective tissue were examined for evidence of mineralization, matrix vesicles, and collagen orientation.

Fig. 18 Toluidine blue-stained thick section of the two-week, lateral metacarpophalangeal sesamoid from the second digit. MC metacarpal, PP proximal phalanx, area of dense connective tissue (arrowhead), mineralizing core of the sesamoid (asterisk), tendon (arrow). Scale bar = 200 μm, magnification: 105x.

Extracellular matrix vesicles were found in the murine two-week-old, lateral-most metacarpophalangeal sesamoid bone examined in the second digit (Fig. 19). However, their association with the surrounding cells, their number, location within the sesamoid, and relation to the collagen were not expected or what had been documented in the literature elsewhere. The results described below indicate the possibility that the observed matrix vesicles were not involved in the sesamoid mineralization process.

Fig. 19 Numerous matrix vesicles in the spaces between cells composing the dense connective tissue of the proximal border of the two-week metacarpophalangeal sesamoid (arrowheads). Extensive endoplasmic reticulum was characteristic for these fibrocartilage cells. N nucleus, ER endoplasmic reticulum. Lead citrate/uranyl acetate stain. Scale bar = 1 μm, magnification: 16,000x.

The majority of the vesicles were associated with long cell processes originating from both chondrocytes and fibrocartilage cells (Fig. 19). However, several matrix vesicle-like structures were also found near the membranes of these cells where others were observed at far distances from the cells altogether. One to as many as seven matrix vesicles were documented at a time near cell extensions or membranes.

In addition, most matrix vesicles were found associated with cells that were near the proximal, distal, and ventral borders of the sesamoid as opposed to near the calcifying front. However, small numbers of matrix vesicles were discovered embedded deep

within the proximal border of the sesamoid near the calcifying center associated with chondrocytes (Fig. 20). These vesicles were linearly arranged in two groups of three, isolated from each other. A short distance away from the vesicles, the interior of the sesamoid appeared to be calcifying cartilage and contained hypertrophic chondrocytes

(Fig. 21). Matrix vesicles were not observed, or any evidence of their presence, in this region of the sesamoid.

Fig. 20 Matrix vesicles near the surface of a chondrocyte in proximity to the calcifying core of the two-week metacarpophalangeal sesamoid (arrows). Lead citrate/uranyl acetate stain. Scale bar = 1 μm, magnification: 33,000x.

Fig. 21 Chondrocytes within the calcifying cartilage of the two-week metacarpophalangeal sesamoid. Distinct lacunae are visible surrounding the cells, but some shrinkage artifact is present. The extracellular matrix is composed of collagen, proteoglycans, and other molecules. No matrix vesicles were documented in the calcifying cartilage areas. Lead citrate/uranyl acetate stain. Scale bar = 6 μm, magnification: 13,000x.

The collagen orientation in the areas where matrix vesicles were found was predominately perpendicular to the plane of section (Fig. 22). Very few regions of parallel collagen fibers, except where sectioned through the tendon away from the border of the sesamoid and at intervals in areas believed to be composed of fibrocartilage, were documented (Fig. 23). No matrix vesicles were found in these parallel collagen fiber regions. In the calcifying cartilage in the interior of the sesamoid, an abundant network of proteoglycans and occasional small collagen fibers was documented.

Fig. 22 Cross-sectioned collagen fibers in the two-week metacarpophalangeal sesamoid composed the majority of the extracellular space (left of the cell). Perpendicular fiber orientation was documented in the ligament along the distal edge of the sesamoid. Lead citrate/uranyl acetate, scale bar = 1 μm, magnification: 18,000x.

Fig. 23 The proximal border of the two-week metacarpophalangeal sesamoid contained parallel collagen fibers (asterisks), but fibers were perpendicular to the plane of section in the majority of the sample. Note the long cell processes, characteristic of cell-cell communication. Lead citrate/uranyl acetate, scale bar = 1 μm, magnification: 19,000x.

Three cell phenotypes were distinguished in the two-week samples, similar to those described in the literature review section and histology results of this thesis. The tendon and early fibrocartilage regions, just outside the proximal and distal borders of the sesamoid, contained fibroblast cell types. The cells located in fibrocartilaginous regions had a well developed endoplasmic reticulum and numerous, random extensions. Cell-cell communication was frequently documented in these areas (Fig. 23). As fibrocartilage cells moved closer toward the mineralizing core of the sesamoid, they appeared to resemble chondrocytes in various stages of development. These cells were larger and rounder closer to the center of the sesamoid and they resided in well defined lacunae (Fig.

20). The calcifying cartilage in the center of the sesamoid was associated with hypertrophic chondrocytes that were receding from the edges of their territorial matrix.

Several cell extensions, although much shorter, were also observed (Fig. 21).

The three-week-old sesamoid, in comparison to the two-week samples, was significantly more mineralized. This was obvious not only in the TEM examination, but also in the toluidine blue-stained thick section that was obtained before thin sectioning ensued (Fig. 24). Matrix vesicles were not found in the proximal, distal, or medial regions of the sesamoid, a result also documented in the two-week-old samples.

Fig. 24 Toluidine blue-stained thick section of the three-week-old metacarpophalangeal sesamoid from the second digit. MC metacarpal, PP proximal phalanx, area of dense connective tissue (arrowhead), mineralizing core of the sesamoid (asterisk), fibrocartilaginous area (arrows). Scale bar = 200 μm, magnification: 95x.

The limited number of matrix vesicles found in the developing sesamoid of the two-week-old mouse were not clearly associated with mineral deposition or the mineralization process as a whole. Located far from the calcifying front and in a few isolated groups, the matrix vesicles likely contributed to other cell functions not involved with initial mineral formation. Evidence of collagen-mediated mineralization events were not specifically detected in the tendon and fibrocartilage regions although mineral formation had occurred in the sesamoids by two weeks of development. Such mineral deposition was clearly associated with the sesamoid extracellular matrix comprised principally of collagen and proteoglycans.

Discussion

The three methods utilized here to investigate the general morphology, anatomy, and age at initial calcification of murine metapodophalangeal sesamoid mineralization were more powerful and useful than using a single method alone. The resolution of radiography was insufficient for identifying early mineral formation of bones under the age of three weeks. However, it did provide information regarding sesamoid mineral development in older mouse paw samples, identification and location of mineral at the metapodophalangeal joints, and apparent differences in bone growth between front and hind paws of the animals.

Complimenting radiography, histology provided a more in-depth view of the sesamoid developmental process. Safranin-O, toluidine blue, and Picrosirius red staining of decalcified sections allowed data to be collected regarding the proteoglycan content, general morphology, cell arrangement, and interaction between the sesamoid, tendon, and other bones of the joint. This information strongly correlated with the abundant literature regarding tendon, fibrocartilage, and bone. Light microscopy and polarized light images also illustrated intimate structural relationships between the sesamoid bone and tendon at the metapodophalangeal joint. The most important information to this thesis, the early mineral content of the tissues, was regrettably lacking in the decalcified samples used for

histology. Therefore, it was determined unnecessary to examine histologically samples of mice older than three weeks, the age at which the metacarpophalangeal sesamoids were clearly ossified.

Providing macroscopic information like radiography, whole mount staining was able to detect mineral development in early ages of mice and in specific locations within the bones of their paws. This method further contributed information that the low- resolution radiography was unable to yield. Cleared samples unequivocally showed the first signs of mineral formation when staining was carried out with Alizarin red S as a marker for calcium. The earliest mineralizing sesamoid was identified in the second digit of the front paws of the one-week-old mouse, a result that was not apparent from radiography and histology. Compared to the other samples of the same age, the hind paw counterparts of these samples were developmentally advanced in their epiphyseal plate development, which apparently precedes sesamoid mineral formation. Sesamoid mineralization continued through at least three weeks of age. No indication of mineral was observed in the fibrocartilaginous insertion of the tendon to the sesamoid bone in samples of all age groups. In addition, cleared and stained samples allowed observation of the development of the epiphyseal plates and the position of the sesamoid bones at the joint in a three-dimensional representation.

The early formation of a mineralizing sesamoid in some of the one-week-old front paws indicates that developmental differences occur in the calcification of these bones.

Sex, litter size, environmental, and/or individual animal differences could be responsible for early sesamoid development, a result supporting the findings of Spark and Dawson

(1928). In addition, the presence of a sesamoid at one week of age confirmed the findings of Wirtschafter (1960). The disparity associated with the absence of these sesamoids at two weeks of age reported by Kaufman (1992) could be a result of developmental differences between strains of mice.

In addition to individual differences, variations in mineralization were documented in the paws and digits of the mice studied here. The differential onset of calcification of the front and hind paws has been widely documented in both the mouse and rat (Johnson, 1933; Spark and Dawson, 1928; Strong, 1925). The hind paw is reported to begin mineralization in advance of the front paw shortly before and after birth. However, seven days after birth, the paw gives the appearance of delayed development because of its greater size compared to that of the front (Spark and Dawson,

1928). This developmental pattern was observed in the one- to six-week-old murine front and hind paws examined in this study.

Furthermore, the second digits are reported to mineralize before the others in the mouse and rat (Patton and Kaufman, 1995; Spark and Dawson, 1928; Strong, 1933). The literature indicates that the second digit is closely followed by the development of the third and fourth digit, then the fifth, and lastly the first in a matter of days in these animals (Patton and Kaufman, 1995; Spark and Dawson, 1928; Strong, 1933). These observations support the results found here regarding advanced development of the lateral metacarpophalangeal sesamoid of the second digit and its associated joint. A similar calcification pattern is documented in a variety of animals, including humans

(Davies and Parsons, 1927). The similar progression of the mineralization of the autopod

in different species signifies the genetically controlled and regulated development of the paws, digits, and sesamoids, much like the other elements of the skeleton (Atchley and

Hall, 1991; Karaplis, 2002).

The compilation of data obtained from radiography, histology and whole mount staining provided an informed selection of the best sample age group of mice to investigate for the presence of matrix vesicles in sesamoid mineralization. As a result, two-week-old samples were found to be an optimal age group for investigating the potential roles of matrix vesicles and collagen in sesamoid calcification using TEM.

Examination of these samples revealed a few small clusters of matrix vesicles present in the fibrocartilaginous areas of the sesamoid, except in one case where they were associated with chondrocytes near the periphery of the calcifying core.

Matrix vesicle association with the mineralization process of tendon, fibrocartilage, or cartilage in the metapodophalangeal sesamoids was inconclusive. Very few matrix vesicles were found in these tissues, an observation that was surprising in comparison to their large numbers in tendon, calcifying cartilage, and early bone reported in the literature. In these tissues, matrix vesicles were documented to be abundant within the extracellular spaces between their respective cells (tenocytes, chondrocytes, fibrocartilage cells, or osteoblasts) (Anderson, 1969; Landis, 1986; Yamada, 1970). In addition, the vesicles found here were in areas distant from the mineralizing interior of the sesamoid, another result raising doubt about their involvement in the mineralization process in the sesamoid samples.

Cell arrangement and phenotype associated with the sesamoid were comparable to that found in the literature. Fibroblasts were typical within both the sesamoid-associated tendon and fibrocartilage. The flat, spindle-shaped tenocytes, arranged in extensive columns parallel to the orientation of collagen fibers, possessed extensions protruding from the proximal and distal surfaces of their enclosing outer cell membranes.

Fibrocartilage cells transitioned from tenocytes into chondrocytes in columnar groups, characteristic of that documented for the four zones within entheses. Chondrocytes composed the articular surfaces and interior of the sesamoids. Hypertrophic chondrocytes within the calcifying core were similar to those within epiphyseal growth plates of other tissues. Thus, the cellular arrangements and phenotypes found within sesamoids correlated well with the findings from the literature.

An examination of collagen orientation, like the relative absence of matrix vesicles, gave unexpected results. At insertion points subject to force, collagen fibers are aligned in the direction of the loading force (Bancroft and Gamble, 2002). In this thesis, collagen was anticipated to be found in a parallel arrangement continuous with that of the associated tendon and portions of the insertion site with the sesamoid. An interweaving network of collagen was expected in the transition from fibrocartilage to cartilage.

However, although columnar cellular arrangement within the tissues examined was consistent with that reported in the literature, collagen in the entheses of the metapodophalangeal sesamoids was predominately viewed in cross section in most areas.

It is possible that, although the cells appeared to be arranged in columns, the plane of section of embedded specimen blocks was not exactly parallel to collagen alignment.

This misalignment could have complicated a clear examination of matrix vesicles and collagen in TEM analyses.

The necessary decalcification of the bone samples prevented identification of specific mineral nucleation and deposition events associated with either matrix vesicles or collagen fibers. No signature evidence of the remnants of mineralization, such as ghost crystals described in Lowenstam and Weiner (1989), were observed in either structure. Therefore, it could not be determined conclusively whether the tendon and/or fibrocartilage insertion site contributed to mineralization of the metapodophalangeal sesamoid bones of mice.

From the results of this study, the sesamoids appear to mineralize from the central core of the cartilaginous model, which expands gradually as the calcification progresses within the tissue. Little evidence was observed suggesting that matrix vesicles were involved in this process. However, the abundant proteoglycan networks, both in the one- week-old samples stained with Safranin-O and the two-week mineralizing sesamoid core in the TEM analysis, appear to be important in the mineralization process. In addition, type II collagen, the most abundant extracellular product of chondrocytes and expected to compose the majority of the calcifying cartilage of the sesamoid, should be a principal mineralization mediator in the absence of matrix vesicles.

Conclusion

Metapodophalangeal sesamoid bones of CD-1 mice began mineralization as early as one week of age. The calcification of the epiphyseal plates of the metapodia and proximal phalanges began in advance of sesamoid mineral formation. Mineralization occurred first in the one-week metacarpophalangeal joint of the second digit as a mineral nodule within the cartilaginous core of the sesamoid. Complete calcification was obtained between three and four weeks, resulting in a large, oblate sesamoid.

Ossification of these small bones occurred before the epiphyseal plates were fully fused, as observed in the five- and six-week-old samples. No evidence of calcified fibrocartilage was documented in the entheses of the sesamoids.

In the TEM analysis of the developing sesamoid, extracellular matrix vesicles that were found were not believed to be involved or associated with mineralization nucleation events of this bone. The lack of abundant matrix vesicles near the calcifying core of the sesamoid and the surrounding tissues was inconsistent with the existing literature on vesicle roles in calcifying cartilage, tendon, and fibrocartilage. In addition, the perpendicular orientation of the collagen fibers in the tissues examined differed from the results of other studies. Therefore, the role of matrix vesicles in the mineralizing sesamoid bones was determined to be inconclusive. It is assumed that extensive

proteoglycan networks and type II collagen fibers initiate mineralization in murine metapodophalangeal sesamoid bones.

Future Research

To understand the process of mineralization fully in murine metapodophalangeal sesamoid bones, future research should focus on the shorter age intervals between one and three weeks of age. It was determined that most of the initial mineral formation occurred in the sesamoids between one and two weeks of age and complete mineralization was established by three or four weeks of age. Investigation of sesamoid development at shorter intervals could identify the specific age range of their mineralization and contribute to our knowledge of individual and sex differences in mice.

Decalcification of samples for ease of sectioning prevented identification of mineral formation in the histology and TEM samples and precluded a direct determination of the role of matrix vesicles in mineralization of the sesamoids.

Examination of undecalcified samples, in both histological and TEM prepared samples, would provide valuable information regarding initial mineral formation. Histological samples stained with Alizarin red S, sectioned without prior decalcification, would determine specific areas of mineral at specific animal ages. TEM analyses of undecalcified specimens would also provide critical information regarding the exact location of the mineralizing front and nucleation events associated with matrix vesicles or collagen fibers in the tendon, fibrocartilage, and cartilage of the sesamoid.

Further studies are necessary to determine the role of collagen and proteoglycans in mineralization of sesamoid bones. Matrix vesicle presence should first be re-examined in samples known to be sectioned parallel to the collagen orientation in the tendons and their insertions into sesamoids. It is possible that matrix vesicles were obscured from view in the apparently perpendicular orientation of the collagen fibers examined in this study. In addition, the role of type II collagen and proteoglycans in the mineralization process is largely unknown. Lowenstam and Weiner indicate that each of these molecules may be key players in mineralizing structures (1989). The results presented here suggest further research should be conducted on type II collagen and proteoglycans to determine their possible involvement in calcification of the metapodophalangeal sesamoid bones of mice.

Investigating the development and mineralization of sesamoid bones has many important implications in a variety of fields of science, including biological anthropology, developmental biology, biomineralization studies, and biomedical science.

Few studies have been conducted on sesamoids other than those concerning the patella.

Other sesamoids may mineralize differently, especially when considering species variability. Further studies of sesamoid mineralization could pinpoint the potential role of matrix vesicles in initial mineral formation and their relationship with collagen fibers within these small bones.

Comprehending the mineralization process fully could facilitate the identification and treatment of human pathological conditions that affect the skeleton, such as rheumatoid arthritis. Furthermore, biological anthropology, concerned with human

evolution, obtains data from human (or human ancestor) skeletal remains. The mineralization of bones and the processes that affect them (diet, aging, and disease) are vastly important for reconstructing the lifestyle of that individual. Understanding how physiological and environmental factors affect the initial mineralization process of bones is important to the field of biological anthropology.

It is in this context that a more complete understanding of the mineralization process of murine metapodophalangeal sesamoid bones will contribute to many fields of science. The ultrastructural anatomy of developing sesamoids and their associated tendons and entheses are ideal for further calcification studies. These small bones remain inadequately researched in their variety both within and between species. It is very possible that sesamoid bones will end the debate between matrix vesicle function and presence in mineralizing tissues.

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Appendix

Solution 1: Decalcification Solution (Bancroft & Gamble, 2002) 5.5 g EDTA disodium salt 10 mL Formaldehyde (37-40% stock) 90 mL dH2O

Solution 2: 3% Ammonium Oxalate Solution (Bancroft & Gamble, 2002; Brown, 1978) 3 g Ammonium oxalate 100 mL dH2O

Solution 3: Toluidine Blue Solution (Clark, 1973) 0.5 g Toluidine blue 500 mL dH2O

Solution 4: Picrosirius Red Solutions (Puchtler et al, 1973; Junqueira et al, 1979) Picrosirius: 0.5 g Sirius red F3B (C.I. 35782) 500 mL Saturated aqueous solution of picric acid *Add a little solid picric acid to ensure saturation Acidified Water: 5 mL Glacial acetic acid 995 mL dH2O

Solution 5: Safranin O Weigert's Iron Hematoxylin: Solution A: 1 g Hematoxylin 100 mL 95% EtOH Solution B: 4 mL Ferric iron chloride stock

29 g Anhydrous FeCl3

100 mL dH2O

95 mL dH2O 1 mL Concentrated hydrochloric acid *Note: Store each solution seperately until use as a 1:1 solution 0.1% Safranin O: 1% Light Green Stock: 0.1 g Safranin O 1 g Light green

100 mL dH2O 100 mL dH2O 1 mL Glacial acetic acid Light Green Working Solution: 10 mL Light green stock 40 mL dH2O

Solution 6: Whole Mount Staining Solutions (Wassersug, 1976) 1% KOH: Acetic Acid/Alcohol: 4 g KOH 60% EtOH (100%)

400 mL dH2O 40% Glacial acetic acid 0.1% Alizarin Red S: 1% Alcian Blue: .01 g Alizarin red S 0.1 g Alcian blue 10 mL KOH 6 mL 100% EtOH 4 mL Glacial acetic acid

Solution 7: 0.1 M Cacodylic Buffer Solution *Cacodylic acid is a highly toxic and carcinogenic substance 2.14 g Cacodylic acid

100 mL ddH2O Mix solution in hood and pH to 7.4

Solution 8: TEM Decalcification Solution (7.5% EDTA in 2.5% Glutaraldehyde) 10 mL of 8% Glutaraldehyde 22 mL of 0.1M Cacodylic buffer 2.4 g EDTA Mix solution in hood for 30+ minutes until fully dissolved

Solution 9: Osmium Tetroxide *Osmium tetroxide is a neurotoxin

.5 g Osmium (OsO4) 50 mL 0.1M Cacodylic buffer (pH 7.4) In hood, put vial of osmium on dry ice to reduce vapors Gently tap osmium crystals off wall of vial Score vial and break Pour crystals into brown, aluminum foil wrapped bottle of 0.1M cacodylate Seal bottle with parafilm Let sit overnight to go into solution

Solution 10: EMbed 812 Resin Solution A: 20 mL Embed 812 30 mL Dodecenyl succinic anhydride (DDSA) Solution B: 27 mL Embed 812 23 mL Nadic methyl anhydride (NMA) Mix solutions in separate plastic disposable beakers for 15 minutes each using a teflon-coated helical rotor in a hood. Mix on low to avoid creating bubbles. Add Sol. B to Sol. A during continuous mixing Add 2% 2,4,6-tri-dimethylaminomethyl phenol (DMP-30) as a catalyst Mix for 15 minutes on low speed

Solution 11: 6% Formvar Solution 6 g Formvar 100 mL Chloroform Mix in glass container in hood

Solution 12: 2% Uranyl Acetate Solution *Use extreme caution, Uranyl Acetate is very toxic 2 g Uranyl acetate

100 mL dH2O In hood, shake solution vigorously in an aluminum foil-wrapped, brown bottle Leave the solution overnight to go into solution

Solution 13: 2.6% Reynold's Lead Citrate Solution 1 N Sodium Hydroxide (NaOH): 1 g NaOH

25 mL ddH20 Shake until in solution Lead Citrate: 1.76 g Sodium citrate

50 mL ddH20 1.33 g Lead nitrate In hood, place 1.76 g sodium citrate into flask

Add 30 mL dH2O, gently invert flask until into solution Add 1.33 g lead nitrate, gently invert flask at intervals for 30 minutes Add 5 mL 1 N NaOH, invert to clear pH solution to 12.0 +/- 0.1 using drops of 1 N NaOH

Add dH2O to make 50 mL Pour into 3 scintillation vials, seal with parafilm, and store at 20º C