ARTICLE IN PRESS
Journal of Biomechanics 36 (2003) 1409–1424
Survey
‘‘Whither flows the fluid in bone?’’ An osteocyte’s perspective
Melissa L. Knothe Tatea,b,c,d,*
a
Department of Biomedical Engineering, ND 20, The Lerner Research Institute, Cleveland Clinic Foundation, 9500 Euclid Avenue, Cleveland,
OH 44195, USA
b
Department of Orthopaedic Surgery, Orthopaedic Research Center, The Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA
c
Department of Biomedical Engineering, Case Western Reserve University, Cleveland, OH, USA
d
Department of Mechanical Engineering & Aeronautics, Case Western Reserve University, Cleveland, OH, USA
Accepted 27 March 2003
Abstract
Bone represents a porous tissue containing a fluid phase, a solid matrix, and cells. Movement of the fluid phase within the pores or
spaces of the solid matrix translates endogenous and exogenous mechanobiological, biochemical and electromechanical signals from
the system that is exposed to the dynamic external environment to the cells that have the machinery to remodel the tissue from
within. Hence, bone fluid serves as a coupling medium, providing an elegant feedback mechanism for functional adaptation. Until
recently relatively little has been known about bone fluid per se or the influences governing the characteristics of its flow. This work
is designed to review the current state of this emerging field. The structure of bone, as an environment for fluid flow, is discussed in
terms of the properties of the spaces and channel walls through which the fluid flows and the influences on flow under physiological
conditions. In particular, the development of the bone cell syncytium and lacunocanalicular system are presented, and pathways for
fluid flow are described from the systemic to the organ, tissue, cellular and subcellular levels. Finally, exogenous and endogenous
mechanisms for pressure-induced fluid movement through bone, including mechanical loading, vascular derived pressure gradients,
and osmotic pressure gradients are discussed. The objective of this review is to survey the current understanding of the means by
which fluid flow in bone is regulated, from the level of the skeletal system down to the level of osteocyte, and to provide impetus for
future research in this area of signal transduction and coupling. An understanding of this important aspect of bone physiology has
profound implications for restoration of function through innovative treatment modalities on Earth and in space, as well as for
engineering of biomimetic replacement tissue.
r 2003 Elsevier Ltd. All rights reserved.
Keywords: Bone; Interstitial fluid; Fluid flow; Osteocyte; Lacuna; Canaliculus; LCS; Shear stress; Bone cells; Mechanotransduction; Tissue
engineering
1. Introduction
Bone is a living ecosystem comprised of 25% fluid.
The remaining 75% of bone’s structure contains organic
and mineral components (Piekarski, 1970). These liquid
and solid components are optimally configured as a
composite structure, organized hierarchically from a
system (the skeleton) to an organ, tissue, cellular,
subcellular and molecular level, to allow for dynamic
*Corresponding author. Department of Biomedical Engineering,
ND 20, The Cleveland Clinic Foundation, Lerner Research Institute,
9500 Euclid Ave., Cleveland, OH 44195, USA. Tel.: +1-216-445-3223;
fax: +1-216-444-9198.
E-mail address: [email protected] (M.L. Knothe Tate).
0021-9290/03/$ - see front matter r 2003 Elsevier Ltd. All rights reserved.
doi:10.1016/S0021-9290(03)00123-4
optimization of structure for function (Fig. 1, Table 1).
As a system of bones, the skeleton fulfills scaffolding,
armoring, and damping functions necessary for survival
of mobile, terrestrial organisms (Pauwels, 1973; Piekarski, 1973). At an organ and tissue level, bone provides
more surface area for exchange and filtering of solutes,
to and from the vascular and lymphatic systems, than
any other organ in the body (Robinson, 1964; Tami
et al., 2003) (Fig. 1A,B). Furthermore, bone, as an organ
and tissue, provides metabolic and hematopoetic (blood
building) physiological functions (Fig. 1A,B). Cells,
ensconced throughout the mineralized matrix tissue of
bone (osteocytes, Fig. 1, C1; Fig. 2), and residing along
vascular spaces and bone resting surfaces (osteoblasts
and osteoclasts, Fig. 2) represent the living component
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M.L. Knothe Tate / Journal of Biomechanics 36 (2003) 1409–1424
Fig. 1. The hierarchical organization of bone is necessary to allow for dynamic optimization of structure for function. The interaction between the
solid and fluid phases of bone underlies much of the mechanical (e.g. viscoelastic damping, elasticity) and metabolic (e.g. molecular sieving, storage
and mobilization of calcium, hematopoiesis) behavior of bone but is poorly understood. Relevant dimensions for fluid spaces in bone are summarized
in Table 1. (A) Principal stress trajectories in the femoral head (left) (after Gray, 1973). Corresponding radiograph of a human femoral head with
trabecular organization corresponding to principal stress trajectories (right). (B) Schematic diagram of a wedge of cortical bone between the
endosteum and periosteum (after Junqueira et al., 1995). (C1) Three-dimensional reconstruction of confocal image stacks from an osteocyte and its
processes. (C2) Scanning electron micrograph of the interior surface of a lacuna. Holes are canaliculi entering the lacuna perpindicular to the plane of
the image. Branching canaliculi can be seen on the fractured surface, in the plane of the image. Spaces between the mineralized matrix, as seen in this
image, and the surface of the osteocyte and its processes, as seen in the previous image, are filled with interstitial fluid (Image courtesy of R.G.
Richards, Ph.D., AO Research Institute, Davos). (D) In relief, the canaliculi appear striated due to collagen fibrils lining the inner wall surface (after
Reilly et al., 2001). Molecules such as hyaluronan (E), control the osmotic pressure, hydration and permeability properties of the tissue (after
Sheehan and Almond, 2001).
of the tissue (Junqueira et al., 1995). The cells provide
the machinery for remodeling, allowing for adaptation
of bone structure to fulfill dynamic functional demands
(Wolff, 1892; Pauwels, 1973; Frost, 1963). Osteocytes,
osteoblasts, and to some degree osteoclasts, form a
functional ‘‘syncytium’’, or interconnected network
(Fig. 2), in their own right (Rasmussen and Bordier,
1974; Aarden et al., 1994, Curtis et al., 1985), allowing
for communication and transport between cells deep
within the tissue (i.e. osteocytes) and those located
within the vicinity of vascular spaces and bone surfaces
(i.e. osteoblasts and osteoclasts) to which loads are
imparted via the muscles, tendons and ligaments (Fig. 2)
(Knapp et al., 2001). This interconnectedness is intrinsic
to both the cell structure, e.g. the cytoskeleton, as well as
the common fluid space shared by the cells, i.e. fluid
within the lacunocanalicular system and intermatrix
porosity (Fig. 1, C2). Finally, molecules (Fig. 1, D), e.g.
hyaluronan, control the osmotic pressure, hydration and
permeability properties of the tissue (Knothe Tate and
Niederer, 1998; Midura et al., 1994; Sheehan and
Almond, 2001).
The ecosystem bone is subjected to a dynamic
environment in which functional adaptation is necessary
for survival. Bone tissue health depends on the ability of
bone cells to recognize and respond to mechanical and
chemical stimuli, a process referred to as mechanochemical transduction (Wang et al., 1993; Berthiaume,
2000; Steck et al., 2001, 2003). Remodeling activity,
coordinated between osteocytes, osteoclasts, and osteoblasts (Fig. 2), provides a basis for adaptation.
Osteocytes, the most abundant cells in bone, are
Table 1
Approximate limiting dimensions for vascular and extravascular spaces within bone (see Fig. 1): based on experimental study with molecular tracers of known dimension (after Knothe Tate, 2001);
relative to tracer size
Bone space/structure
(A)
Bone
Haversian system
Trabeculae
Lamellae
Secondary
Epithelial fenestrae of
marrow cavity vessels
Capillaries
5–7 10 5 m (Atkinson and Hallsworth,
1982; Cooper et al., 1966)
5–12 10 5 m (Atkinson and
Hallsworth, 1982)
2.5–5 10 5 m (Atkinson and
Hallsworth, 1982)
1 10 5 m (Dillaman, 1984; Landis and
Pappenheimer, 1963)
1 10 7 m (Hughes et al., 1978)
3–4.5 10
9
m (Hughes et al., 1978)
(B)
Lacunocanalicular system
Lacunae
99m
Tc-Red blood cells
(McDougall, 1970)
Microspheres
Glycogens
Saccharated iron oxide
particles
Ferritin (native)
1–1.3 10 5 m (Atkinson and
Hallsworth, 1982)
Space between osteocyte
surface and mineralized
wall of lacuna
Canaliculi
3–4 10 5 m (Chakkalakal, 1989)
5–7 10 6 m (Ascenzi et al., 1965)
2.7 10 6 m (Wasserman and Yaeger,
1965)
6
m (Gross et al., 1981)
12–15 10 9 m (Simionescu and
Simionescu, 1987)
7–10 10 9 m (Simionescu and
Simionescu, 1987)
1–10 10 9 m (McDougall, 1970)
5.5 10 9 m (Simionescu and
Simionescu, 1987)
11 10 9 m (De Bruyn et al., 1985)
15,000–300,000
445,000–46,000 (Adamson,
unpublished data)
440,000 (Simionescu and
Simionescu, 1987)
340,000 (Handagama et al., 1989)
9
5.22 10
data)
m (Adamson, unpublished
IgG
9
LDH
4.6 10
data)
Myeloperoxidase
4.4 10 9 m (Simionescu and
Simionescu, 1987)
240,000 (Simionescu and
Simionescu, 1987)
180,000 (Handagama et al., 1989)
m (Adamson, unpublished
160,000
Aldolase
158,000
Transferrin
76,000–88,000 (Morris et al., 1982)
1411
1–10 10 7 m (Atkinson and
Hallsworth, 1982; Cooper et al., 1966;
Boyde, 1972)
5–6 10 7 m (Knapp et al., 2001; Reilly
et al., 2001)
15 10
Fibrinogen
Catalase
Intralacunar distance
M.W. (g/mol unless otherwise
indicated)
2.5 10 4 m (Chakkalakal, 1989)
1.5–2.0 10 4 m (Ham, 1965)
3–7 10 6 m (Ham, 1965)
Dextrans
Capillary wall pores
Diameter
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Volkmann canal
Tracer
M.L. Knothe Tate / Journal of Biomechanics 36 (2003) 1409–1424
Vascular system
Haversian canal
Primary
Limiting dimension (diameter, unless
otherwise indicated)
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Table 1 (continued )
Bone space/structure
Limiting dimension (diameter, unless
otherwise indicated)
Osteocyte process
8.5–10 10
8
m (Cooper et al., 1966)
Tracer
Diameter
Serum albumin
(4 14) 10
Lactoperoxidase
Hemoglobin
1.42–5.0 10 8 m (Cooper et al., 1966)
1–20 10 7 m (Wasserman and Yaeger,
1965)
3.25 10
Postulated gap junctions
between cell processes
(C)
Bone matrix pores, i.e.,
collagen–hydroxyapatite
porosity
8
5–12 10
9
m (Holmes et al., 1964)
(Small fraction 12–25 10
et al., 1964))
9
3.55 10 9 m (Adamson, unpublished
data)
3.6 10 9 m (Simionescu and
Simionescu, 1987)
2.8 10 9 m (Simionescu and
Simionescu, 1987),
3.1 10 9 m (Adamson, unpublished
data)
3.25 10 9 m (Pappenheimer, 1955)
64,000 (Morris et al., 1982)–69,000
(Pappenheimer, 1955)
82,000 (Simionescu and Simionescu,
1987)
68,000 (Pappenheimer, 1955)
Thorotrast (colloidal thorium
dioxide) (Guzelsu and Walsh,
1990)
Horseradish peroxidase
5–6 10
3.0 10
data)
m (Cooper et al., 1966)
2 10 8 m (Atkinson and Hallsworth,
1982)
B2 10 9 m (Doty, 1981)
m (De Bruyn et al., 1985)
Colloidal gold spheres
3–7 10
Orosomucoid
3.0 10
data)
Ovalbumin
2.76 10
data)
2.74 10
data)
2.25 10
data)
2.08 10
data)
2.06 10
data)
2.02 10
data)
1.87 10
data)
9
9
40,000 (McDougall, 1970)
m (Adamson, unpublished
9
9
m (McDougall, 1970)
m
m (Adamson, unpublished
38,000 (Adamson, unpublished
data)
9
43,000
data)
35,000
data)
25,000
data)
13,683
data)
14,100
data)
14,176
data)
12,284
data)
m (Holmes
b-lactoglobulin
Chymotrysinogen A
Ribonuclease
Lysozyme
Alpha-lactalbumin
Cytochrome c
m (Adamson, unpublished
19
m (Adamson, unpublished
9
m (Adamson, unpublished
9
m (Adamson, unpublished
9
m (Adamson, unpublished
9
m (Adamson, unpublished
9
m (Adamson, unpublished
(Adamson, unpublished
(Adamson, unpublished
(Adamson, unpublished
(Adamson, unpublished
(Adamson, unpublished
(Adamson, unpublished
(Adamson, unpublished
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Space between
mineralized and
unmineralized zone of
lacunae
1 10 7 m (Wasserman and Yaeger,
1965; Weinbaum et al., 1994)
9
M.L. Knothe Tate / Journal of Biomechanics 36 (2003) 1409–1424
Canaliculus wall to
osteocyte process surface
M.W. (g/mol unless otherwise
indicated)
Myoglobin
Saccharated iron oxide
particles
[14C]Inulin
Microperoxidase MP-11
(haem undecapeptide)
1.5 10 9 m (Simionescu and
Simionescu, 1987)
1.75 10 9 m (Adamson, unpublished
data)
1–10 10 9 m (Weinstein et al., 1967)
12,800 (Simionescu and Simionescu,
1987)
17,800 (Simionescu and Simionescu,
1987)
1.48 10 9 m (Pappenheimer, 1955)
2.0 10 9 m (Cowin unpublished),
5000 (Weinstein et al., 1967)
1800–1900
8.9 10
data)
Patent blue violet
1.13 10 9 m (Cowin unpublished)
1.08 10 9 m (De Bruyn et al., 1985)
4.0 10 8 m (Hughes et al., 1978)
6.4 10
data)
5 10
data)
10
10
m (Adamson, unpublished
m (Adamson, unpublished
‘‘Fluorescent dyes’’
Azure C
Biotin
Pyrophosphate
125
I-labeled antipyrine
Tritiated glycine
Glucose
Calcium ion (Ca2+)
Strontium isotope(85Sr)
4.3 10
data)
10
m (Adamson, unpublished
1630
1550
860
342
566 (Adamson, unpublished data)
336 (McDougall, 1970)
376 (Adamson, unpublished data)
300–400 (Greenwald and Haynes,
1969;Nachemson et al., 1970;
Knothe Tate et al., 1998)
277 (Adamson, unpublished data)
250 (Handagama et al., 1989)
235
188 (McDougall, 1970)
1.6–2.1 10 10 m (Adamson,
unpublished data)
8.4 10 10 m
139
87.62
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51
Cr-labeled EDTA
Na fluorescein
m (Adamson, unpublished
M.L. Knothe Tate / Journal of Biomechanics 36 (2003) 1409–1424
MP-9
MP-8
Ruthenium red
Sucrose, [14C]sucrose
10
1413
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M.L. Knothe Tate / Journal of Biomechanics 36 (2003) 1409–1424
Fig. 2. Confocal micrograph of a remodeling event on the periosteum
of a dog ulna, superimposed on a DIC image of the same field of view.
At a prior timepoint, osteoclasts carved out the area in which the
osteoblasts (Ob) are laying down osteoid (Os), i.e. unmineralized
matrix, which mineralizes over time and fluoresces strongly. Typically,
resorption occurs intracortically, whereby the osteocytes (Ot) are
thought to signal the osteoclasts and osteoblasts on resting surfaces.
Osteoclasts remove areas of bone, and the osteoblasts fill in those areas
with osteoid. A nascent osteocyte (Not) is apparent in the area of bone
recently filled in with osteoid (Specimen courtesy of Mitchell B.
Schaffler, Ph.D., Mount Sinai School of Medicine, New York, after
Knothe Tate et al., 2002).
‘‘actively involved with the maintenance of the bony
matrix, and their death is [typically] followed by
resorption of this matrix (Junqueira et al., 1995). In
addition, osteocytes are thought to be mechanosensors
in bone (Aarden et al., 1994; Burger and Klein-Nulend,
1999). Transmission of mechanical signals to the
osteocyte cytoskeleton via cell surface receptors (Wang
et al., 1993) can occur directly through the solid matrix
structure of the tissue (Goodship et al., 1979; Carter,
1987; Huiskes and Hollister, 1993; Burr et al., 1989;
Frost, 1983; Harter et al., 1995; Burger and KleinNulend, 1999) as well as indirectly via fluid pressure
(Thompson, 1936) and shear stresses (Ajubi et al., 1996;
Cowin et al., 1991; Weinbaum et al., 1994; Luo et al.,
1995; Duncan and Turner, 1995; Forwood and Turner,
1995; Turner et al., 1994; Smalt et al., 1997) imparted by
fluid moving through the lacunocanalicular system due
to load-induced fluid flow (Salzstein and Pollack, 1987a;
Knothe Tate and Knothe, 2000). Translation of
mechanical signals at the cellular level may further
involve triggering of integrin force receptors and/or
changes in the conformation of membrane bound
proteins (Berthiaume, 2000) that affect membrane
fluidity (Haidekker et al., 2000) and trafficking. In
addition to these mechanical signals, chemical signals,
modulated through diffusive, convective and active
transport mechanisms, are transported intracellularly
(Donahue, 2000) as well as through the extracellular
fluid in which the cells are immersed (Knothe Tate,
2001; Steck et al., 2003). The lacunocanalicular system
provides an ideal milieu for transfer of exogenous and
endogenous signals via mechanical, electrical and
chemical mechanisms (Kelly and Bronk, 1990; Bassett,
1966). The cell signaling pathways leading to release of
second messengers, transcription factors, and finally
gene expression are not yet fully elucidated and are the
subject of much current research (Klein-Nulend et al.,
1995; Johnson et al., 1996; Hung et al., 1996a, b; Jacobs
et al., 1998; Burger and Klein-Nulend, 1999; You et al.,
2000; Allen et al., 2000; Haidekker et al., 2000).
Hence, the interaction between the solid and fluid
phases of bone underlies much of its mechanical (e.g.
viscoelastic damping, elasticity) (Piekarski, 1973; Buechner et al., 2001), metabolic (e.g. molecular sieving,
storage and mobilization of calcium, hematopoiesis)
(Knothe Tate, 2001), and adaptive behavior. At a
system and organ level, salient flow pathways include
the vascular system through which the blood flows as
well as the medullary space through which marrow is
squeezed during physiological loading activity. The flow
of blood through the vascular system is an important
mechanotransduction agent for endothelial cells (refer
to Berthiaume for a review of these effects). The study of
marrow flow and its effect on osteoprogenitor cells is in
its nascency. Although these flow regimes are important
in their own right, the objective of this survey is to
review the current understanding of fluid–structure
interactions within bone, with particular emphasis on
interstitial fluid flow and its implications for osteocyte
mechanobiology.
2. Structural influences on the fluid flow environment
within bone
2.1. Origins of the cellular syncytium and
lacunocanalicular network
The lacunocanalicular network is not only the largest
reservoir for fluid within bone, but it is also the fluid
space in closest proximity to bone cells. Pericellular fluid
in the lacunocanalicular system is the coupling medium
through which mechanical forces are translated into
mechanobiological, biochemical, mechanochemical and
electromechanical effects at a cellular level. The
structure of the lacunocanalicular system is dominated
by the syncytium of cells contained therein, the majority
of which are osteocytes. In situ, the local mechanical and
biological environment of an osteocyte is defined by its
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M.L. Knothe Tate / Journal of Biomechanics 36 (2003) 1409–1424
morphological history as well as the metabolic state of
the tissue in which it is incorporated (Figs. 3 and 4)
(Holtrop and Weinger, 1972; Knothe Tate et al., 2002).
Osteocytes derive from osteoprogenitors, a fraction of
which differentiate into active osteoblasts (Fig. 2).
Osteoblasts synthesize and secrete collagen and other
organic components of the bone matrix, which is called
osteoid in the unmineralized state. Of the active
osteoblasts, a fraction become incorporated within the
newly laid down matrix (Menton et al., 1984; Junqueira
et al., 1995) and remain ensconced as osteocytes within
spaces called lacunae. Nascent osteocytes maintain
direct contact with the overlying bone lining cells and
Fig. 3. Osteocytic syncytium in a normal (left) and advanced stage
osteoporotic (left) human femur (after Knothe Tate et al., 2002).
Fig. 4. Portion of a cross section from the metacarpus (I) of a 180-dayold rat 10 min after procion red injection (left). The bone tissue shows
cortical structure. Tracer appears to delineate vascular pathways from
the intramedullary vessels (black arrow) to those of the endosteum
(black arrowhead) and the inner cortex (white arrowhead). Numerous
periosteocytic spaces show presence of tracer as well (original
magnification 120). Bone tissue of the metacarpus in a 60-day-old
rat 10 min after procion red injection (right). Adjacent metacarpi (III
and IV), exhibiting predominantly cancellous bone structure with
peripheral areas of undecalcified cartilage. Tracer appears to be
concentrated in areas of calcified tissue apposing vascular areas (white
arrows) (original magnification 75) (after Knothe Tate et al., 1998).
1415
osteoblasts, as well as with previous generations of
osteocytes through cell processes that are elaborated
before and during matrix synthesis (Fig. 2) (Menton
et al., 1984). In mature bone, the osteocyte body and its
processes are contained within spaces called lacunae and
channels called canaliculi, respectively. Derived from the
stellate shape of the osteocytes (Aarden et al., 1994;
Tanaka-Kamioka et al., 1998) and their interconnectivity, the lacunocanalicular system (LCS) is a conduit for
metabolic traffic and exchange (Cooper et al., 1966;
Copenhaver, 1964; Simionescu and Simionescu, 1987;
Knothe Tate et al., 1998). The extended osteocytic
network, comprising cells interconnected by multiple
cell processes that are joined at gap junctions (Doty,
1981), forms a ‘‘functional syncytium’’ (Johnson and
Highison, 1983; Martin, 2000; Knapp et al., 2001).
Thus, in addition to intercellular communication via the
gap junctions (Doty, 1981; Donahue, 2000), the cells
making up the syncytial network remain in contact via
their common environment that is defined by a
contiguous bone fluid space. What remains is a network
of cells, most of which are isolated physically from one
another while remaining connected to syncytial and
lacunocanalicular networks via cell processes and a
common fluid medium.
The lifecycle and health of individual osteocytes
influences the state of the cellular syncytium within
bone tissue. Osteocytes have an approximate average
half-life of 25 years (Frost, 1963), although their life
expectancy may be highly variable (Marotti et al., 1990).
The state of a given lacuna depends on the viability of
the osteocyte contained within it. As osteocytes lose
viability, their size and shape changes and their pyknotic
remains may persist within the lacunae for some time.
Thereafter, the remodeling cycle may be initiated to
remove nonviable cell remains and surrounding tissue or
a lacuna may remain empty, become mineralized and/or
lose its patency (Frost, 1960; Currey, 1964). Hence, at a
tissue level, the cellular syncytium and the common fluid
space defined by this syncytium are interrelated and
change, depending on the viability of individual cells
and the state of the tissue (Knothe Tate et al., 2002).
Whereas the local structure of the matrix dictates the
local mechanical environment of an osteocyte, the
periosteocytic fluid defines its local biochemical environment and the viscosity of this fluid influences the drag
forces imparted to the cell surface via fluid flow. Just as
remodeling activity and osteocyte osteolysis change the
local mechanical environment of the osteocyte, metabolic activity of these cells influences the biochemical
milieu of their surrounding fluid. The exact biochemical
composition and viscosity of this fluid is unknown due
to the practical difficulties of obtaining a sample size
sufficiently large for analysis. Hence, bone fluid is often
idealized as being analogous to interstitial fluid, which is
defined as that fluid ‘‘interposed between the plasma and
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the cellsy [with an] ionic composition similar to that of
plasma’’ (Aukland, 1984.). This idealization is inappropriate, given early work by Neuman and colleagues
in which important differences between bone extracellular fluid and plasma were shown; in particular the
concentration of K+ is much higher in bone fluid than
in plasma. Moreover, the amount of K+ (mM/l) in bone
extracellular fluid decreases with age and in states of
metabolic deficiency (Canas et al., 1969), providing a
basis for differences in ionic content of periosteocytic
fluid and the extravascular fluid bathing osteoprogenitor
cells and osteoblasts on bone surfaces (Rasmussen and
Bordier, 1974; Geisler and Neuman, 1969; Matthews
et al., 1978) (which is more directly in contact with the
vascular–extravascular interface). Variations in water
content have been documented as a function of species,
age and underlying pathology of the specimen under
examination (Timmins and Wall, 1977). Hence, the
lacunocanalicular network provides a microcirculatory
system for periosteocytic fluid that is distinct from the
blood plasma and lymph fluid.
2.2. Pathways of fluid movement through bone tissue
Although periosteocytic fluid flow is essential for bone
vitality, the pathways for influx from the blood supply
and efflux to the lymphatic system have not yet been
completely elucidated (Knothe Tate et al., 1998). In
cortical bone interstitial fluid originates from the
vascular system (Fig. 4. left) and is most likely formed
via filtration from the arterial end of Haversian
capillaries (Montgomery et al., 1988). Effluence is
hypothesized to proceed via the venous ends of
neighboring Haversian systems, by a prelymphatic
system leading through the matrix to the intial
lymphatics of the periosteum (Montgomery et al.,
1988), and/or by way of periosteal venules (McCarthy,
1997). The blood supply entering the Haversian system
is thought to derive primarily from the vessel system of
the medullary canal (Fig. 4A) and exit via the periosteal
blood supply (Cooper et al., 1966; Montgomery et al.,
1988). In cancellous bone, major pathways of influx and
efflux are presumed to be associated with the medullary
blood supply (Fig. 4, right). Path distances between the
medullary cavity and the extravascular space of a given
osteocyte are much shorter in cancellous bone
(McCarthy, 1997; Gatzka et al., 1999) than in cortical
bone (Knothe Tate and Knothe, 2000; Knothe Tate
et al., 1998).
In both cancellous as well as cortical bone, transudance of interstitial fluid from the blood supply to the
periosteocytic space occurs primarily via the lacunocanalicular system, and to a lesser extent via the
intermatrix porosity (Knothe Tate et al., 1998; Knothe
Tate, 2001). Starting at the source, the fluid passes first
through the basement membrane and fenestrae of the
capillary blood vessel epithelium (Landis and Pappenheimer, 1963; Cooper et al., 1966), the dimensions of
which are uncertain (in bone marrow vessel walls
epithelial fenestrae are circa 9–10 mm in diameter)
(Landis and Pappenheimer, 1963; Dillaman, 1984),
and enters the extravascular space. Prior to entering
the lacunocanalicular system (Fig. 1A,B), the fluid
passes through the layer of so-called bone lining cells,
or flattened osteoprogenitor cells, that line internal (i.e.
along the endosteum and Haversian canals) and external
(i.e. along the periosteum) surfaces of bone (Rasmussen
and Bordier, 1974; Maximow and Bloom, 1952; Van der
Weil et al., 1978; Miller et al., 1989) and may play a role
in maintenance of bone fluid composition, modulation
of the ion flux between vascular and extravascular
compartments (Miller et al., 1989; Baltadzhiev, 1994),
and/or regulation of interstitial fluid pressure (Hillsley
and Frangos, 1996). Thereafter the fluid enters the
canalicular openings (measured diameter 500–600 nm,
(Knapp et al., 2001) or the intermatrix porosity
(estimated average pore size 5–12 nm) of the bone
surface (Fig. 1C).
The lacunocanalicular system provides one conduit
for osteocytes to receive nourishment and to rid
themselves of waste products via extracellular transport
(Wasserman and Yaeger, 1965; Baud, 1968; Piekarski
and Munro, 1977; Knothe Tate et al., 1998). The LCS
flow volume comprises the roughly annular space
between the walls of lacunae and canaliculi and the
surface of osteocytes and their processes (postulated
dimension 14–100 nm (Cooper et al., 1966; Weinstein
et al., 1967). Furthermore, this space is likely to be
partially occupied by a molecular network, e.g. of
collagens and proteoglycans, that influence osmotic
pressure and flow conditions in situ. Flow conditions
within this annulus depend not only on the state of this
molecular network but also on the viscosity of the fluid
as well as morphological characteristics including the
surface roughness of the canalicular wall, the presence of
junctions between the cell surface and the canalicular
wall and/or fibril networks (e.g. proteoglycans) within
the fluid space, as well as physicochemical surface
interactions (Reilly et al., 2001). Based on atomic force
microscopy measurements of methyl methacrylate filled
casts of the lacunocanalicular system, canaliculi are 500–
600 nm in diameter. Their wall structure is dominated by
collagen fibrils that may be arranged regularly and form
ridges spaced approximately 100 nm apart (Knapp et al.,
2001). The canalicular wall is smooth, but the regular
dips and ridges caused by the collagen that lines the wall
are a source of roughness which may influence shear
stresses imparted by the fluid on the cell surface as well
as mixing of solutes within the lacunocanalicular system.
Finally, the collagen lining may affect molecular charge
interactions between the fluid and bone matrix (Reilly
et al., 2001; Anderson and Eriksson, 1968).
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2.3. Alterations in fluid movement due to ‘‘structural’’
changes in environment
Bone provides a dynamic flow environment that is
influenced by ‘‘structural’’ changes at the organ, tissue
(e.g. changes in macroscopic porosity), matrix (e.g.
degree of matrix hydration), cellular (e.g. state of the
syncytium), and molecular levels (presence of molecules
that influence permeability). At an organ level, the bone
lining cell layer that covers resting surfaces of bone is
postulated to serve a boundary function, gating the
degree to which fluid and solutes enter and exit bone. At
a tissue level, macroscopic changes in porosity or
vascularization have a profound effect on flow regimes.
At a matrix level, the solid phase of bone tissue acts like
a molecular sieve (Knothe Tate et al., 1998; Tami et al.,
2003) with low-pass filtering function, whereby molecules larger than approximately 40,000–70,000 Da are
not transported through the lacunocanalicular system
without fluid flow (Tami et al., 2003). At a cellular level,
it has been suggested that osteocytic process exert a
plugging action at the junction between the canaliculus
and lacuna, resulting in a valve mechanism favoring
efferent over afferent flow with respect to the cell body
(Arnold and Frost, 1971). Subcellularly, if the osteocyte
surface is tethered to its surroundings (You et al., 2001)
like leukocytes and neutrophils during rolling, it is
reasonable to expect that changes in number or patterns
of tethers, resulting from trauma or disease, would
influence shear stresses (Tami et al., 2003) imparted by
the fluid through drag force. Finally, it has been
suggested that the inherent ‘‘hydraulic conductivity’’ of
bone matrix is affected by both the microarchitecture of
bone tissue (Mishra and Knothe Tate, 2003) as well as
by the incorporation of molecules such as hyaluronan
and chondroitin sulphate into the matrix. Hence,
‘‘structural’’ changes in bone that are manifested
through increases in fluid space or incorporation/
exclusion of molecules in the matrix may influence the
global and local flow field through bone; this reiterates
the concept of the fluid medium serving a coupling
capacity within bone through mechanobiological, biochemical, and electromechanical interactions at a
cellular level.
3. Endogenous and exogenous mechanisms of fluid flow in
bone
Several biophysical and electrochemical mechanisms
have been implicated as motive forces for fluid in bone.
These involve endogenous mechanisms including active
transport by osteocytes,1 hydraulic conductivity effects,
1
Bundles of microfilaments that line the surface of the osteocyte and
fill its cell processes (King and Holtrop, 1975; Aarden et al., 1994) are
1417
pressure gradients inherent to pulsatile pressure and
osmotic pressure, as well as exogenous mechanisms
associated with mechanical loading of bone and effects
of electromechanical and acoustic energy. Regardless of
the specific mechanism for translation of biophysical
and electrochemical effects to the cellular level, the
pericellular fluid serves as a coupling medium.
3.1. The concept of hydraulic conductivity
Hydraulic conductivity, defined as volumetric fluid
flux per unit pressure drop, is an inherent property of
bone tissue (Neuman, 1969; Neuman and Neuman,
1980; Peterson et al., 1985; Bushinsky et al., 1989;
Hillsley and Frangos, 1996; Hui et al., 1996) and plays a
dominant role in establishing baseline levels of endogenous fluid flow through bone, i.e. excluding fluid flow
induced through exogenous effects such as pressure
gradients due to mechanical loading (Mishra and
Knothe Tate, 2003). It is a function of i.e. tissue
architecture and porosity, matrix biochemistry, and
pericellular fluid properties and is modulated by the
bone lining cells or surface osteoblasts. (Neuman, 1969;
Neuman and Neuman, 1980; Peterson et al., 1985;
Bushinsky et al., 1989; Hillsley and Frangos, 1996; Hui
et al., 1996; Mishra and Knothe Tate, 2003). In cell
culture experiments using osteoblasts, hydraulic conductivity has been shown to be affected by a number of
agonists including calcitonin and parathyroid hormone
(Hillsley and Frangos, 1996). Similarly, tissue permeability is affected by the presence of specific osteotropic
agents including hyaluronidase and chondroitin sulphate (Otter et al., 1988; Guzelsu and Walsh, 1990;
Guzelsu and Regimbal, 1990; Hillsley and Frangos,
1996). Given that parathyroid hormone has been shown
to stimulate the production of hyaluronan by osteoblasts in culture (Midura et al., 1994) and in situ
(Noonan et al., 1996), an increase in hydraulic
conductivity in response to PTH treatment (Hillsley
and Frangos, 1996) may be influenced by the increase in
this macroporous glycosaminoglycan.
3.2. Flow via pressure gradients
3.2.1. Pressure gradients generated by mechanical
loading
Mechanical loading of bone results in a tissue stress
state comprising cyclic dilatational and deviatoric
components. The dilitational component is responsible
for the development of fluid pressure and the ensuing
(footnote continued)
thought to aid in active intracellular transport (King and Holtrop,
1975; Civitelli, 1995) and may actively pump fluid through the
extracellular space through contraction and expansion of the cell
processes (Aarden et al., 1994).
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fluid flow through deformation of the fluid-filled
lacunocanalicular and intermatrix porosities within
bone tissue.
Biot’s theory of poroelastic solids (Biot, 1955)
describes the behavior of fluids in porous materials
and provides the biophysical basis for load-induced fluid
flow in bone. According to this theory, compression
deforms the solid matrix of a porous material, instantaneously increasing the pressure in the fluid within the
pores. Disparate pressures between the interior and
exterior of a porous solid cause a net flow of fluid. In
turn, fluid flow out of the porous solid causes the solid
matrix around the pores to relax. Concomitant to
matrix relaxation around the pores, fluid pressure within
the pores decreases until it equilibrates with surface
pressure of the solid, at which point there is no pressure
gradient to drive fluid flow. Removal of load results in a
pressure gradient as well, moving fluid back into the
sample until the pressure gradient reduces to zero once
again. The rate of fluid pressure reduction during matrix
relaxation is an exponential decay function (Biot, 1955)
(refer to Table 2 for other parameters related to
poroelastic behavior of bone).
Hence, bone tissue is analogous to a stiff and dense,
fluid-filled sponge. Bassett was one of the first to
describe the concept of mechanical load-induced fluid
flow in bone (Bassett, 1966, 1968). Thereafter, Piekarski
and Munro applied Biot’s theory of poroelasticity to a
model of the lacunocanalicular system and postulated
that load-induced fluid flow increases perfusion between
the blood supply of the Haversian canal and osteocytes
(Piekarski and Munro, 1977). Many researchers have
developed analytic and computer models to simulate
fluid flow phenomena in idealized model systems of
bone tissue (Salzstein and Pollack, 1987a, b; Keanini
et al., 1995; Steck et al., 2000, 2001) and the
lacunocanalicular network (Johnson et al., 1982; Johnson, 1984; Pollack et al., 1984; Petrov et al., 1989;
Kufahl and Saha, 1990; Weinbaum et al., 1994; Cowin
et al., 1994; Mak et al., 1997; Knothe Tate and Niederer,
1998; Wang et al., 1999; Knothe Tate, 2001). The
existence of mechanical load-induced fluid flow has been
proved recently through visualization of fluid displacements induced through controlled mechanical loading of
cortical and trabecular bone (Gatzka et al., 1999;
Knothe Tate and Knothe, 2000; Knothe Tate, 2001).
3.2.2. Pressure gradients generated by venous and
intramedullary pressures
In addition to mechanical load-induced pressure
gradients in bone, it is expected that endogenous
pressure gradients affect movement of fluid through
the tissue. Streaming potential measurements indicate
that the heartbeat causes transcortical pressure gradients
(Kelly and Bronk, 1990; Otter et al., 1990). Furthermore, disparities between capillary and intramedullary
pressure may provide pressure gradients that drive
interstitial fluid flow through bone. In 1896 Starling
formulated the law that increased capillary pressure
increases transudation through tissue (Starling, 1896).
Given the fact that medullary pressure is influenced
primarily by venous resistance in the tissue, venous and
intramedullary pressure are necessarily interrelated
(Bier, 1915; Willans and McCarthy, 1986). A number
of experiments have been conducted to examine these
effects. Application of a low-pressure venous tourniquet
to the tibiae of growing dogs has been shown to increase
fluid movement from the capillaries to the interstitial
Table 2
Constants related to bone as a poroelastic material
Diffusion constants
Diffusion constant: glucose through cartilage 1.4 10 6–2.3 10 6 cm2/s (Maroudas, 1968; Maroudas et al., 1968)
Free diffusion coefficient of glucose at 37 C: 8.8 10 6 cm2/s (Gladden and Dole, 1953)
Diffusion constant of glucose through cortical bone, based on in vitro experiments: 10 8 (Amprino, 1952)
Extrapolated diffusion constant of glucose through cortical bone: 3 10 9 cm2/s (Lang et al., 1974)
Free diffusion constant for albumin: B9 10 7 cm2/s
Constants related to bone as a poroelastic material
Porosity, f (porosity connected to the free surface of the sample): 0.2 (Karnovsky, 1967)
Permeability (m2), k
10 12–10 14 (Johnson et al., 1980)
Adult canine bone: 3.32 10 7 cm3/(mm min g/cm2) (Li et al., 1987)
Puppy bone: 20.83 10 7 cm3/(mm min g/cm2) (Li et al., 1987)
Time constant (s)
Haversian system: 10 4 (Johnson and Highison, 1983)
Lacunocanalicular system (cleared of debris and filled with water): 10 3 (Johnson and Highison, 1983)
‘‘Bone’’: order of 1 s (Kufahl and Saha, 1990)
Fluid flow rates
11.4–400 ml/g per hour (McCarthy and Lang, 1992)
600 ml/g per hour, based on permeability constants from Li et al. (1987) and McCarthy and Lang (1992)
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space of bone tissue (McDougall, 1970; Kelly and
Bronk, 1990). In contrast, short term (i.e. hours, days)
studies applying femoral vein ligation showed no
significant influence on fluid movement from the
capillaries to the interstitium (McCarthy, 1997). Nonetheless, in longer term studies (weeks) it could be shown
that venous ligation increases bone marrow pressure
significantly in the hindlimb of a tail-suspended rat,
causing a concomitant and lasting induction of interstitial fluid flow through the ligated bone (Bergula et al.,
1999).
3.2.3. Pressure gradients generated by osmotic gradients
Given the differences in composition and concentration between the extravascular, intracellular and intravascular fluid compartments, it is assumed that
osmotic pressure due to concentration gradients also
plays a role in influx and exudation of fluid from one
compartment to the next in bone, as is the case in
cartilage and encapsulated organs. By controlling the
flow of solutes through the tissue, local osmotic pressure
gradients build up and influence the flow of fluid
through the tissue. Arnold and Frost have suggested
that fluid flow through bone tissue occurs by way of a
‘‘osteocyte actuated osmotic pump’’ (Arnold and Frost,
1971; Frost, 1973), whereby water enters and leaves the
system through the LCS as well as the microporosity of
the bone matrix. Recently, it has been shown in vitro
that osmotic stress affects the viscoelastic and physical
properties of articular chondrocytes (Guilak et al.,
2002). Hence, in addition to the role of osmotic pressure
in bulk flow regulation and subsequent swelling
pressures, it is expected that local osmotic stresses affect
the conformation of the osteocyte cytoskeleton.
3.3. Electromechanical and acoustic energy effects
Electromechanical and acoustic (Knothe Tate, 2001;
Simionescu and Simionescu, 1987) energy have been
applied clinically to improve healing of fractures (Rubin
et al., 2001) and nonunions (Kristiansen et al., 1998), as
well as to ameliorate bone quality in osteoporotic
patients (Warden et al., 2001). In addition to hypothesized cellular ‘‘triggers’’ that may be turned on by
imparting such energy, it is assumed that energy
application causes changes in fluid flow that play a
putative role in stimulation of anabolic activity. This
is an emerging field of translational research, the
underlying mechanisms of which are currently being
elucidated.
Electromechanical behavior per se is an endogenous
characteristic of bone. Collagen confers piezoelectric
behavior to bony and tendonous tissue (Bassett, 1966,
1968; Anderson and Eriksson, 1968; Fukada and
Yasuda, 1957); just as deformation of bone tissue
produces an electrical field, application of an electrical
1419
field results in mechanical deformation or strain
(Fukada and Yasuda, 1957). In addition, bone exhibits
electromechanical behavior related to flow of extracellular fluid through the charged matrix (Lanyon and
Hartman, 1977; Johnson et al., 1980; Gross and
Williams, 1982; Otter et al., 1985); these are referred
to as streaming potentials. In concurrent studies of
piezoelectric and streaming potential effects in wet bone
samples, potentials caused by interstitial fluid flow
dominate over those caused by piezoelectric effects
(Huiskes and Hollister, 1993; Goodship et al., 1979).
Application of electrical and electromagnetic energy to
bone could conceivably alter fluid flow regimes within
bone, thereby affecting concentration gradients and
shear stresses imparted to the cells. In addition,
application of such energy may affect cell membrane
permeability as well as trans- and intracellular calcium
fluxes. Hence, streaming potentials and other electromagnetic effects are recognized as further potential
mechanisms of transduction in bone (Bassett, 1966;
Chakkalakal, 1989; Weinbaum et al., 1994; Cowin et al.,
1994; Knothe Tate, 2001).
In order to apply exogenous signals to induce flow
through bone, it is necessary to understand the intrinsic
electromechanical behavior of bone tissue, e.g. the
degree to which the mechanical load-induced fluid flow
and the electromechanical streaming potential coincide.
There is controversy regarding whether streaming
potentials in bone occur exclusively via fluid flow
through the lacunocanalicular system, through the
matrix micrporosity or through a combination of both.
Experimental studies in wet bone specimens have
targeted the collagen–hydroxyapatite porosity of the
bone matrix as the likely site for streaming potentials in
situ (Salzstein and Pollack, 1987a, b). In contrast, some
theoretical models of fluid flow assume that streaming
potentials occur exclusively within the bone canaliculi
(Cowin et al., 1994). The crux of the issue pertains to
mobility of bone fluid in situ (i.e. to what degree it is
bound to the mineral crystal and collagen) (Eriksson,
1974) as well as the state of the walls through which the
fluid flows. Experimental studies suggest that a fraction
of the water within bone exists in the free liquid state
and that the remaining water is chemically bound to
collagen and apatite, existing in at least three different
bound states (Eriksson, 1974). Load-induced fluid flow
has been shown experimentally to occur through the
intermatrix porosity as well as the lacunocanalicular
system (Tami et al., 2003). Based on recent atomic force
microscopy studies, the walls of canaliculi may be at
least partially sheathed by a collagen layer (Reilly et al.,
2001; Knapp et al., 2001) that would be expected to
insulate the flow of the charged fluid from the charge of
the canalicular wall (Guzelsu and Walsh, 1990).
Whereas data supporting the idea of ‘‘bound’’ water
within the matrix pores lend credence to the idea that
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the bone canaliculi, rather than the micropores within
the matrix, are the site of streaming potentials, flow
visualization and AFM observations strengthen the
argument that the matrix microporosity plays an
additional role in the development of streaming
potentials as well.
4. Summary and future directions
A review of the current understanding of fluid–
structure interactions and fluid flow in bone delineates
myriad roles for fluid flow, from the level of the skeletal
system down to the level of osteocyte mechanobiology.
At a systemic level, the vascular/extravascular interface
allows exchange of fluids between the skeleton and other
organ systems, which is critical for maintenance of
metabolism and hemapoiesis. At a tissue level, the
common fluid space defines the environment of osteoclasts, osteoblasts and osteocytes comprising the functional cellular syncytium in bone. Furthermore, bone
tissue health depends on the viability of the cells
comprising the living component of the tissue. The
system comprising the bone cell syncytium and the
lacunocanalicular fluid space provides a high and low
pass filter for mechanochemical signal reception and
processing via intercellular and intracellular means
(Dodd et al., 1999, Knothe Tate, 2001; Tami et al.,
2003). Local fluctuations in fluid composition (i.e.
concentration gradients) within this common fluid space
determine the local biochemical milieu of the cells. In
addition to its role as a carrier of solutes, fluid flow
provides an indirect mechanism to impart loads,
incurred through physiological activity, via shear
stresses to the surface of cell membranes and the
cytoskeleton.
Conceptually, bone fluid serves as a coupling medium
through which energy is transferred from the system to
the cells that have the machinery to remodel the tissue,
thereby providing an elegant feedback mechanism for
functional adaptation. Much like, in the Earth’s atmospheric flow environment, the wind is harnessed and
transferred by the blades and gears of the windmill into
a form of energy useful for human work, the interaction
between pericellular fluid and bone structure, from a
system to an organ, tissue, matrix, cellular and
molecular level, provides an optimal feedback system
for transfer of energy and signals from the system that is
exposed to the dynamic, external environment to the
cells that have the machinery to adapt to these changes
through internal remodeling. In sum, bone structure
provides a dynamic flow environment for cells. In
flowing through the fluid space of bone, pericellular
fluid acts as a carrier of mechanical, electrical and
chemical energy/signals between the systemic circulation
and bone tissue and cells. It provides an extracellular
network for communication between osteocytes, osteoclasts and osteoblasts. Thus, the flow of this fluid
provides redundant mechanisms for mechanochemical
transduction in bone, the organization and modulation
of which are the subject of much current scientific
exploration.
With growing technological advancements in the
fields of molecular and cell biology as well as sensors
and imaging, in situ observation of flow and its effects
on bone cells will be possible in the near future,
providing a new basis of understanding for this
important aspect of bone physiology. Eludication of
the mechanochemical transduction mechanisms related
to fluid flow in bone will provide a basis for applying
biomimetic principles for engineering of bone replacement tissue, treatment of orthopaedic disorders, e.g.
fractures, osteoporosis, osteolysis, and osteonecrosis, to
restore optimal function, and development of innovative
strategies to mitigate effects of microgravity, i.e.
osteopenia, in space exploration and inhabitation.
Acknowledgements
The author would like to extend thanks to Ron
Midura, Ph.D. (Department of Biomedical Engineering,
Lerner Research Institute, Cleveland Clinic Foundation) for his valuable advice and constructive criticism of
the biological and biochemical aspects of this manuscript. In addition, the author expresses her gratitude to
R. Iwan Alexander, Ph.D. (Department of Mechanical
Engineering and Aeronautics, Case Western Reserve
University; NASA Microgravity Research Center) for
discussions regarding microscopic and submicroscopic
fluid flow regimes and gravitational effects on the
same. During preparation of this manuscript, MKT
was supported by a grant from the Swiss National
Science Foundation 823A-056609 and the Orthopaedic
Research Center of the Lerner Research Institute,
Cleveland Clinic Foundation. Portions of this work
were supported by the EMDO foundation.
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