INFLUENCE AND EFFECTS OF DC
ELECTRIC FIELDS ON BONE CELLS
DISSERTATION
submitted to the Faculty of Electrical Engineering and
Information Sciences at the Ruhr-Universität Bochum to
receive the grade of Doctor of Engineering
Presented
by
Miguel Miron Mendoza
From
Ciudad Mendoza, Mexico
Bochum, Germany 2003
To Palmira
and
To all the Indigenous people of Mexico, for their fighting and
struggling for the acceptation of their rights and recognition as a
fundamental part in the history of Mexico. Particularly to the
Zapatistas (ZapatistaNational Liberation Army, EZLN), for their
courage, dignity, resistance and struggle.
ii
ACKNOWLEDGMENTS
•
I would like to express deep gratitude to Prof. Martin Hofmann for his total support
and assistance to gain my PH. D grade. Furthermore I am also very grateful for his
valuable collaboration in the writing of this work.
•
I would like to thank Prof. Gerhard Schiffner for taking great care in reading my
manuscript and his constructive criticism.
•
I thank Prof. David Jones for the use of all facilities to carry out my experimental
research in his laboratory. Furthermore I am also very grateful to all lab colleagues,
particularly to Marita Kratz, Cornelia Klein, Thorsten Pohl, Eckahrd Bröckman and
mi amigo Weng Tan for their valuable assistance in my experimental research, but
mainly for all moments we shared together during that time.
•
My deep gratitude goes to Catrin Davies and Timo Rieger for their valuable help and
support, but mainly for all nice time we have shared together and their sincere and
appreciated friendship. Special thanks to Miss Davies for improving the style of this
manuscript.
•
My heartfelt thanks go to Michael Mederos for his technical support in my
experimental work with calcium measurements.
•
I would like to thank all the people that contributed to my enjoyment of life in
Germany. Particular thanks go to Margaret and Manfred Jan, my Underwater Rugby
team and Latinos group.
•
I am very grateful to the Mexican people that through National Council for Science
and Technology of Mexico who made possible for me to receive a scholarship for the
realization of this work.
•
My whole gratitude goes to my family, for its total support, but mainly for its
complete love and fond.
iii
ABSTRACT
The aim of this work was to study the effects of DC electric fields on bone cells,
namely osteoblasts, and the mechanism of interaction between them. To date no one has been
able to answer this question. The solution to this phenomenon that is undergone daily in our
skeletal system will aid in developing techniques to help heal bone diseases such as
osteoporosis, pseudoarthrosis, and non-union fractures among others.
The present work utilized a newly developed technique, traction force microscopy, to
study the effects of DC electric fields on cells through mechanical forces. This new technique
was coordinated along with phase and fluorescence microscopy to characterize the physical
behavior of cells under DC electric field exposure. By combining the physical evaluation of
cells with biochemical analysis, it was possible to conclude that cells adapt to a tensional
force generated by DC electric fields. The adaptation is carried out through a process of
retraction and elongation, which causes each cell to become orientated perpendicularly to the
direction of the electric field lines. Adaptation process is carried out in 5 steps:
1. Firstly DC Electric fields exerted a tensional force on the cells at the membrane
zones exposed directly to the positive and negative poles of the electric field.
2. The tensional force caused an increase of calcium ions in the cytoplasm of the cells.
3. The increase of calcium ions caused a reorganization of the cytoskeleton. Cells
retracted their sides exposed directly to the positive and negative poles of the electric
field, exactly where the tensional force had been visualized.
4. Cells underwent a loss of traction force with the substratum at both sides exposed
perpendicularly to the direction of the field lines.
5. Cells carried out an active re-extension of both their long sides exposed
perpendicularly to the direction of the field lines, exactly where the loss in traction
force had been visualized.
The present work did not manage to answer the question of “What is the mechanism
of interaction between DC electric fields and osteoblasts”, but contributed to this field of
work with new results that opened further debate into this area of research. It suggests a new
theory to explain the mechanism exerted by a DC electric field to cause the re-orientation of
the cells. It proposes that cells re-orientate themselves to counteract and adapt to a tensional
mechanical force dependent on the changes of transmembrane potential generated by the
application of an external DC electric field. Future work would involve carrying out the same
experiments with different cells, to determine whether the results obtained were exclusively a
property of bone cells. Followed by an evaluation on whether the tensional mechanical force
generated by a DC electric field is exerted either at the membrane, or at the extracellular
matrix of cells. Finally, implementation of an in vivo system will provide a physiological
basis for future development of bone pathology therapies and more effective devices of
stimulation.
iv
KURZFASSUNG
Das Ziel der vorliegenden Arbeit war es, die Effekte von elektrischen Gleichfeldern
auf Knochenzellen, im speziellen Osteoblasten, und die daraus resultierenden
Interaktionsmechanismen zu erforschen. Bisher konnten noch keine detaillierten Antworten
gefunden werden. Die Lösung dieses Phänomens, das täglich auf unser Skelettsystem
einwirkt, wird helfen neue Techniken zu entwickeln, um Knochenkrankheiten wie z.B.
Osteoporose, Pseudoarthrose, Knochenbruch und andere zu heilen.
In der vorliegenden Arbeit wurde eine neu entwickelte Technik, „die
Traktionskraftmikroskopie“, benutzt, um die Effekte von elektrischen Gleichfeldern
hinsichtlich mechanischer Kräfte in Zellen zu untersuchen. Diese neue Technik wurde mit der
Phasenkontrastmikroskopie und der Fluoreszenzmikroskopie zur Charakterisierung des
physikalischen Verhaltens von Zellen unter Gleichfeldwirkung kombiniert. Bei der
Kombination zwischen physikalischer Evaluierung von Zellen und biochemischer Analyse
war es möglich, zu dem Schluss zu kommen, dass die Zellen durch Spannungskräfte, die
durch elektrische Gleichfelder erzeugt wurden, ausgerichtet werden. Diese Ausrichtung wird
durch einen Prozess von Retraktion und Elongation erreicht. So orientiert sich jede Zelle
senkrecht zu den Feldlinien. Die Zellen führen diesen Prozess in 5 Schritten aus:
1. Zunächst wird eine Spannungskraft in den Zellen an der Zellmembran, die direkt den
elektrischen Feldlinien ausgesetzt sind, durch elektrische Gleichfelder erzeugt.
2. Die Spannungskraft bewirkt einen Anstieg der intrazellulären Kalziumkonzentration.
3. Der Kalziumanstieg führt zu einer Reorganisation des Cytoskelettes. Die Seiten der
Zellen, die direkt den elektrischen Feldlinien ausgesetzt sind, retrahieren sich genau
da, wo die Spannungskräfte beobachtet wurden.
4. Die Zellen verlieren die Verbindung mit dem Substrat, auf dem sie adhärent waren, an
den Seiten, zu denen sie senkrecht zu den Feldlinien ausgerichtet waren. Dies führt zu
einem inneren Spannungskraftverlust.
5. Die Zellen dehnen sich senkrecht nun zu den Feldlinien aus. Und zwar genau dort, wo
der Verlust der Spannungskraft beobachtet wurde.
Auch diese Arbeit konnte keine abschließende Antwort auf die Frage bringen, was der genaue
Mechanismus in der Interaktion zwischen elektrischen Gleichfeldern und Osteoblasten ist,
aber die erzielten neuen Ergebnisse können zu weiterer Diskussion auf diesem
Forschungsgebiet beitragen. Die neue Theorie, die den Mechanismus der „Reorientierung“
der Osteoblasten beschreibt, soll hierzu anregen. Sie geht davon aus, dass die Zellen sich
selbst reorientieren durch Entgegenwirken und Anpassen an die mechanischen
Spannungskräfte. Dies ist abhängig von den Änderungen der transmembranen Potentiale, die
durch die Einwirkung von elektrischen Gleichfeldern erzeugt wurden. Weitere
Forschungsarbeit mit anderen Zellenarten wird in Zukunft nötig sein, um zu erforschen, ob
die gezeigten Veränderungen nur bei Osteoblasten auftreten. Weiterhin stellt sich die Frage,
ob die durch elektrische Gleichfelder erzeugten mechanischen Spannungskräfte an der
Zellmembran direkt oder in der Extrazellulärmatrix ansetzen. Darüber hinaus würde ein invivo-System eine physiologische Basis für zukünftige Behandlungsstrategien hinsichtlich
Erkrankungen des Knochens liefern.
v
PUBLICATIONS
•
How do DC electric fields affect cells? Cellular responses under electric fields depend
on voltage amplitude, time of exposure to the electric field and are dependent on
extracellular calcium. Miron, M. and D. B. Jones. Presented at the World Congress on
Medical Physics and Biomedical Engineering, IEEE. July 23-28 2000. Chicago
Illinois, USA.
•
Cellular Orientation on Osteoblasts Under Electric Fields Depends on the Calcium and
Potential Distribution at the Membrane. M. Miron and D. B. Jones. Presented at the
20th Annual Meeting of the Society for Physical Regulation in Biology and Medicine.
January 10-14, 2001. Charleston, South Carolina, USA.
Student work awarded with a Travel Award by the Program committee
•
Force Distributions Across Osteoblasts Caused by Electric Fields Produce Actin
Disruption, Remodeling and Orientation: A mechanism Regulated by Calcium. M.
Miron, S. Curtze and D. B. Jones. Presented at the 21th Annual Meeting of the Society
for Physical Regulation in Biology and Medicine. January 30-February 1, 2002 San
Diego, California, USA.
Student work awarded with a Travel Award by the Program committee
•
Dynamic Changes in Traction Forces with DC Electric Fields in Osteoblasts-like
Cells. Sami Curtze, Miguel Miron. Micah Dembo, and David B. Jones. Journal of Cell
Sciences. Accepted.
vi
INDEX
LIST OF ABBREVIATIONS AND PARAMETERS
x
INTRODUCTION
1
I THEORICAL CONSIDERATIONS
1.1. Anatomy and Physiology of Bone
1.101. Bone Remodeling
1.101a. Activation
1.101b. Resorption
1.101c. Reversal
1.101d. Formation
1.102. Aspects of Bone Remodeling
1.103. Mechanotransduction in Bone
1.2. Electrical Properties of Bone
1.201. Endogenous Electrical Activity in Bone
1.202. Exogenous Electrical Activity in Bone
1.3. Electrical Properties of Bone Cells
1.301. The Membrane
1.302. Electric Fields at the Membrane Level
1.302a. The Potential Difference Across the
Compact Layer (∆χ)
1.302b. The Diffuse Layer Potential (φGC)
1.302c. The Surface Potential (φ)
1.302d. Resting Potential (VREST)
1.302e. An Endogenous Electric Field, Emem
1.302f. The Electric Field Undergone by Proteins
at the Membrane Et
1.303. Polarization and Dispersion of the Cell Membrane
1.304. Deformation of the Cell Membrane
1.305. Cytoplasm and the Extracellular Medium
1.4. Effects of Modifying the Natural Electric Field
1.401. Distribution of Membrane Potential
1.402. Electrophoresis in the Plasma Membrane
1.403. Electro-osmosis in the Plasma Membrane
1.404. Summary of Effects of Applying an Electric Field
II MATERIAL AND METHODS
2.101. Introduction to Cell Culture and Biochemical Work
2.102. Preparation of Bone Cells
2.103. Electric Field Chamber
2.104. Phase Contrast Microscopy
2.105. Fluorescence Microscopy
2.105a. Calcium Imaging
2.105b. Membrane Potential Measurements
2.105c. Visualization of Actin Filament Dynamic
2.106. Traction Force Microscopy
5
8
8
8
8
9
10
11
14
14
17
18
19
20
22
22
22
22
23
23
23
24
25
26
26
28
30
31
34
35
37
37
40
41
43
44
46
vii
2.106a. Preparation and Characterization
of Flexible Substrates
2.106b. Calculation of Traction Forces
2.107. Analysis of the Calcium Role
2.107a. Thapsigargin
2.107b. Manganese
2.107c. Cadmium
2.107d. Lanthanum
2.107e. Nifedipine and Nitrendipine
2.108. Actin Filament Disruption
2.108a. Latrunculin A
2.108b. Cytochalasin B
2.109. Myosin Light Chain Kinase Inhibition
2.109a. ML-7
2.109b. Wortmannin
2.110. pH Measurements
47
48
48
49
49
50
50
50
50
50
51
51
51
51
52
III RESULTS
3.1. Behavior of Osteoblasts Under Electric Field Exposure
54
3.101. Retraction
55
3.102. Retraction and Elongation
57
3.103. Alignment and Orientation
60
3.104. Summary of Cellular Responses Caused by DC Electric Fields 62
3.105. Measurement of Temperature in Medium
63
3.106. Measurements of pH in Medium, Electrophoresis and
Electro-osmosis
63
3.107. Traction Force Microscopy
65
3.107a. Experiment Control of Traction Force Microscopy
70
3.108. Alignment Was not Programmed in the Cells
70
3.108. Summary of the Behavior of Osteoblasts Under Electric
Field Exposure
70
3.2. Biochemical Evaluation of Osteoblasts Under Electric Field Exposure 72
3.201. Alignment Is Dependent on Actin Polymerization
72
3.202. Membrane Potential Changes
74
3.203. Calcium Measurements
75
3.203a. Characterization of Calcium Responses (Peak Value) 77
3.203b. Manganese
80
3.203c. Thapsigargin
82
3.203d. Summary of Calcium Responses in Osteoblasts
83
3.203e. Calcium and Latrunculin
84
3.203f. Cadmium
86
3.203fa. 10 µM Cadmium
86
3.203fb. 25µM Cadmium
88
3.203fc. 50 µM Cadmium
90
3.203fd. 100 µM Cadmium
92
3.203fe. Summary of Cadmium Experiments
94
3.203g. Nifedipine and Nitrendipine
95
3.203h. Lanthanum
99
3.203i. Summary of Calcium
101
3.204. Myosin Light Chain Kinase
101
3.205. Alignment Is Dependent on Ca2+
103
viii
IV DISCUSSION AND CONCLUSION
4.1. Summary of the Behavior of Osteoblasts Under DC ELF Exposure
4.2. Analysis of Results
4.201. Cytoskeletal Contractility Defined Retraction of Osteoblasts
4.202. Contractile Forces Caused Increases of Ca2+ in Cytoplasm
4.203. Re-orientation was Dependent on Calcium
4.204. Alignment Was Only Dependent on Actin
4.205. Elongation Was only Possible in Low Tension Zones
4.3. Conclusion of Results
4.301. What Is New in this Work?
4.302. Future Perspectives
105
106
107
109
111
112
113
115
116
118
APPENDIX
Solutions and Reagents Used
120
REFERENCES
123
CURRICULUM VITAE
138
ix
LIST OF ABBREVIATIONS
Cytochalasin B
DC ELF
Epidermal Growth Factor Receptor
Fetal Calf Serum
GFP
Latrunculin A
Myosin Light chain Kinase
Nifedipine
Nitrendipine
Polyphosphoinositodes
SGPs
TFM
VSCC
CB
Direct Current Electric Field
EGFR
FCS
Green Fluorescent Protein
Lat A
MLCK
Nif
Nit
PIPs
Stress-Generated Potentials
Traction Force Microscopy
Voltage-Sensitive Calcium Channel
TABLE OF PARAMETERS
Parameter
Capacitance
Charge
Charge density
Concentration
Conductivity
Current
Current density
Electric field
Frequency
Force
Length
Mass
Permittivity
Resistance
Resistivity
Temperature
Time
Velocity
Viscosity
Voltage
Symbol
C
q
ρM
(…)
σ
I
J
E
ƒ
F
L
M
ε
R
ρ
T
t
v
η
V
Name of Unit
Farad
Coulomb
Couloumb per square meter
Mol
Siemens per meter
Ampere
Ampere per square meter
Volts per meter
Hertz
Newton
Meter
kilogram
Farad per meter
Ohm
Ohms meter
Degree Celcius
Second
Meter per second
Newton second per square meter
Volt
Symbol for Unit
F
C
C/m2
M
S/m
A
A/m2
V/m
Hz
N
m
kg
F/m
Ω
Ωm
o
C
s
m/s
N s / m2
V
TABLE OF PREFIXES
Factor
Name
Prefix
109
giga
G
106
mega
M
103
kilo
k
10-3
milli
m
10-6
micro
µ
10-9
nano
n
10-12
pico
p
10-15
femto
f
x
INTRODUCTION
Electric fields have been well established to modulate bone physiology (1-4).
However, despite a multitude of studies of bone-derived cell cultures with electric fields (58), little is known about the mechanism exerted by the cells to sense an electric field, how
they convert the information into a biochemical signal, and generate the appropriate
physiological response. Therefore further research is required to determine this interaction
process between electric fields and bone cells. This was exactly the objective that motivated
the present work “to identify the possible pathway(s) of interaction between electric fields and
bone cells”. Establishment of the exact nature of electric field transduction would not only
solve a fundamental mystery in the field of bone research, but would provide valuable
benefits in the management of bone diseases with electrical therapy.
The approximately 206 bones of the human skeleton, apart from serving as a mineral
reservoir, give the body structural support, protection and locomotive capacity. Every bone is
a dynamic tissue; it undergoes a constant process of reformation. Bones renew themselves in
order to enable the growth, repair, reinforcement and resorption, adjustment of the bone to
stress, calcium ion regulation in the body, and everyday maintenance necessary for a healthy
skeletal and body (9-11). Thus old bone is removed constantly and new bone is created
according to the needs of every bone. This process of renewal of bone, known as bone
remodeling is responsible for keeping the strength of bone throughout our life (9-11), if it
fails, damages result. To activate bone remodeling, bones have to detect the changes of load
in their mechanical environment and generate the appropriate changes in bone architecture
(11, 12). However, the question of how bones sense the mechanical loading has not been
solved yet. A possible theory is through potential differences generated by the mechanical
deformation of bone (13, 14); when bone is mechanically stressed, the bending forces create
pressure gradients that make that bone fluid flows across all canals in the bone, and since
bone fluid is composed of ions, when it flows, it causes a movement of charges in the
direction of flow, thereby giving rise to an electrical current, which, in turn also gives rise to a
potential difference at two different bone sites. The potential difference or voltage measured
is known as stress-generated potential, SGP (13, 15-19). Thus, SGPs have been postulated to
be the signal controlling the response of bone as a mediator of the mechanical loading (13, 14,
19). However, it has not been demonstrated yet, how bone cells can detect those electrical
signals and generate the appropriate physiological responses.
Bone consists of concentric layers of matrix surrounding longitudinal vascular
channels (9, 10). The difference in ion concentration between its different layers generates
differences in electrical potential across or along the bone surface. Therefore bone is also a
tissue electrically dynamic. Thus, when a pair of electrodes are inserted between two different
bone sites, a potential difference or voltage is measured (20, 21). The voltage measured is
usually known as bioelectric or steady-state potential (22). Bioelectric voltages produce
electrical current densities within the range of 0.5 to 12 µA/cm2 (22, 23). Interestingly, the
areas of active growth and repair are electrically negative when measured and compared with
less active areas in bone, or with respect to a reference point (21-23). Furthermore, when bone
is injured or fractured, the entire bone becomes electrically negative, and a peak value of
voltage is measured exactly at the fracture site (21, 23). The electrical currents produced by
the voltages due to the lesion are now called injury or fracture currents, and their densities
increase to approximately 100-130 µA/cm2 (23). Thus, based on these findings, it was
postulated that the endogenous bioelectricity of bone might play a role during bone growth,
repair and healing (22-24).
1
Convinced of the bioelectricity in bone, many laboratories predicted that an external
electrical stimulus would stimulate bone formation. Thereby it was demonstrated that
implanting weak electrical current directly into the bone increased bone formation (25-30).
New bone (callus) formed around the cathode and the optimal bone growth took place with
currents between 5 and 20 µA (25, 27-30). These results seemed consistent with the values of
fracture currents measured, and also with the electrical negativity at the fracture and growth
site measured. Therefore, from all this research, electrical stimulators were developed as a
rehabilitative tool for complete bone repair and growth. By the early 1980’s, electrical
stimulation was firmly established as a treatment for the management of nonunions. To date,
the interest in methods of accelerating bone healing with electricity persists; new techniques
of electrical stimulation have been developed and all have been proven to work (31-35); thus,
electrical stimulators are now used worldwide as a supplemental form of therapy to help
enhance the body’s bone healing process (36-39). However, despite an abundance of
undeniably valid data, from both basic and multicenter clinical trials, demonstrating the
effects of electrical stimulation on skeletal biology and healing, little is known about the
signal transduction that mediates the physiological response of bone to electrical stimulation.
As consequence, a certain amount of skepticism remains among biologists and clinicians, due
to the fact sometimes the method used is not the appropriate one and can thus bring collateral
damages. Therefore scientists need to learn more about how electricity interacts with bone,
thereby it will provide more effective electrical stimulators and prevent principally secondary
damages. Until recently, it was only believed that such alterations in the electrical
environment of bone are perceived by bone cells which in turn initiate a chain of intracellular
events that lead to an adaptive bone formation. However it has not been proved to date. To try
to solve such a dilemma, a multitude of in vitro studies involving bone cells and electrical
stimulation have been performed over the last 20 years. As electrical stimulation has been
used electric and magnetic fields, direct current and ultrasound among others (5-8, 40-43).
Bone cells have responded in different manners that are considered key in the process of bone
formation. Therefore it has not been questionable if bone cells respond to electrical
stimulation, but what is the mechanism that is acting on the cells, and how it is transduced
into biochemical signals for generating an appropriate physiological response.
In light of all research done with bone and electricity, there is enough evidence to say
that electric fields exist into bone and play a role in bone physiology, and that external
electrical stimulation can also alter profoundly many cellular events and modulate the bone
physiology. It leaves no doubt in the relation between bone and electricity. However, until the
mechanism of interaction between electric fields and bone cells can be identified, little
advance can be made in the management of bone diseases with electrotherapy. Bone diseases
that could benefit from this include: osteoporosis (a fail in the bone remodeling process),
pseudoarthrosis (failed fusion of bone growth), delayed fracture healing and non-union
fractures among others. Therefore, the knowledge of the exact nature of interaction between
bone cells and electric fields would not only solve a fundamental mystery in the field of bone
research, but would provide a physiological basis for future development of bone pathology
therapies. Thus all this basic research may add up to more effective devices and ubiquitous
applications.
The work presented here had both purposes: a) to solve a mystery in bone physiology;
and b) to contribute to the fight against bone diseases. To achieve this, experiments were
divided into three objectives:
1. To evaluate the physical behavior of bone cells when exposed to DC electric fields.
2
2. To investigate if a DC electric field stimulates mechanical changes in the cells.
3. To identify and determine the possible signal transduction pathway(s) that causes the
changes in the behavior of bone cells when subjected to DC electric fields.
To achieve these objectives, a new technology was incorporated into this work. This is
the first example of experimental work that utilizes traction force microscopy to evaluate the
mechanical behavior of bone cells (and any other cell) when exposed to DC electric fields.
Other techniques such as phase contrast and fluorescent microscopy were also coordinated to
gain details and obtain a whole overview of the behavior of the cells when exposed to these
fields. Furthermore, diverse experiments were carried out to identify the possible biochemical
transduction pathways responsible for generating the response in cells. New results were
found that open a new perspective in the process of interaction between cells and electric
fields (44). The findings point towards a new hypothesis to explain the behavior of cells when
subjected to DC electric fields.
For clarity, this work was divided into four chapters. The first chapter gives a general
introduction to the anatomy and physiology of bone, and describes the electrical properties of
bone and bone cells. This chapter has two purposes: a) to outline the relationship between
bone and electricity and; b) to describe the possible effects caused on bone cells by the
application of an electric field. The second chapter details the materials and methods used
during this research project. It describes the process of obtaining bone cells and all techniques
that were used to analyze the behavior of cells when subjected to DC electric fields. The third
chapter describes the results; it was divided in two sections: the first section was the
identification of the physical behavior demonstrated by the cells when subjected to DC
electric fields. The physical behavior was visualized at the same time with phase contrast,
fluorescent and traction force microscopy to obtain an overall characterization of events. The
second phase was the identification of the possible signaling mechanism responsible for
causing the observed changes in the behavior of the cells. In this phase biochemical
experiments were carried out. The fourth and final chapter is the discussion and conclusion. It
compares the results obtained with results previously reported; it mentions the new findings
and opens new theories, leading to a final conclusion of this work as well as proposals for
future research in this field.
3
I
THEORICAL
CONSIDERATIONS
INTRODUCTION
Despite the evidences that demonstrate that bone is an electrically dynamic structural
tissue (16-24), little is known about the signal transduction that mediates the physiological
response of bone to electric fields. The purpose of this chapter is to revise those electrical
properties of bone and their possible role in bone physiology. A theorical revision of how a
DC electric field may act at the cellular level is also described. To achieve these purposes, the
chapter was divided into 3 sections: firstly it examines the anatomy and physiology of bone;
the second section revises the electrical properties of bone and their possible role in bone
physiology; the last section revises the electrical properties of single bone cells.
4
1.1. ANATOMY AND PHYSIOLOGY OF BONE
The adult human skeleton has approximately 206 bones (9). Bones gives the body
structural support and protection, locomotive capacity, and serve as a reservoir for minerals,
especially calcium and phosphorus. Bone also harbors the bone marrow, where blood cells are
formed. The bones of the body fall into four general categories: long bones, short bones, flat
bones, and irregular bones (45). At the macroscopic level the human skeleton consists
predominantly of two types of bone: cortical bone, also called compact or dense bone; and the
spongy bone, also called cancellous or trabecular bone (10). A specific bone usually consists
of varying amounts of compact bone surrounding varying amounts of trabecular bone (figure
1). Compact bone is dense and hard. It serves as a protective outer shell covering and
surrounding every bone in the body. Therefore it has primarily mechanical and protective
functions, which is also responsible for the skeleton's strength. Compact bone represents
nearly 80% of the skeletal mass (9). Spongy bone is less dense and more elastic than compact
bone. It is composed of an irregular three-dimensional array of vertically and horizontally
orientated bundles of various widths and lengths of strands, plates, rods, arches and struts of
bone fused together known as trabeculae (10). Trabeculae make bones look very porous,
similar to a honeycomb (full of tiny holes). Spongy bone is present in most bones, generally
within the confines of a cortical bone shell, however it only represents 20% of the skeletal
mass (9). Both cortical and trabecular bone contain bone marrow. Bone marrow fills the
spaces of the pores in the spongy bone; in cortical bone it is found in the marrow cavity.
Marrow contains special cells that are responsible for the production of blood cells. However
only certain bones like breastbone, spine, skull, hips, and ribs contain blood-forming marrow
(9, 10).
Figure 1. A schematic representation of a bone. All bones in the skeleton are composed with different
amounts of cortical and trabecular bone (Gray’s Anatomy, ref. 45).
All bones are lined on both external and internal surface by layers of connective tissue
called periosteum and endosteum respectively (figure 2). Periosteum consists of an outer layer
of collagen fibers and cells. The inner, more cellular layer of the periusteum is composed of
flattened cells with the potential to divide and differentiate into bone cells (osteoblasts). These
flattened cells are also called osteoprogenitor cells and play a prominent role in bone growth
and repair. The endosteum lines all internal surfaces of bone and it is also composed of
osteoprogenitor cells and a small amount of connective tissue. Thus endosteum is
considerably thinner than periusteum. The principal functions of periosteum and endosteum
5
are nutrition of osseous tissue and provision of a continuous supply of new osteoblasts for
repair or growth of bone (9, 10). In the present work, osteoblasts were obtained of periosteum
pieces of bovine ulnae and steer radii. Chapter 2 will give the details of the process to harvest
the cells.
Bone is a complex living tissue; it is composed of cells, vessels and mineralized
extracellular matrix (45). The extracellular matrix has two principal ingredients, a protein
matrix (collagen) and calcium phosphate crystals (hydroxyapatite). Collagen provides the
basic structure into which hydroxyapatite is deposited; this gradually hardens into the protein
mesh forming bone. Collagen fibers and calcium phosphate crystals are arranged in flattened
plates known as lamellae that are variously oriented so as to give bone its strength (10). At
the microstructural level both trabecular and compact bones have a lamellar organization.
Trabecular bone is composed of a collection of more or less parallel lamellae. In compact
bone the lamellae may be organized either in parallel fashion or concentrically in quasicylindrically shaped structures called osteons or harvesian systems (9). The Haversian
system or osteon forms the basic unit of cortical bone (figure 2 and 3). Osteons are long
bifurcated cylinders (figure 2B). They are typically about 200 µm in diameter and about 1 or
2 cm long with a central canal known as harvesian canal that contains blood vessels, nerve
fibers and loose connective tissue (figure 2B, 2C and 3). The Haversian canal is from 20 to
150 µm in diameter. The variation in diameter can be due to the age of the osteon and its
relative position within the bone (10). One or two small blood vessels occupy the canal along
with a nerve and possibly a lymphatic vessel. The harvesian canal is surrounded by concentric
lamellae. These structures composed of collagen and tiny calcium phosphate crystals are
about 3 to 7 µm in width. Osteons consist from between four to about twenty lamellae. Lying
between or within the lamellae are special small holes called lacunae. Each lacuna provides
enough space for an individual bone cell (osteocytes) to reside. The osteocyte inside the
lacuna is responsible for secreting the bone salts surrounding it. Osteocytes within their cavelike lacunae have many cytoplasmic processes. These processes are approximately 15 mm
long and are arrayed three-dimensionally in a manner that permits them to interconnect with
similar processes of up 12 neighboring osteocytes (9, 45). These processes lie within passages
called canaliculi. The outer perimeter of the osteon has a special coating of mineralized
matrix that is known as a cement line (figure 2B and 2C). The harvesian canals communicate
with the marrow cavity, with the periusteum, and with each other through transverse or
oblique Volkmann’s canal (figure 2). Volkmann’s canals do not have concentric lamellae.
Instead, they perforate the lamellae (45).
Bones undergo a constant process of reformation (11). Bones renew themselves
throughout life in order to enable the growth, repair and maintenance necessary for a healthy
skeleton and body. This constant dynamic renewal of bone is responsible for bone strength.
Old bone is removed (resorption) and new bone is created (formation). During childhood and
the beginning of adulthood, bone becomes larger, heavier and denser; bone formation is then
more important than bone resorption (9, 10). During the first three decades of our life we also
deposit calcium, phosphate and other minerals in our bones. The Bone Mass actually
increases until the age of 25-30 where it reaches its maximum value: the peak bone mass
(maximum bone density and strength). The higher the Peak Bone Mass is, the lower the risk
of fracture and osteoporosis is later in life (11). Osteoporosis is characterized by reduced
bone mass and is thought to result from an imbalance between bone formation and resorption
(11). Bone mass remains stable for a few years; a perfect equilibrium between bone formation
and resorption exists yet. After a certain age (about 40), bone mass starts to decrease (10). For
women, this bone loss starts a few years before menopause and becomes more and more
important till death. Bone loss occurs at an average rate of 0.5% every year. The process of
6
resorption and formation in bone is the role of two different cells that remove old bone and
replace it with new. The technical term for this process is 'The Bone Remodeling Cycle', and
the cells that are responsible for this recreation and removal of bone are the osteoblasts and
osteoclasts. Osteoblasts are involved in bone formation and the osteoclasts are involved in
bone resorption (11).
Figure 2. A schematic picture of the wall of a long bone. Observe 4 types of lamellar bone: harvesian
system or osteon, outer and inner circumferential lamellae, and interstitial lamellae. The protruding harvesian
system in A shows the orientation of collagen fibers in each lamella. Picture B (longitudinal) and C (cross
section) are of an osteon system showing lamellae, a central blood capillary, and many bone cells (osteocytes)
with their processes (modified from Gras´s anatomy, ref 45).
Figure 3. Cross section of a harvesian system. Image is a high photomicrograph of a harvesian system
or osteon within compact bone. At the center of the osteon is the Haversian canal (HC) containing blood vessels.
Osteons are composed of circular layer known as lamellae (LA); Osteocytes (OC) embedded in the Lacunae
(LAC) from adjacent lamellae communicate through narrow channels or canaliculi (CAN) in the bony matrix.
VC is the Volkmann’s canal and CL is the cement line. Picture was taken in the School of Anatomy and Human
Biology in the University of Western Australia.
7
1.101. Bone Remodeling
Bone remodeling is the process by which bone is deposited, resorbed and formed
through precisely controlled functions of osteoblasts, osteoclasts and their associated
activation factors and cofactors (11). Remodeling is involved in bone growth, reinforcement
and resorption, changes in bone shape, adjustment of the bone to stress, bone repair, calcium
ion regulation in the body. Remodeling keeps the strength of bone as well as mineral
homeostasis by transferring calcium and other ions into and out of bone. Approximately 25%
of trabecular bone and 3% of cortical bone is renewed annually in the mature adult, and this
remodeling activity decreases with age (9). At any one time, about 10% of available bone
surfaces are undergoing remodeling. The events in the bone remodeling cycle have been
described as activation, resorption, reversal and formation (figure 4).
1.101a. Activation
Activation is the initial event in the bone remodeling cycle. The bone surface changes
from the resting state to an activated state. The activated state signals mononuclear precursor
cells (pre-osteoclasts) to migrate to the remodeling site at the bone surface. Pre-osteoclasts
cells originate in the bone marrow. They are released as monocytes into the bloodstream and
collect at sites of bone resorption, where they fuse together to form differentiated,
multinucleated osteoclasts. Although the specific factors are not known, microdamage or
microfracture might predispose a site to be selected for remodeling. It is thought that
osteoblasts are the responsible to induce the formation of osteoclasts (46).
1.101b. Resorption
During the resorption phase of the remodeling cycle, the osteoclasts attach to bone and
seal it off from the extracellular space. Osteoclasts begin to remove both the organic matrix
and the mineral component of bone by the secretion of bone-resorbing enzymes and
subsequent acidification by the release of protons into the bone-resorbing compartment.
Thereby osteoclasts are capable of excavating a tunnel into the old bone, forming cavities that
are then invaded by other cells. Osteoclasts act at a rate of about 50 µm per day. In trabecular
bone, the cavity created is called a Howship's lacuna, which is a saucer-shaped cavity about
40 to 50 µm deep. In cortical bone, the cavity created is called a "cutting cone", which is a
cylindrical tunnel about 2.5 mm in length and 150 to 200 µm in diameter, parallel to the long
axis of the bone. On average, the resorption phase takes about 33 days in trabecular bone and
about 23 days in cortical bone (10).
1.101c. Reversal
During reversal phase resorption ceases and osteoclasts detach from the bone. This
phase reflects a transition period during which the critical coupling of bone formation to bone
resorption occurs. The mechanisms responsible for this coupling remain unclear. The fate of
osteoclasts also remains unclear. It is possible that they apoptose (i.e. programmed cell death)
or they may undergo fission to return to the mononucleated state. This phase of the
remodeling cycle is the least well understood. The reversal phase lasts for about 9 days in
trabecular bone and about 4 days in cortical bone (10).
8
1.101d. Formation
During the formation phase repair begins (11). The walls of the tunnel become lined
with a layer of osteoblasts (figure 4). Osteoblasts appear in the cavities to smooth off the
resorbed surface and deposit a cement-like substance (bone matrix synthesis) that binds the
new bone to the old. At the same time a blood capillary grows down the center of resorption
tunnel bringing the nutrients to the bone cells to survive. The freshly formed substance
deposited by osteoblasts (consisting chiefly of type I collagen strands) form the osteoid: spiral
fiber of bone matrix. It is rapidly converted into hard bone matrix by the deposition of
calcium phosphate crystals (hydroxyapatite). Osteoblasts cause calcium salts and phosphorus
to precipitate from the blood; these minerals bond with the newly formed osteoid to
mineralize the bone tissue. To produce the osteon, osteoblasts lay down concentric layers of
new bone, which gradually fill the cavity, leaving only a narrow canal (harvesian canal)
surrounding the new blood vessel. Many of the osteoblasts become trapped in the new bone
matrix and, at this point, the original bone-forming cells, are referred to as osteocytes. The
osteocyte has no opportunity to divide, although it continues to secrete further matrix in small
quantities around itself. The osteocyte occupies a lacuna in the matrix. From each lacuna
radiate tiny channels (canaliculi) where cell process from the resident osteocyte enables it to
form junctions (gap junctions) with adjacent osteocytes (figure 2b and 3). The remodeling
sequence is complete and that particular site returns to a quiescent state (11).
Osteoblasts need about 150 days to refill the resorption cavity (9). Osteoblasts
deposite layers of matrix at a rate of 1 to 2 µm per day. After an interval of about 25 days in
trabecular bone and about 35 days in cortical bone, mineralization of osteoid begins under the
control of osteoblasts. After the mineralization process starts, the mineral content of the
matrix increases rapidly (over the first few days) to about 75% of its final mineral content. It
may, however, take up to 1 year for full mineralization of the matrix (10). At the same time as
some tunnels are filling up with new bone, others are being bored by osteoclasts, cutting
through older concentric systems (45). The tunnel functions as a route of access to osteoclasts
and osteoblasts; besides as it contains one or more blood vessels, it brings also nutrients to the
bone cells in order for them to survive.
Figure 4. A schematic picture of bone remodeling. Osteoclasts resorb a discrete area of mineralized
bone matrix. Osteoblasts enter the tunnel behind them, lining its walls, and then begin to form new bone matrix
(osteoid). A blood capillary grows down the center of the resorption tunnel bringing nutrients to the bone cells to
survive. Osteoblasts become trapped in the new bone, and then mature into terminally differentiated osteocytes.
(Design based in ref. 11).
9
1.102. Aspects of Bone Remodeling
In the skeleton of the normal adult, bone formation occurs only where bone resorption
has already occurred (11). Normally, because the resorption of old bone by osteoclasts and the
subsequent formation of new bone by osteoblasts are closely linked, the two processes are
coupled. Thus, bone resorption and bone formation are not random and independent events;
they occur sequentially as part of the turnover mechanism replacing old bone with new bone.
It is thought that the coupling of resorption followed by formation can be mediated by factors
produced during resorption. Following the completion of bone resorption, these factors recruit
and activate osteoblasts for the next stage of remodeling bone formation. What is the
mechanism activating osteoblasts to migrate to the resorption site? This is unknown yet.
When a bone breaks, the fracture breaks the continuity of the bone and of important
attached soft tissues including blood vessels, which spill their contents into the surrounding
tissue. The body automatically seeks to repair the injury. Bone healing occurs in three distinct
but overlapping stages: 1) the early inflammatory stage; 2) the repair stage; and 3) the
remodeling stage. Firstly inflammatory cells rush to destroy, dilute or isolate invaders and
injured tissue (45). Tiny new blood vessels called capillaries begin to grow into the site.
Surrounding cells proliferate. New tissue bonds the fractured bone ends with a soft callus, a
mass of connective tissue and exudate (matter escaped through blood vessel walls). Finally
remodeling begins. Within a few months, a hard callus replaces the soft one. Remodeling
restores the inner canal. Once restoration is completed, which may take years, the healed area
is brand new, without a scar. Usually thicker, the new bone may be even stronger than the old
one, thus, if the bone should break again, it is unlikely to be at the same place (9).
The skeleton, despite constant remodeling, provides a rigid framework whose
dimensions scarcely change (11). This is partly because the parts of a bone are renewed not all
at once but little by little, rather like a building whose bricks are replaced one at time. Besides
such conservatism in the mode of renewal, active homeostatic mechanisms are at work. Thus
a bone, like the body as a whole, is a dynamic system, maintaining its structure through a
balance between the opposed activities of a variety of specialized cells. Any dynamic system
poses a problem of stability, and this lead us to a general question about maintenance of body
structure. It has been previously described how new cells are produced in a controlled fashion
to replace those that are lost, and how the extracellular matrix is remodeled and renewed. But
why do the different types of cells not become progressively jumbled and misplaced? Why
does the whole structure not sag, warp, or otherwise change its proportions as new parts are
substituted for old? How does a fracture detect whether to activate its repair? What gives the
order to trigger the differentiation of the cells? What is responsible for activating the
migration of cells to the remodeling site? What controls the remodeling process? Bone
remodeling is not fully understood yet. While much is known about the kinetics of bone
turnover at the cellular level, little is understood about the local regulation of bone remodeling
at the molecular level. Presumably, specific factors regulate each step of a remodeling cycle,
the coordination of remodeling with the local differentiation of osteoclasts and osteoblasts
from their respective precursor cell population, and the integration of bone remodeling with
the endocrine control of bone turnover (11, 46). It is very important to understand these
various levels of control because deregulated bone remodeling lies at the basis of many
metabolic bone diseases, such as osteoporosis or fractures that do not heal. Although
osteoporosis has been associated with endocrinological imbalances such a reduced estrogen,
how these systematic factors influence the local regulation of bone remodeling is still not
known.
10
The integrity of bones depends on their use (9). If bones provide mechanical support
and are the site of muscle attachment for movement, then in order for bones maintain their
mass, grow, repair, or renew, they need to be stimulated mechanically. This simple conclusion
suggests that the bones should have the ability to respond to changes in their mechanical
environment to renew themselves. In 1892, the anatomist Julius Wolff proposed that
mechanical stress is the responsible for determining the bone architecture. Wolff related into a
specific mathematical law the changes in the form and the function of bones to the changes in
mechanical stress acting in them; subsequently this law has become known as "Wolff's Law"
(12). Wolff stated simply that a bone grows in response to mechanical stress so as to produce
an anatomical structure best able of resisting the applied stress. This proposal formulated that
mechanical stress determines the form and function of bone has become profoundly accepted
(47-52). In support this theory, experimental studies have shown the response of bones to
weightlessness during spaceflight (47, 48). Loss of bone is common in astronauts during
extended spaceflight. On the other hand, increased stress on bone, as occurs in elite athletes in
high-impact sports such as gymnastics, results in increased bone (48). However, despite the
phenomenon of bone gain or loss with changes in skeletal loading is well established (49-52),
the mechanisms by which bone senses load and adjusts to it are not so clear. What actually is
the stimulus and what are the sensors? Which are the target cells? How do the sensors
communicate the message to the cells, and by what pathways do the cells respond? None of
these questions have been answered accurately. There are many theories on how bone may
sense the mechanical loading, however the mechanism by which bone senses the information
in order to initiate the adaptive response remains yet unsolved. Therefore, establishment of
the exact nature of load transduction signals would not only solve a fundamental mystery in
the field of bone research, but would provide a physiological basis for future development of
bone pathology therapies.
1.103. Mechanotransduction in bone
Loading of bone can come of external forces applied to the body or internal forces
generated by the muscles and connective tissue. Thus, when loads are applied to bones, for
example, during normal activity such as walking and running where the various parts of the
skeleton are subjected to mechanical forces that vary with time and place, bones have to adapt
to the changes in load applied to keep serving their functions. This process is known as bone
adaptation, and if it fails, damages result (49). Therefore bone adaptation requires that bones
detect the amount of mechanical loading and generate the appropriate changes in bone
architecture. Research over the past 30 years has attempted to discover what triggers this
process of functional adaptation in accordance with Wolff´s law; however it remains unclear
yet. The first mystery is the nature of the signals controlling the response of bone to
mechanical loading. Is the mechanical load directly the signal? Are other signals controlling
the response of bone? The second mystery is what biological system in bone senses those
signals and how is this information transmitted to cells to make new bone or destroy old bone
(bone remodeling).
To try to solve such a dilemma, the first step should be to identify the physical signals
generated by mechanical loading in bone, which could be the responsible for triggering the
functional adaptation. There are five primary modes of loading that can be imposed on bone:
bending, compression, tension, shear, and torsion (9). When loads are imposed on bone, all
cause an elastic deformation of bone (49). This is because the forces between the atoms and
molecules making up the bones act like tiny springs, stretching and compressing in response
to the applied force. Thus as result, when a force is applied, the bone changes its length (49).
In mechanics, the length changes of objects in response to an applied force are defined as
11
strain. Therefore, the bone deformation is also quantified as strain (49); thereby strain (S) is a
measure for the change in bone length, and is defined as the change in length, ∆L, divided by
the original length, L.
S = δL / L
(1)
Where δL = ∆L – L
∆L is the new length and L is the original length.
Strain is dimensionless, however it is reported as units of microstrains, which respond
to the percentage of change in length; 10000 microstrains respond to 1% change in length. By
convention negative strain values are referred to compressive loading, whereas positive strain
values are referred to tensile loading.
Bone is rarely subjected to pure bending, pure compression, pure tension, pure shear,
or pure torsion, but instead “combined loading”. In reality bone is subjected to a combination
of compression and tension. For example, when physiologic bending forces are applied to a
bone, tensile forces are generated on the convex surface and compressive forces are generated
on the concave side (figure 5). A continuous gradient of stress distribution from tension to
compression exists through the bone’s cross section. Therefore, a bending force will cause a
portion of bone to act in tension while another portion acts in compression.
Figure 5. Schematic representation of a bending force imposed on bone. The top portion of the bone is
elongated (∆L > L), and the bottom portion of the bone is shortened (∆L < L). Thus, the top is under tension and
the bottom is under compression.
Bone is strain rate sensitive and the magnitude, orientation, and distribution of strains
encountered by bone during functional acivities are deemed to be extremely important in
controlling bone architecture (49, 53). Bone tends to be more strain rate sensitive than other
biological tissues. To give an idea how sensitive bone is to strain, compare it to other tissues
such as skin and muscle. The mechanical loads on skin and muscle can normally result in
strains up to 120 % on the cells. In bone the mechanical loading results in strains up to 0.5%
(at which point bone begins to fracture), but more typically 0.3% (10). This places the straindetecting mechanism(s) in bone at among the most sensitive.
Since strains are produced always that bone is mechanically loaded, it was firstly
postulated that mechanical strain could be the signal controlling the response of bone to
mechanical loading (51, 53). However it has not been demonstrated yet.
When bone is loaded, the resulting compressive strain squeezes the interstitial fluid,
which causes the fluid pressure to rise, that in turn causes the fluid to flow to regions of lower
pressure (54). Therefore, a pressure gradient drives the fluid flow from regions of
compression to tension in bone when it is mechanically deformed as in figure 5 (49). Bone
12
cells (osteocytes) live in fluid-filled hollows within the bone matrix (lacunae figures 2 and 3)
and are interconnected by fingerlike extensions through microscopic tunnels (canaliculi
figures 2-4). Strain-derived flow of interstitial fluid through the canaliculi and along the
osteocytes processes (cell membrane) is thought to mechanically activate the osteocytes (54,
55). The narrowness of the canalicular annulus ensures that even the minute physiological
bulk strains in bone produce considerable fluid shear stress over the osteocyte cell “finger”
(55). This flow of fluid over the cell surface, besides being crucial to renewing nutrients to
cells, subjects the cell to fluid-induced drag forces known as fluid shear stresses. Therefore it
has been also postulated that fluid shear stress could be the signal controlling the response of
bone to mechanical loading (54, 55).
Bone fluid immersed in the diverse bone channels is composed of nutrients and ions as
Ca2+ and Na+ among others, when it flows, it causes a movement of ions in the direction of
flow. Therefore when compressive strain causes bone fluid to flow from regions of high
pressure to lower pressure between two sites in the bone tissue, the charge is convected with
the fluid, giving rise to a electrical current between both sites in the bone tissue (15-17). This
electrical current, known as streaming current gives rise to a potential difference between the
same two bone sites, which is know as streaming potential (16, 17). Therefore when
electrodes are placed on two different bone sites and then bone is deformed, a voltage is
measured. The potential differences or voltages generated when bone is deformed by the
application of a mechanical load are called stress- or strain-generated potentials, SGPs
(56). SGPs have also been postulated to be the signal controlling the response of bone to
mechanical loading (13, 14).
In conclusion, there are three physical variables that are generated when a mechanical
load is applied to bone tissue: mechanical strain, shear flow and stress generated
potentials. Each of these tissue-level effects of mechanical loading probably plays some role
controlling the response of bone. To solve such dilemma, the next step should be to identify
the biological system that senses those physical variables, then isolates and subjects it to
similar magnitudes of mechanical strain, shear flow, and voltages as those thought to exist in
vivo.
Bone is a structural dynamic living tissue; it modulates its shape in response to
changes in load. Thus if bone undergoes remodeling when subjected to mechanical load, then
the process of bone remodeling appears to be governed by a feedback system in which the
bone senses the state of strain and either adds or removes bone as needed to maintain the
strain within normal limits. In the simplest analysis, some component of the bone itself may
be considered to have the function of sensing the signals generated by the mechanical load.
Figure 2, 3, and 4 show that the only living portion of the bone is the population of the bone
cells. Osteocytes are immersed in the lacunae; osteoblasts are covering the walls of the
harvesian canal; and osteoclasts are excavating new tunnels. In the adult human skeleton,
only 1% of the bone surface is covered by osteoclasts (9, 11, 46). Thus the greatest bone
surface is covered by osteocytes and osteoblasts. Therefore the biological living system in
bone that could be responsible for sensing the signals generated by mechanical loading are the
osteoblasts or osteocytes. Osteocytes were osteoblasts that became trapped in the bone matrix
and that have undergone a final differentiation (section 1.101d, figure 4). However osteocytes
remain connected with osteoblasts each other via long slender cell processes that connect by
means of gap junctions across canaliculi, thereby allowing for biochemical signal
communication between cells (figure 4). Since osteocytes represent the great majority of the
cells of bone tissue and are the only type of bone cells that are encased in the mineralized
bone matrix, they seem to be the cells responsible to sense the mechanical loading. However
13
because osteocytes are embedded in hard matrix and are post-mitotic (come of osteoblasts),
they are difficult to study. Therefore, research with osteoblasts-like cells (which may contain
possibly osteocytes) and mechanical strain, shear flow or electric fields, is the only possibility
to provide valuable information in the search of the biological system that senses the
mechanical loading in bone.
Experimental studies have shown that osteoblasts in culture respond to mechanical
strain (51), shear flow (54), and electric fields (5-8). However, the amount of mechanical
strain needed to activate bone cells in culture is 10-100 times larger than in vivo bone strains
(14, 49). It now appears that the most important tissue-level effect in bone physiology is the
fluid flow. It may be reasonable since the mineralized bone matrix is not directly attached to
the plasmalemma membrane of bone cells; bone fluid is placed between both. Are shear flow
or stress generated potentials the signals responsible for generating the appropriate response
from bone cells? Are only the stress generated potentials the signal? How do bone cells sense
those difference potentials? What is the process of mechanotransduction in bone cells?
Mechanotransduction is the conversion of a biophysical signal into a cellular response. It
includes the initial physical stimuli that must be sensed by the cells and then be converted into
a biochemical signal, which is transmitted through intracellular signaling to generate the
appropriate physiological responses. The process of mechanotransduction in bone cells with
shear flow or under electric field exposure is poorly understood. Thus more research is
required to try to determine this process.
1.2. ELECTRICAL PROPERTIES OF BONE
To evaluate the effectiveness of electrical stimulation in bone cells, it is important to
understand the electrical properties of bone. Bone has been found to be a source of
endogenous electrical currents when it is mechanically stressed or injured (16-24); and
second, it has been demonstrated that bone formation and repair can be modified or
stimulated by exogenous electrical currents (25-29). The purpose of this section is to
familiarize with those electrical properties of bone to understand the electrical stimulation of
bone and bone cells in culture.
1.201. Endogenous Electrical Activity in Bone
When bones are mechanically stressed, they generate potential differences or voltages
(56). At one time it was thought that bone might be piezoelectric (57), and that the
mechanically produced electrical signals were the stimulus that produced bone growth and
remodeling according to Wolff's law. The piezoelectric effect is the production of electrical
polarization in a material by the application of mechanical stress. The electrical polarization
results by the displacement of charges within the material. Piezoelectric materials also display
the converse piezoelectric effect (mechanical deformation upon application of electrical
charge). Piezoelectricity in bone was discovered by Yasuda and Fukada (58); subsequently
many others researchers also verified the piezoelectricity in bone (56, 59-61). The discovery
of piezoelectricity in bone aroused great interest because it seemed to provide an important
key in understanding bone physiology. Thus piezoelectricity became a candidate for the
process of bone adaptation to mechanical loading. The source of piezoelectricity in bone has
been attributed to the nonsymmetrical collagen fibers in bone (56, 62). However, although
bone exhibits small but definite piezoelectric properties, actually piezoelectricity is believed
to be insignificant in living bone, as piezoelectricity does not explain the amplitude and time
dependence of potentials measured in fluid-saturated bone (14, 17, 19). In effect, the first
measurements were carried out only in dry bone and not in fluid-saturated bone. Therefore, it
14
is now clear that piezoelectricity is not the source of voltages generated when bone is
mechanically stressed. What is the source of those voltages?
Bound and unbound electric charges exist in bone tissue (55). Wall channels and the
extracellular matrix in bone tend to be negatively charged due to the negative charge in
collagen and the negative fixed charges on carbohydrates and proteins respectively (62).
Thus, a fluid electrolyte bound by the extracellular matrix and wall channels will have a
diffuse double layer with an excess of positive ions (figure 6). When bone is loaded, the
resulting compressive strain causes bone fluid to flow across all canals from regions of high
pressure to lower pressure between two different sites in bone tissue. The bone fluid with its
excess of positive charge is convected, thereby developing a convective current that is called
the streaming current (15, 18). The potential difference produced by this convective current
between two different bone sites is called the streaming potential (16, 17). Thus, when
electrodes are placed on two different bone sites and then bone is deformed, the voltage
produced by the streaming current is measured. These potential differences or voltages
generated when bone is deformed by the application of a mechanical load are called stress- or
strain-generated potentials, SGPs, and are simply the result of the streaming currents and
potentials (56). Since SGPs dominate over those of piezoelectric origin in fluid-saturated bone
(13, 14), it means that those voltages measured in fluid-satured bone when loaded are due to
streaming potentials rather than piezoelectric polarization. SGPs are now recognized as
dominantly electrokinetic phenomena explained by an extension of poroelasticity (15).
Poroelasticity is a theory that models the interaction of deformation and fluid flow in a fluidsatured porous medium, as the found in bone tissue (15). The deformation of the medium
influences the flow of the fluid and vice versa. The electric potentials generated are directly
related to the pressure gradients that produce the fluid flow, thus, the potential difference
measured is proportional to a pore pressure difference between the same two points used to
measure the potential difference (18). Until relatively recently, it was widely believed that
SGPs were due to the small pores in the mineralized matrix (63). However, research in bone
poroelasticity predicted recently that the size of the pores required to satisfy the measured
magnitude and phase of the SGPs is approximately between 15-30 nm of radius (14, 15, 64).
It therefore excludes the mineralized matrix, as the sizes of the pores that were hypothesized
to exist are of maximum 10 nm. In support this theory, experiments with tracer molecules
have shown that tracers never penetrate the mineralized matrix and are confined to the
lacunar-canalicular porosity (14). Therefore the anatomical site in bone that contains the fluid
source responsible of SGPs seems to be the lacunae-canaliculi system, exactly where
osteocytes with their respective processes are located (figure 2-4).
Figure 6. An electrical current and potential difference are generated when the electrical charge in a
bone canal is convected. Wall channels in bone tend to be negatively charged due to collagen (62). Therefore
bone fluid will have a double layer of positive charges bound to walls as shown in figure. When bone is loaded,
the resulting compressive strain causes bone fluid to flow across all bone canals from regions of high pressure to
lower pressure between two different sites in the bone tissue.
The duration of the voltage generated in bone when mechanically loaded is determined
primarily by the rate of compression and the mechanical relaxation properties of the bone
15
(15). SGPs have been measured across individual osteons as a function of frequency across
bone samples in a four-point bending apparatus and in vivo, as shown in figure 7 (17, 65).
One microelectrode is moved across a surface of exposed bone while viewing through a
microscope, and other one is placed in a reference point, normally in the harvesian channel
(mittle point in an osteon). The value of the potential rises steeply from the harvesian canal to
a peak at the cement line (end side of a osteon, figure 2 and 3), and then decreases to the
harvesian canal of the next osteon. The potential difference profile is thus a cusp. It has been
found that voltages generated surrounding the harvesian canal are 10-30 times larger than
across the entire bone specimen. The typical values of voltage densities measured in
individual osteons are between 20-50 mV/mm. This observation also suggests that the
dominant fluid flow occurs into or out of the harvesian canal (15). Therefore, the bone fluid
flows from the harvesian canal towards the lacunae-canaliculi when the bone is mechanically
stressed, giving rise thus to the potential difference. The question is if these voltages play a
possible role in transducing the mechanical stimulus into a biological response during
adaptative remodeling and maintenance of bone. It has not been proved yet.
A
B
Figure 7. Measurement of stress-generated potentials in vivo (A) and in vitro (B) in bone. In picture A
the voltage is measured with periosteal electrodes when compressive loading (arrows) is applied. Picture B
shows a four bending system to apply mechanical loading to bone in vitro (adapted from ref 65).
When bone is not stressed (intact bone), the difference in ion concentration between its
layers produces a potential difference across or along bone surface. This voltage is usually
measured with a pair of electrodes, where one electrode is usually held at a single position
and used as reference, while the other electrode is displaced across or along bone surface. The
voltage measured is known as bioelectric or steady-state potential and was originally
described by Friedenberg and Brighton (20). Thinking on images 2-4, it is clear that the
electrochemical gradient will not be the same at areas where bone undergoes growth,
resorption or exchange of nutrients. Thus, an intact bone usually display a potential difference
between two different points. Bioelectric voltages produce electrical current densities within
the range of 0.5 to 12 µA/cm2 (22, 62). Interestingly, it was found that areas of active growth
and repair are electrically negative when measured and compared with less active areas in
bone, or with respect to a reference point (21-23, 66). Therefore bioelectric potentials have
been implicated to play a role during growth (66). Furthermore, when bone is injured or
fractured, the entire bone becomes electrically negative, and a peak value of voltage is
measured exactly at the fracture site (22-24). The electrical currents produced by the voltages
due to the lesion are now called injury or fracture currents, and their densities increase to
approximately 100-130 µA/cm2 immediately after the lesion (23, 62). However, after one
hour of lesion, current densities decline about 10% of the initial peak value and weaken
slowly (62). Injury or fracture currents are thought to play a role during healing (23).
Therefore based on these findings, it was postulated that the endogenous bioelectricity of
bone might play a role during bone growth, repair and healing (22-24).
16
In conclusion, two distinct types of endogenous electrical activity in bone have been
observed and measured. The first type described in this work were the voltages and electrical
currents produced when bone is mechanically loaded, giving rise the so called stress
generated potentials. The magnitude and duration of the voltage and currents generated are
determined primarily by the rate of compression and the mechanical relaxation properties of
the bone. SGPs are thought to play a role during bone remodeling and mechanotransduction
in bone. The second type of endogenous electrical activity in bone are the voltages and
currents measured in intact and injured bone, which are generated by the difference in ion
concentration between the different bone layers. The voltages are relatively steady state or
slowly changing, and are measured when a pair of electrodes are inserted between two
different bone sites. These voltages, usually termed DC biopotentials are thought to play a
role in bone healing and growth.
1.202. Exogenous Electrical Activity in Bone
After Yasuda discovered the so called “piezoelectric effect” in bone, convinced of the
role of these potentials in bone remodeling, he predicted that an externally applied electrical
stimulus would stimulate bone formation. He implanted a pair of direct current electrodes in a
rabbit femur. Over three weeks, a ridge of bone (callus) formed between the electrodes sites,
more specific in the vicinity of cathode. He took this as confirmation that electricity takes a
role in bone formation (67). Almost at the same time, in the early1960s, Basset also observed
bone formation with implanted electrodes (25). Based on these findings, and as bone
generates its own currents and potentials when broken, many laboratories around the world
began studying the effects of electricity on bone (68-70). Research showed that implanting
weak electrical current directly into the bone increases bone formation around the cathode and
decreases bone around the anode. The optimal bone growth took place with currents between
5 and 20 µA. Levels below 5 µA did not enhance growth, while above 20 µA caused cell
necrosis and bone death. The results seemed consistent with the values of fracture current
measured and with the electronegativity at the fracture and growth site. It also suggested that
osteoblasts were activated by negative charges and perhaps osteoclasts by positive charges
(62). From all this research, bone growth stimulators were developed as a rehabilitative tool
for complete bone repair (26-30). Bone growth stimulation is the technique of promoting bone
growth in difficult to heal fractures by applying a low electrical current or other electrical
stimulation technique. By the early 1980s, electrical stimulation was firmly established as a
treatment modality for the management of nonunions (26, 62). To date, electrical bone growth
stimulators are a supplemental form of therapy to help enhance the body’s bone healing
process. Initially, the method of electrical stimulation was a direct contact with the zones of
interest. Actually there are three different techniques of bone growth stimulators: Invasive,
semi-invasive and non-invasive (62). An invasive technique requires that cathode, anode and
battery supply are surgically implanted into the fracture site. In a semi-invasive technique the
cathode is inserted into the fracture site, while anode and battery are placed on the skin. A
noninvasive technique, also known as capacitive coupling, requires that anode and cathode
are placed on either side of the fracture site on the skin. This latter technique has shown to be
more suitable, its advantages are: no surgery, no risk of infection, most conservative and least
expensive. Therefore it is the most commonly used to assist fractured bone repairing and
healing.
Since electrical stimulation has demonstrated consistently high success rates even in
complicated bone healing situations (26, 29, 31), the interest in methods of accelering bone
formation with electricity has increased enormously. Initially only DC electrical currents were
used in bone growth stimulators. Actually different forms of electrical stimulation are used:
17
capacitive and inductive coupling, electric and magnetic fields, DC, AC and pulsating among
others (31-33). Furthermore, recently stimulators through ultrasound have increased their use
as bone formation stimulators (34, 35). All these techniques of stimulation have been proven
to work and their uses depend on pathology and decision of doctor and patient (36-39, 71). In
conclusion, the effect of electricity on bone cell physiology leaves no doubt that electricity in
its various forms can alter many cellular events. However, despite an abundance of
undeniably valid data, from both basic and multicenter clinical trials, demonstrating the
effects of electrical stimulation on skeletal biology and healing, little is known about the
signal transduction that mediates the physiological response of bone to electrical stimulation.
As consequence, a certain amount of skepticism remains among biologists and clinicians, due
to the fact sometimes the method used is not the appropriate one and can thus bring collateral
damages. Therefore scientists need to learn more about how electricity interacts with bone,
thereby it will provide more effective electrical stimulators and prevent principally secondary
damages.
Until recently, it was only believed that such alterations in the electrical environment
of bone are perceived by bone cells which in turn initiate a chain of intracellular events that
lead to an adaptive bone formation. However it has not yet been proved. To try to solve such
dilemmas a multitude of in vitro studies with bone cells and electrical stimulation have been
performed over the last 20 years. As electrical stimulation has been used electric and magnetic
fields, direct current and ultrasound among others (5-8, 40-43). Bone cells have responded
with proliferation (8), differentiation (72), initialization of mineral formation (73), calcium
increase (40), and migration (5) among others, all these cellular responses are considered key
in the process of bone formation. Therefore it has not been questionable if bone cells respond
to electrical stimulation, but what is the mechanism that is acting on the cells, and how it is
transduced into biochemical signals for generating an appropriate physiological response.
Further research is required to characterize the process of electrical transduction of bone cells
subjected to electrical stimulation. A first step should be to understand how a single bone cell
might react to electricity, specifically with electric fields. Attempting to answer this question
was the scope of this thesis. To try to understand how a bone cell could react to an electric
field, the next section focuses on explaining the electrical properties of single bone cells and
the cellular effects reported until recently by the application of electric fields to bone cells.
1.3. ELECTRICAL PROPERTIES OF BONE CELLS
All living matter consists of cells that have a similar structure; bone is not an
exception. Bone cells as any other living cell consist of a cytoplasm surrounded by a
membrane. Membrane and cytoplasm are composed of many components. Details about cell
architecture are out of the scope of this thesis. Only components that have been reported
previously to play a role in the electrical properties of the cells are mentioned here.
Cells can be modeled as a uniform, ellipsoidal shells enclosing the cytoplasm and
surrounded by the extracellular medium (figure 8). The difference in ion concentrations
between the extracellular and intracellular medium gives rise to a potential difference known
as the transmembrane potential or simply membrane potential (74). All living cells have a
membrane potential with the inside of the cell being negative relative to its external surface.
The membrane potential is typically in the range of –20 to –200 mV according to the type of
cell; bone cells have normally a membrane potential of –70 mV (9). The cell membrane
potential is strongly linked to the transport mechanisms, membrane architecture and signaling
mechanisms of the cells; thus a change in the membrane potential will profoundly influence
changes in the behavior of the cells.
18
Figure 8. Cells are modeled as spherical shells enclosing the cytoplasm. All cells have a membrane
potential caused by the ion concentration differences between the extracellular and intracellular medium.
The cell membrane conductivity is several orders of magnitude lower than those of the
cytoplasm and the physiological extracellular medium. Since high resistivity of the plasma
membrane to current flow, electric fields are unlikely to affect significantly the cytoplasm of
cells (75, 76). In contrast, electric fields are known to have distinct physical effects on the cell
membrane, mainly because most of the electric field is concentrated on it. Therefore all
research with electric fields has concentrated around the membrane. A description of the
membrane composition, its electrical properties, and how it contributes to the generation of
the transmembrane potential are mentioned in the next section. Furthermore, the different
effects that have been reported for applying an electric field to the cells are also mentioned
here.
1.301. The Membrane
Biological membranes are extremely thin complex mixtures of lipids and proteins (410 nm in thickness). Biological membranes have been characterized as a fluid mosaic model
(figure 9, ref. 77). The lipid bilayer is the main fabric of the membrane; it is a sheet like
structure composed of two layers of phospholipid molecules. Each phospholipid molecule has
a polar (hydrophilic) head and two nonpolar (hydrophobic) tails. These phospholipids are
aligned tail to tail so the nonpolar areas form a hydrophobic region between the hydrophilic
heads on the inner and outer surfaces of the membrane. The major driving force for the
formation of phospholipid bilayers is hydrophobic interaction between the fatty acyl chains of
glycolipid and phospholipid molecules. Van der Waals interactions among the hydrocarbon
chains favor close packing of these hydrophobic tails. Hydrogen bonding and electrostatic
interactions between the polar head groups and water molecules also stabilize the bilayer. The
hydrophobic core impedes the transport of hydrophilic structures, such as ions and polar
molecules but enable hydrophobic molecules, which can dissolve in the membrane, cross it
with ease. Thus membranes are semi-permeable structures.
Proteins are embedded at irregular intervals in the membrane and determine most of
the membrane's specific functions (77). Proteins are held by hydrophobic interactions between
the membrane lipids and hydrophobic domains in the proteins. Membrane proteins are
classified into two major categories, Integral proteins and Peripheral proteins. Integral
proteins are generally transmembrane proteins, with hydrophobic regions that completely
span the hydrophobic interior of the membrane. The hydrophilic ends of the molecule are
exposed to the aqueous solutions on either side of the membrane. Peripheral proteins are not
embedded in the lipid bilayer at all; they are loosely bound to the surface of the membrane,
often to the exposed parts of integral proteins.
The plasma membrane also has carbohydrates, which are restricted to the exterior
surface(77). Membrane carbohydrates are usually branched oligosaccharides with fewer than
19
15 sugar units. Some of these oligosaccharides are covalently bonded to lipids, forming
molecules called glycolipids. Most are covalently bonded to proteins, which are thereby
glycoproteins (77). The oligosaccharides on the external side of the plasma membrane vary
from species to species, among individuals of the same species, and even from one cell type
to another in a single individual. The diversity of the molecules and their location on the cell's
surface enable oligosaccharides to function as markers that distinguish one cell from another.
Membranes are also asymmetric; they have distinct inside and outside faces. The two
lipid layers may differ in specific lipid composition, and each protein has directional
orientation in the membrane. Furthermore membranes are not static sheets of molecules
locked rigidly in place. Most lipids are randomly mobile in the plane of the membrane with an
average migration rate of 22 µm per second. Proteins are much larger than lipids and move
more slowly, but some do drift. Some membrane proteins seem to move in a highly directed
manner, however, many others seem to be held virtually immobile by their attachment to the
cytoskeleton.
Figure 9. Biological membranes are modeled as a fluid mosaic. Biological membranes consist of a lipid
bilayer, diverse carbohydrates and proteins (Internet Classnotes for Biologists, Texas A&M UniversityKingsville, Department of Biology. http://ntri.tamuk.edu/cell/membranes.html)
1.302. Electric Fields at the Membrane Level
The difference in ion concentrations between the extracellular and intracellular
medium gives rise to a potential difference known as the transmembrane potential (74). Since
electrical resistance of the membrane is high, it can be regarded as an insulator separating two
conducting aqueous phases. It automatically generates an electric field with electrical charges
distributed on both membrane surfaces. Many models have been proposed to explain how
electric fields should be integrated at the membrane level (78-82). However, none of those
models have been demonstrated experimentally. The following is a summary of some of the
common models.
The phospholipid bilayer, besides providing a hydrophobic barrier separating the intrafrom the extracellular aqueous compartment, has a negative charge on both sides, which is
mainly due to the negatively charged phospholipids (roughly 10% of membrane lipids are
negatively charged) (78, 79). Positively charged lipids are rare. Most commonly occurring
lipids with net charges are fatty acids, phosphatidic acids, phosphatidylserines,
phosphatidylethanolamines and cardiolipins (80). Other constituents of membranes, such as
proteins and carbohydrates, also have net charges at neutral pH. These charges do not always
(even normally) cancel one another. Further superficial negative charges are provided by a
20
variety of molecules inserted in, or attached to, the membrane (proteins, glycolipids,
proteoglycans), which contribute to bring the charge density ρM up to about -10 µC/cm2 (81,
82). The simplest conceivable electrostatic model for a bilayer is a thin slab of low dielectric
material (i.e. a hydrocarbon, with a relative permittivity or dielectric constant (εr) ranging
from typically εr ≈ 2-10) immersed in a continuum of high dielectric material (i.e. water εr ≈
80). The dielectric constant is the ratio of the permittivity of a substance to the permittivity of
free space (εr = ε / εo ). The dielectric constant is dimensionless, and can be defined as an
expression of the extent to which a material concentrates electrostatic lines of flux. In other
words, the electric charge is hold in the material preventing thus its flux. As the dielectric
constant increases, the electric flux density increases. Therefore the low dielectric interior in
the membrane presents a high (roughly 40 kcal/mol) energy barrier to the passage of ions
across it (83). To avoid such cost of energy, membrane-spanning proteins containing ionselective pores, the so-called ion channels, facilitate selective ion transport across membranes,
which is critical for many biochemical processes.
In this electrostatic model, on each side of the membrane there is a positive ionic cloud
facing the negative charge layer and providing the system with electroneutrality. Physically,
this makes the membrane behave as a series of plane plate capacitors, with the electrostatic
profile outlined in figure 10.
Figure 10. The electrostatic profile across the plasma membrane and at interfaces (model proposed by
Olivotto, ref. 79). At each interface of the membrane with the internal (left) and external (right) electrolytic
milieux, the fixed negative charges (here represented on the polar heads of phospholipids) can be thought of as
constituting a layer that attracts a cationic “diffuse layer” (the Gouy-Chapman layer, GC) in which cations are
more concentrated than in the bulk aqueous phase. Interleaved with these layers there is a “compact layer” (CL)
of water molecules (circles containing arrows) in contact with the membrane and strongly polarized by the
membrane surface charge. The surface potential (φi or φe to intra- extracellular respectively) is the sum of the
difference potentials in ∆χ and φGC. VREST is the potential difference between the extracellular and intracellular
medium, measured at distances larger than the Debye distance (76, 80, 84) from the corresponding side of the
membrane.
21
1.302a. The Potential Difference Across The Compact Layer (∆χ)
Whatever the model adopted to describe the electrostatic profile of the plasma
membrane, it is always necessary to make a distinction between the first monolayer of water
molecules in contact with the surface and the water of the bulk solution bathing the surface
itself (figure 10). This layer usually referred as the Compact Layer (CL), has a peculiar
structural and electrostatic properties. In fact, the presence of the surface breaks the spatial
symmetry of the liquid phase and induces a structural arrangement of the molecules lying
close to the surface (85). The details of this arrangement depend on the nature and the charge
of the surface. In the case of the plasma membrane, the charge density of the order of –10
µC/cm2 generates a strong electric field, which orientates the water dipoles, producing a
potential difference (∆χ) across the CL, between the surface and the bulk solution bathing the
surface (see figure 10). When in this layer, water molecules are replaced by absorbed organic
solute molecules endowed with their own polarizability (different from that of water), the
change in the induced dipole moment generates a variation in ∆χ. Hence absorption of
substrates to the internal or external compact layer can produce variations of the potential
across the corresponding compact layer, and therefore variations in the corresponding
potential difference ∆χi (internal) or ∆χe (external) respectively.
1.302b. The Diffuse Layer Potential (φGC)
At each interface of the membrane with the internal and external electrolytic milieux,
the fixed negative charges can be regarded as constituting a layer that attracts an ionic
“diffuse layer” the so called Gouy-Chapman cloud, where cations are much more
concentrated than in the bulk solution (86). The ensuing “double layer”, which is known as
Gouy-chapman layer, GC, together with the interposed water (circles containing arrows in
figure 10), forms a capacitor, where the membrane is the dielectric. The potential difference
between the compact layer and the bulk solution on a same side of the membrane (internal or
external) is known as the diffuse layer potential or Gouy-Chapman potential, φGC (79).
1.302c. The Surface Potential (φ)
The potential difference between the membrane surface and the bulk of the electrolytic
medium (on a same side of the membrane) is the sum of two contributors: the electrical
potential difference across CL (∆χ) and the electrical potential difference across the GouyChapman Layer (φGC). The sum of ∆χ + φGC constitutes an overall electrostatic parameter
usually indicated as “surface potential”, φi and φe at the intra- or extracellular surface of the
membrane respectively.
1.302d. Resting Potential (VREST)
The bulk to bulk transmembrane potential is the electric potential difference between
the extra and intracellular bulk aqueous phases, which completely drops in the region of the
plasma membrane. The potential difference arises from protein-mediated processes, which
generate a movement of charged particles across the membrane so that a net separation of
charge takes place across the membrane. The movement of charged particles is carried out by
electro-diffusion through ion channels immersed in the membrane, or by active transport
operated by electrogenic ion pumps (e.g. the Na+,K+ ATPase) (87, 88). Under equilibrium
conditions, when the cell is at rest (when 0 current is passing across the membrane or when
the sum of all ion currents across the membrane is zero), the voltage difference between bulk
to bulk is called the resting potential (VREST), with the internal side being negative in relation
22
to the external side of the membrane. This potential difference across the plasma membrane of
animal cells varies between 20 and 200 mV (87, 88), and it corresponds to a given amount of
charge separation across the membrane. Resting potential is the most familiar potential
difference to cell biologists, typically described in neurons, muscle, and bone cells, and is the
only one measurable by classical electrophysiological techniques. Resting potential is also
known as transmembrane or membrane potential.
To measure the potential difference across the cell membrane and to establish the sign
of VREST, the positive lead of the voltmeter is always placed inside the cell. This is usually
accomplished by inserting the positive lead (typically a Ag/AgCl wire) of the voltmeter into a
micropipette filled with a conducting solution, such as 1.5 M KCl. The tip of the pipette has a
very small opening (typically less than 0.1 µm) that provides electrical continuity with any
solution it contacts. The micropipette is attached to a micromanipulator that allows accurate
control of its position. The micropipette is pushed through the cell membrane so that its tip
comes into contact with the intracellular medium. The tip of the micropipette is so small that
it does not damage the plasma membrane; the membrane tends to adhere to the outside of the
micropipette and seal the hole. The negative lead of the voltmeter is placed in contact with the
solution on the outside of the cell. This is usually accomplished by inserting the negative lead
into a second micropipette (also filled with 1.5 M KCl), the tip of which is in contact with the
extracellular medium.
1.302e. An Endogenous Electric Field, Emem
Since high resistivity of the plasma membrane to current flow, an electric field is
created as a result of separating charge across the membrane. Thus, all cells are exposed to an
electric field created by their transmembrane potential between the two sides of the cell
membrane. The transmembrane potential, typically in the range of –20 to –200 mV, imposes
an electric field strength of roughly –40 to –200 KV/cm on the cell membrane if the thickness
of the membrane (d) is assumed to be 5-10 nm (Emem = VREST / d). The transmembrane
potential is the potential difference measured between the two bulk aqueous phases (VREST).
1.302f. The Electric Field Undergone by Proteins at the Membrane Et
If one takes a macromolecule (an integral protein) spanning the membrane, the true
electric field (Et) affecting this molecule across two points located at the opposite sides of the
membrane will be given by the sum of the resting potential plus the difference between the
intra- and extracellular surface potentials. Hence,
Et = VREST + (∆χi - ∆χe) + (φGCi - φGCe)
(2)
This equation shows that any component of Et contributes to the functional modulation
of voltage-dependent proteins that are placed in the membrane.
1.303. Polarization and Dispersion of the Cell Membrane
The membrane, because of its large lipid content, is relatively impermeable to ions.
Phospholipids provide the capacitive property of the membrane (Cm), which allows charge,
once separated, to remain separated across the membrane. Thus cell membrane can be
regarded as an insulator separating two conducting aqueous phases, i.e. as a dielectric in a
parallel plate capacitor. Therefore, in an equivalent electrical circuit the cell membrane can be
represented as a capacitance and a resistance arranged in parallel. All biological membranes
23
seem to have the same specific capacitance (i.e. capacitance per unit area), which assumes a
value of 1 µF/cm2 (76, 89). Higher values for the specific capacitance have been described in
very few rare exceptions (90). Like any material, a membrane will be polarized in the
presence of an electric field across it. Polarization most often occurs due to formation of
dipoles or orientation of permanent dipoles within the membrane or at interfaces between the
membrane and the electrolyte solution. Orientated polarization occurs in a dielectric
containing polar molecules with a permanent dipole as in the case of a biological membrane.
These dipoles normally may be randomly oriented, but align themselves in direction to the
field lines in the presence of an electric field as an extent depending on field strength and
thermal agitation. At high field strengths dielectric saturation can occur i.e. orientation of all
permanent molecules. Insofar as polarization involves the movement of matter, it does not
occur instantly after a change in the electric field, therefore, all dielectrics show loss (76, 89,
91). Under a more rapidly changing voltage, such as an AC field of high frequency, fewer and
fewer dipoles will be formed or reoriented in time, and, in consequence, dielectric dispersion
will occur. The dispersion is characterized by the dielectric relaxation time, given by τ =
1/2Πƒ where ƒ is the frequency of the AC field at the dispersion. In the frequency range
where the dispersion occurs, the dielectric absorbs energy from the AC field. The dispersion
frequency depends on the dielectric and the mobility of the dipoles in it.
Cell membranes show more than one dispersion. The dependence of the dielectric
constant and the conductivity of a living system on the frequency exhibit four remarkable
dispersions: these are known as α, β, δ and γ dispersions (92) (figure 11). Thus with
increasing frequency the dielectric constant and the specific resistance very often decrease.
The α dispersion is carried out in low frequencies (1-104 Hz), and is caused by the tangential
flow of ions near membrane surfaces. The β dispersion is centred in the range of 104-108 Hz,
and results from the build-up of charges at cell membrane due to the Maxwell-Wagner effect.
The phenomenon known as the Maxwell-Wagner interfacial arises from the free charges that
accumulate at the two cell membrane surfaces through intracellular and extracellular medium,
and that induced the dipole, i.e. at the inner and outer boundaries of the plasma membrane
(92). Above the β dispersion, the cell membranes have negligible impedance and the current
passes through. The δ dispersion is centred in the range of 108-109 Hz, and is produced by the
rotation of macromolecular side-chains and bound water. Finally, the γ dispersion is centred
in the rage of 1010 Hz, and is due to the dipolar rotation of small molecules, particularly water.
This dispersion is the same as the found in pure liquid water.
1.304. Deformation of the Cell Membrane
The polarization of the cell membrane produces an electric force that acts on the
membrane, and as a result the cell membrane is deformed, elongated or compressed at the
direction of the field (93).
The deformation force exerted on the membrane, called the dielectrophoretic (DEP)
force (92), is caused by the electric dipole induced within the cell and a nonuniform electric
field. The induced dipole arises from the free charges that accumulate at the two cell surfaces,
i.e. at the inner and outer boundaries of the plasma membrane. The resulting deformation
force is caused by the Maxwell stresses applied to the cell interfaces. The polarity of this
force depends on the orientation of the induced dipole (i.e. of the interfacial charges), which,
in turn, is determined by the effective polarizability of the cells relative to the suspending
media, thereby resulting in compression or elongation of the cell membrane. Thus, both the
generated dipole and the deformation force are conductivity-dependent (92). However due to
24
the layered structure of the cells, the magnitude and polarity of the force can occur in a few
nanoseconds or vary with time in a complicated manner (92).
Figure 11. Idealized spectrum of the dielectric properties of cell suspensions and tissues. The step
changes in permittivity are called dispersions and are due to the loss of particular polarization processes as
frequency increases. The α dispersion is due to the tangential flow of ions across cell surfaces. The β dispersion
results from the build-up of charge at the cell membrane due to the Maxwell-Wagner effect. The δ dispersion is
produced by the rotation of macromolecular side-chains and bound water, and the γ dispersion is due to dipolar
rotation of small molecules, particularly water. (Graphic taken from reference 92).
1.305. Cytoplasm and the Extracellular Medium
The cytoplasm of cells is highly complicated, it contains not only large amounts of
salts, proteins, nuclei acids and smaller molecules, but in many cases a large number of
organelles and various membranous structures (nucleus, vacuoles, etc), which can also affect
the dielectric properties. The values of relative permittivity or dielectric constant of cytoplasm
of cells has been measured ranging from about 50 to 200 (76, 94-96). In most cases, however,
the cytoplasm can be approximated as a highly conducting salt solution with a large
concentration of dissolved organic material. The extracellular medium is always also regarded
as a highly conducting salt solution (94).
An electric field cannot be sustained in a medium with high electrical conductivity, as
those composed by the cytoplasm and the extracellular medium (Debye theory) (76, 80, 84).
When an electric field is applied to a conducting medium, as the cytoplasmic or extracellular
medium that exists in biological cells, it polarizes, and the new charge distribution will reduce
the magnitude of the electric field inside the medium. The free charges according to their
polarity move towards the perturbing charge (the positive or negative pole of the electric
field), thereby reducing (shielding) the electric field that every pole makes. At a sufficient
distance, a neutralization of the electric field is carried out. This distance, known as Debye
length, is thus a measure of the distance over which an individual charged particle can exert
an effect (84). Beyond the Debye length the individual charged particle appears as if it is not
there because it has been neutralized. To a better understanding of this effect, consider the
same medium with mobile positive and negative charged particles; if a positive test charge is
introduced in this medium, it will attract negative charges and repel positive charges, making
a cloud of net negative charge around itself. The net effect is to ``screen'' the test charge,
thereby the electric field of this positive test charge will be partially shielded, and over a
sufficientely large distance, this shielding (known as Debye shielding) becomes complete.
The distance beyond which the test charge is effectively screened is called Debye length, and
it is considered a measure of the effective shielding length (76, 84). Therefore, the Debye
25
length is a measure for the electrostatic screening in a conducting medium. Thus charged
particles placed beyond the Debye length do not feel the electrostatic effect. Therefore,
electric fields applied to extracellular medium or cytoplasm of cells are subject to a strong
attenuation and decrease rapidly over a distance of Debye length (76, 84).
In summary, since the high ionic strength and electrical conductivity of physiological
medium, an electric field cannot be sustained at distances greater than Debye length from the
originating charge distribution. However, since high electrical resistance of the membrane,
membranes of cells are the privileged sites for electric fields. Therefore, the whole electric
field will concentrate only at the membrane and will extinguish into the cytoplasm or
extracellular medium at distances greater than Debye length. In most cells the Debye length is
typically 1 nm (76, 80). Thus at distances larger than 1 nm from cell membranes, the electric
fields are practically extinguished.
1.4. EFFECTS OF MODIFYING THE NATURAL ELECTRIC FIELD
The application of electric fields to biological cells has resulted in a diversity of effects
in the cells. For example, application of a large DC or low-frequency AC electric field to a
cell induces a large potential drop across the plasma membrane that can cause dielectric
breakdown. This rupturing of the membrane has been used to kill cells, and has also found
applications in electro-diffusion for the creation of new hybrids and electro-poration for the
introduction of new genetic material into cells (90, 97). On other hand, it has been
demonstrated theorically and experimentally that cells experience an asymmetric disturbance
of transmembrane potential toward the poles of an applied electric field. Cathode-facing
membrane surfaces are depolarized while anode-facing surfaces experience a
hyperpolarization (76, 98, 99). Additionally, electric fields act on plasma membrane surfaces
parallel to the field, causing the electrophoretic (100, 101) and/or the electro-osmotic (102,
103) lateral redistribution of several classes of plasma membrane glycoproteins.
1.401. Distribution of Membrane Potential
Application of an uniform electric field, DC ELF, in a conductive medium to cells of
any size and of geometry that can be modeled as a uniform, ellipsoidal shell enclosing
conductive cytoplasm, should induce a calculable spatial alteration of the transmembrane
potential (76). The direct target of the steady field action on a single, relatively iso-diametric
cell should be the plasma membrane since practically all of the resistance across such cells is
generally across this membrane. Consider such uniform spherical cell placed in a conductive
medium (figure 12).
Figure 12. Field lines around a nonconducting sphere placed in a DC ELF (Cole, 76). Tangential field at
the cell surface is related to the uniform field applied, Eappl.
Assuming the membrane is nonconductive, or its resistivity is much higher than those
of the internal and external media (76), the transmembrane potential generated by an applied
26
DC electric field may be calculated simply by the well-known relation, often referred to as the
steady-state Schwan’s equation (91).
Vind = 1.5 EappRcosθ
(3)
Where Vind is the induced transmembrane voltage, Eapp is the external DC electric field
applied, R is the cell radius and θ is the polar angle measured from the center of the cell with
respect to a point of interest on the cell membrane.
If the membrane conductivity is not negligible, the transmembrane potential changes
given by Eq. 3 are, to first order, in the quasisteady state, reduced by the correction factor Βo
(104).
Vind = 1.5BoEappRcosθ
(4)
Where Bo = 2σeσi / [(2σe+σm)(2σm+σi) + (R/d) σm (2σe+σi)]
σe, σi and σm are the electrical conductivities of extracellular, intracellular and membrane
respectively, and d is the membrane thickness. The ratio σm/d is the effective membrane
permeability (104).
Note that if σm= 0 (ideal nonconductive), or << σe, σi, Bo = 1 and Eq 3 stays without
change. The only condition is that the membrane conductivity has to be much lower than the
conductivity of extracellular and intracellular media. With physiological values of the
conductivities, the factor 1.5 differs at most by several parts per thousand from the exact
result (104). Thus that Bo =1 for most healthy cells (100, 104, 105).
If θ = 0o or 180o, cosθ = 1. Thus, the maximal transmembrane voltage change that
occurs at both sides is
Vind = 1.5 EapplR
(5)
For an usual DC ELF (1-10 V/cm) applied to a cell of 10 µm of radius, the potential of
the membrane facing the anode will be hyperpolarized by 1.5 - 15 mV and the facing the
cathode depolarized by the same amount.
The total transmembrane potential measured in a cell after application of a DC ELF
will be given by
Vtm = VREST + Vind
(6)
Where VREST is the transmembrane potential before the imposition of the electric field
(resting potential).
The effect of an applied DC electric field on transmembrane potential is not negligible,
and the possible involvement of membrane potential dependent mechanisms in altering the
topography and mobility of membrane components should not be overlooked (106, 107).
In case that the applied electric field is alternating, AC; the situation becomes more
complex and the induced transmembrane voltage becomes strongly dependent on the
frequency of the applied electric field (108. 109). With very high frequencies, the high
27
electrical resistance of the membrane will be decreased, and therefore the membrane will not
be able to sustain the induced electric field, resulting thereby in the pass of current through.
The critic frequency until where the induced transmembrane voltage will be independent of
frequency, is that comparable with the reciprocal of the time constant, T (109). The
transmembrane voltage induced by the application of an alternating electric field was
theorically derived by Schwan (108).
Vind = (1.5EapplRcosθ) / (1+ ωΤ)
(7)
Where:
Τ = RCm(ρi + ρe / 2)
R = Radius of the cell
ω =2πƒ
ƒ = frequency of the applied electric field
Cm= Capacitance of the membrane measured in terms of the area of the membrane (F/m²).
Cm = εm / d. εm is the permittivity of the membrane (F/m), and d is the thickness of the
membrane (m).
ρI, ρe = resistivities of the internal fluid in cytoplasm and external medium respectively.
Resistivities are given in ohms meter.
At lower frequencies than 1/Τ, the induced electric field is independent of the AC
frequency, and at higher frequencies than 1/Τ, membrane polarization is reduced and
electrical current begins to flow into membrane (109).
1.402. Electrophoresis in the Plasma Membrane
The most convincing evidence that the cell membrane is basically fluid in structure
comes from studies that showed that many macromolecular components undergo long-range
movement in the plane of the cell membrane (110). DC electric fields cause the lateral
redistribution of several types of plasma membrane glycoproteins, including receptors for
plant lectins (111), low-density lipoprotein (112) and Epidermal Growth Factors (113-115)
amongst others. One of the most studied class of electro-mobile glycoproteins are the
receptors for the lectin Con A (116), which undergo electrophoretic or electro-osmotic
redistribution in a wide variety of cells. It has been shown that concanavalin A alters the
morphology and locomotory behavior of several cell types. Con A causes the rearrangement
of cytoskeletal components in cells as fibroblasts (117), inhibits the migration of corneal
epithelial cells (118), neural crest cells (119) and macrophages (100). While it is clear that
Con A receptors may be involved in the control of cellular architecture and migration, it has
yet to be determined whether or not electric field-induced asymmetries of plasma membrane
glycoproteins can cause structural polarization and directional migration of the cells.
The term electrophoresis (-phoresis = Greek, phoros - act of carrying) refers to the
migration of a charged particle or molecule in an electric field. The theoretical analysis of in
situ electrophoresis of membrane-bound charged molecules has been carried out using a
spherical cell as a simple model. The molecules are envisioned to possess a hydrophobic
portion embedded in the membrane and a charged portion exposed to the extracellular
aqueous phase. The cell is immobilized by its adhesion to the surface of a substratum, and it is
assumed that adhesion does not distort the spherical shape of the cell. The interaction between
the adjacent molecules is ignored (100). The factors that can effect the migration of a charged
molecule include the size of the charge, the electrical potential, and the frictional resistance of
28
the particle. The driving force on a particular molecule, F1, is the product of the charge, q, and
the electric field, E. This force will displace the molecule at a distance from its initial
position. However as the molecule moves, it also receives a damping force, F2, which is
dependent of the molecule’s velocity, v, and the attenuation coefficient for the molecule’s
movement, κ. The molecule continues its movement whenever the driving force is greater
than damping force. At equilibrium, F1 = F2, thus
Eq=vκ
(8)
v = (E q) / κ
(9)
It is clear that the molecule’s velocity depends upon the applied electric field and the
attenuation coefficient for the molecule’s movement, which depends upon the internal friction
coefficient (viscosity), η, of the medium in which the molecule moves (that is the cytoplasm
or extracellular medium) and the radius, R, of the molecule (96). If the molecule or particle is
spherical (Stoke's law), then
κ = 6ΠηR
10 in 9
(10)
Units of viscosity are given in Kg/m • s. Therefore units of κ are Kg/s. Replacing Eq.
v = (E q) / (6ΠηR)
v / E = q / (6ΠηR)
(11)
The expression v / E is the electrophoretic mobility (U), and is defined as the ratio of
the velocity of particles to the field strength in a solution with determined viscosity (100,
101). Thus
U = q / (6ΠηR)
(12)
Therefore, the mobility of the charged particle or molecule depends on its charge, its
size, and the viscosity of the extracellular medium or cytoplasm.
Under the influence of a DC electric field, the distribution of the molecules at the cell
surface reaches an equilibrium state when fluxes of electrophoretic migration, UC(θ), are
balanced by a diffusional flux in the opposite direction, D∇C(θ)
U Vind C(θ) = D∇C(θ)
(13)
Where U is the electrophoretic mobility coefficient of the molecule defined at eq. 12;
Vind is the applied electric field that produces effective electrophoretic force at angle θ; C(θ)
is the surface concentration of the mobile molecule at the angle θ at time t; and D is the
diffusion coefficient of the molecule. Assuming the cell resembles a nonconducting sphere
(76), the magnitude of the effective electric field at angle θ is given by equation 3.
Vind = 1.5 EapplRcosθ
29
Therefore, the analysis of the redistribution of membrane components under the
influence of a DC electric field deals with two characteristic properties of a charged
component: the effective electrophoretic mobility (U) and the diffusion coefficient (D). Thus
from the value of D/U obtained by measuring the steady state surface distribution from
equation 13 one can calculate the effective electrophoretic mobility.
Macromolecules such as charged proteins will migrate through an electrophoresis
medium when an electrical field is applied. The electrophoretic mobility depends on how
easily the protein can move through the medium and on the strength of the electrical field.
Small proteins move more readily than larger proteins, therefore have greater mobility. When
a mixture of proteins is subjected to electrophoresis, these proteins migrate through the
medium at different rates and separate on the basis of size.
Proteins are enormously complex molecules and exhibit a variety of properties in
addition to size, which can affect electrophoretic mobility. For example, proteins vary in
amino acid composition and consequently do not all have the same charge to mass ratio (77).
Proteins with a high proportion of amino acids with acidic side chains (aspartic acid, glutamic
acid) carry a net negative charge while proteins with a predominance of basic amino acids
(lysine, arginine, histidine) carry a net positive charge. Depending on the number and ratio of
acidic and basic amino acids, some proteins are weakly charged while others are strongly
charged. Oppositely charged proteins will migrate in opposite directions in an electric field.
Weakly charged proteins will have less mobility than strongly charged proteins.
Proteins also vary in tertiary and quaternary structure (77). Some proteins are globular,
others fibrous; some exist as multiple protein complexes; and some proteins, such as insulin,
consist of two or more polypeptide chains held together by covalent disulfide linkages of
cystine residues. The conformation of a protein can affect the rate of migration.
When proteins are subjected to electrophoresis, they will migrate and separate as a
function of size, charge density, and conformation. Thus each protein will migrate differently
depending on those characteristics.
1.403. Electro-osmosis in the Plasma Membrane
It has been pointed out that an electric field parallel to the surface of a cell should
redistribute charged macromolecules that are free to move laterally in the plasma membrane.
The surface of most cell membranes bears net negative charge. As a consequence, in its
vicinity the electrolyte will contain an excess of ions of opposite charge (counter-ions) and
thus will have a net charge density (figure 10). At each interface of the membrane with the
internal or external electrolytic milieux, the fixed negative charges at the membrane can be
regarded as constituting a layer that attracts an ionic layer the so called Gouy-Chapman cloud,
where cations are much more concentrated than in the bulk solution (79, 81). This "ion cloud"
exists as two parts forming an electrical double layer, and extends into solution around
charged surface of the membrane. The first layer is an inner region where the ions are strongly
bound. The second layer is an outer (diffuse) region where ions are less firmly associated, and
the ion distribution will be determined by a balance of electrostatic forces and random thermal
motion (102). In other words, since negatively charged surface of the membrane, very few
negative charges can get close to the surface because of repulsion from the charges; therefore
only positive charges tend to congregate (inner region). However, farther away from the
membrane the negative charges suffer less repulsion and besides are also attracted by the
positive charges that are bound to the negative charge of the membrane (diffuse region). At a
30
sufficient distance away from the membrane surface, the number of positive and negative
charges are evenly balanced. The imaginary surface separating the inner region from diffuse
region is known as the slipping plane, within which the counter-ions strongly bound (inner
region) with the fixed negative charges of the membrane act as a single entity showing an
elastic behavior; they act as if are attached to the negatively charged components of the
membrane. By contrast, ions outside of the plane act as if they are part of the surrounding
solution showing normal viscous behavior (81).
Inside of slipping plane, the oppositely charged ions in the inner layer decrease
linearly the electric surface potential of the membrane. Outside of slipping plane, the surface
potential decrease is not linear; it drops off exponentially as the distance from the membrane
increases until, at sufficient distance, it reaches the bulk solution value, conventionally taken
to be zero (86). That means the ion concentration decreases with the increase of the distance
and reaches to the normal concentration of the solution. The electric potential on the slipping
plane is called the Zeta Potential, ζ, and it is that potential which is measured, when it
measures the velocity of the particles in a DC electric field (102). The zeta potential depends
on the surface charge density in the double layer and the double layer thickness. The surface
charge density, in turn, depends on the concentration of electric potential-determining ions.
The thickness of the double layer depends on the concentration of electrolyte in solution
(intra- extracellular medium).
The imposition of an electric field along the membrane surface produces flow of
positive counter-ions toward cathode, dragging the bulk fluid along. Such fluid movement
results in an electroosmotic flow of fluid towards the cathode and parallel to the surface to the
cell (102, 120). The electroosmotic flow exerts a hydrodynamic force on the portion on the
mobile macromolecules that protrude from the lipid bilayer of the membrane, resulting thus in
the movement of charged and uncharged molecules, which causes that even negatively
charged macromolecules to accumulate at the negative side of the cell. The electroosmotic
flow is typically plug-like and its velocity is proportional to the zeta potential of the surface
and to the applied electric field when the surface charge distribution is uniform. The velocity
of the fluid adjacent to the cell surface is
vf = (ε ζ Vind) / η
(14)
Where ε is the permittivity of the medium; ζ is the zeta potential; Vind is the induced
transmembrane voltage (from Eq. 3); and η is the viscosity of the medium.
In conclusion, the migration of charged molecules by the application of an electric
field is under the influence of both, the electroosmotic fluid drag and electrophoretic forces on
its own charge.
1.404. Summary of effects of applying an electric field
These are the most important effects caused by the application of an electric field to
living cells, including bone cells. They are a change in the transmembrane potential,
electrophoresis, and electro-osmosis of membrane components. These effects have been
demonstrated experimentally in a variety of cells as indicated by the papers referred. When an
electric field is applied, the three effects will result on the cell at the same time. In this work,
only DC electric fields were used; therefore it is assumed that a DC electric field will polarize
the cell. Thus membrane side exposed to cathode will be depolarized, while membrane side
expose to anode will be hyperpolarized. Furthermore, a redistribution of membrane
31
components will also occur. But effects such as dispersions and deformations at the
membrane that are only dependent on the frequency will not be carried out. Therefore the
effect of applying a DC electric field will be a constant polarization of the cell and cell
components. The question is which of these effects is the cause of specific cellular responses.
This question has not been answered to date and more research is required. The problem is the
difficulty in isolating just one cellular response from one effect caused at the membrane by
the electric field. Furthermore the difficulty also lies in the fact that every cellular response
involves a coordinated system of molecules including extracellular matrix receptors and
cytoskeletal and membrane proteins. The present work contributes to the determination of the
mechanism of interaction between bone cells and DC electric fields, evaluating the
mechanical responses of the cells when exposed to the fields.
32
II
MATERIAL
AND
METHODS
INTRODUCTION
This chapter presents the material and methods used in this work. The first objective
of this chapter is to give a brief introduction of cell cultures and working in laboratory.
Subsequently it describes the details of the process to obtain bone cells of periosteum pieces
of bovine ulnae and steer radii, as well as all techniques used to analyze the behavior of cells
when subjected to a DC electric field. A brief explanation of every substance used during
biochemical studies is also mentioned here. A list of all solutions and reagents used is
presented in the Appendix.
33
2.101. Introduction to Cell Culture and Biochemical Work
In order to work with cells, cell biologists have developed ways of dissociating cells
from tissues and separating the various types, resulting in a relatively homogenous population
of cells that can be analyzed. The population cell number can also be greatly increased by
allowing cells to proliferate in culture.
The first step in isolating cells of a uniform type from a tissue that contains a mixture
of cell types is to disrupt the extracellular matrix and intercellular junctions that hold the cells
together. The resulting cut from the tissue can be then cultivated and thereby cells can
proliferate in culture. The resulting cut can also be dissociated directly into single cells by
treating the tissue with proteolytic enzymes (such as trypsin and collagenase) and chemical
agents (such as EDTA), which disrupt the binding between cells and tissue. Either method
given appropriate conditions, cells live, multiply and even express differentiated properties in
a tissue culture dish, giving rise to the establishment of cell cultures (121). The cells can be
watched under microscope or analyzed biochemically, and the effects of adding or removing
specific molecules can be explored. Experiments on cultured cells are sometimes said to be
carried out in vitro (literally, “in glass”) to contrast them with experiments in intact
organisms, which are said to be carried out in vivo (literally, “in the living organism”).
Cells proliferate, develop and survive in a culture dish if they are provided with a
suitable medium containing nutrients and specific protein growth factors. This liquid medium
is commonly referred as extracellular medium and contains specific quantities of small
molecules such as salts, glucose, amino acids, vitamins and growth factors. The extracellular
medium can be used either in combination with serum (horse serum of fetal calf serum, FCS)
or free from serum.
If the cultures were prepared directly from the tissue of an organism they are called
primary cultures. In most cases cells in primary cultures can be removed from the culture
dish and used to form a large number of secondary cultures; they may be repeatedly
subcultured in this way for weeks or months. Such cells often display many of the
differentiated properties to their origin. Most vertebrate cells die after a finite number of
divisions in culture (77). Human skin cells, for example, divide only 50 to 100 times before
they die out. It has been suggested that this limited life-span is related to the limited life-span
of the animal which cells are derived (121). Occasionally, however, some cells in a culture
will undergo a genetic change that makes them effectively immortal. Such cells proliferate
indefinitely and can be propagated as a cell line. Although all the cells in a cell line are very
similar, they are often not identical. The genetic uniformity of a cell line can be improved by
cell cloning, in which a single cell is isolated and allowed to proliferate to form a large
colony. A clone is any such collection of cells that are all descendants of a single ancestor
cells. All cells used in this work were obtained of primary cultures and they were only
allowed to divide 20 times.
Populations of cells can be analyzed physically and biochemically. Cells can be
stimulated and their physical behavior with a normal phase contrast microscope observed and
recorded. This classical method in microscopy gives good views of cell architecture, but it
provides little information about cell chemistry. In cell biology it is often important to
determine the quantities of specific molecules and to know where they are in the cell and how
their level or location change in response to extracellular signals. The molecules of interest
range from small inorganic ions, such as calcium to large macromolecules, such as specific
proteins or DNA sequences.
34
A large number of techniques are now available for detecting, measuring and
following almost any chosen molecule in a cell. Fluorescent indicators dyes, for example, can
be used to measure the concentration of specific ions in individual cells and even in different
parts of a cell. Calcium, Ca2+, is the ion most frequently measured, as Ca2+ plays an important
part in allowing cells to respond to extracellular signals (121, 122). To monitor the Ca2+
activity in the cells, fluorescent indicators are introduced into the cells and then are excited at
slightly longer wavelengths when they are free of Ca2+ than when in their Ca2+–bound form.
By measuring the ratio of fluorescent intensity at two excitation wavelengths, the
concentration ratio of the Ca2+–bound indicator to the Ca2+-free indicator can be determined;
this provides an accurate measurement of the free Ca2+ concentration. Ca2+ measurements will
be discussed in detail in section 2.106.
To introduce the fluorescent indicators to the cells many techniques have been
developed (121). One approach is to microinject the molecules into the cell through a glass
micropipette. A second approach is to create pores in the membrane by electrical shock. The
pores remain open for minutes and allow the molecules to enter to the cell. A third method is
to create membrane vesicles that contain the indicators to fuse with the cell membrane. A
fourth method is simple diffusion; it is the simplest method. Solutions with the indicators are
created to diffuse across the membrane. In this work all methods used were simple diffusion.
To analyze the dynamic behavior of intracellular components in living cells, such as
the movement of cytoskeletal proteins, sensitive methods have been developed. Green
fluorescent protein, GFP, is the most popular in the last 5 years (123). The principal
advantage of GFP is the ability to detect fluorescence in living specimens with real-time
kinetics. GFP is a protein that generates a highly visible and efficiently emitting fluorophore
and that was originally identified in the jelly fish Aequorea victoria as the responsible of its
bioluminescence (123). The cloning of the GFP and its subsequent expression in heterologous
systems established GFP as a novel genetic reporter. When expressed in cells as fusions to
many proteins provides a “fluorescent tag” on the protein, which allows its localization in
living cells when is illuminated by blue or UV light. Light-stimulated GFP fluorescence is
species-independent and does not require any cofactors, substrates, or additional gene
products.
Phase contrast microscopy provides valuable information about physical behavior and
architecture of the cells, but it does not provide information about the mechanical behavior in
the cells. Complex mechanical interactions take place at cell-substrate adhesion sites. Forces
generated at these sites are involved in vital cellular functions from migration and cell shape
to signal transduction (124). A newly developed technique, traction force microscopy, makes
it possible to visualize the dynamic characteristics of mechanical forces exerted by cells,
including the magnitude and direction (125). In traction force microscopy, living cells are
cultured in flexible polyacrylamide gels where fluorescent beads are embedded. Displacement
of the beads can be analyzed mathematically, thereby traction forces can be calculated and a
spatial distribution analyzed. Thereby the mechanical forces exerted by the cells can be
evaluated.
2.102. Preparation of Bone Cells
In chapter I it was described that all bones are lined in the external surface by a layer
of connective tissue called periosteum (see figure 1 and 2, section 1.1). Periosteum consists of
an outer layer of collagen fibers and cells. The inner, more cellular layer of the periusteum is
composed of flattened cells with the potential to divide and differentiate into bone cells
35
(osteoblasts). In this work, osteoblasts were obtained from the inner cellular layer of the
periusteum and the process used to separate them is explained here.
Osteoblasts were prepared using the outgrowth method described by Jones in 1991
(126). The preparation of periosteum pieces of bovine ulnae and steer radii obtained at the
local slaughterhouse led to a homogenous culture of osteoblasts like cells after 4-5 weeks.
Having removed all the muscles from the bone, it was cleaned using sterile phosphate
buffered saline, PBS. Then scraping of the periostal surface was done cautiously and
periosteum was slit and removed in strips carefully from the bone surface. The periosteum
strips were put in Earles solution and washed 3x10 min at 37 oC. After washing, periosteum
strips were put in new Earles medium plus antibiotics (Penicillin, Streptomycin and
Amphotericin B) and then washed 20 min at 37 °C. Periosteum strips were cut in pieces of
about 5X5 mm and 8-10 pieces were placed in a Petri dish (diameter: 14 cm) with High
Growth Enhancement Medium (HiGEM) containing 10% fetal calf serum, FCS (figure 13).
The periosteum pieces were incubated at a CO2- incubator at 37 oC (5 % CO2). After 24 hours
of incubation, the HiGEM medium was changed, and then periosteum pieces were left in the
incubator for 4-5 weeks. During this time, medium was changed once a week. After these 4-5
weeks, the outgrowing osteoblasts reached confluence. Bone cells adhered to the dish and
proliferated.
Figure 13. Periosteum strips were removed from bone and then were washed and disinfected (A). After
washing periosteum strips were cut in 5X5 mm pieces (B) and cultivated in petri dishes (C) under 5% CO2 and
37oC. After 4-5 weeks cells proliferated in the dishes.
To remove cells from the dish a trypsin solution was applied. Trypsin is a proteolytic
enzyme that acts to degrade the proteins responsible for the attachment of cells to substratum,
thereby disrupting the binding between cells and substratum. Periosteum pieces were removed
with the medium and then cells were washed two times with PBS solution. 5 ml trypsin was
applied to the cells and incubated 2-3 min for activating the disruption. Cells with the solution
were aspired with a pipette and placed in a 50 ml falcon tube with 20 ml HIGEM medium to
centrifugation. Centrifugation permits to group all cells forming the so called pellet.
Centrifugation was carried out in a maximal speed of 500g or lower for 10 min. After
centrifugation, medium was removed and cells were placed in 50 ml culture containers with
HAM’S F-10 medium plus vitamin C, Penicillin and Glutamin. HAM’S F-10 is a culture
medium containing a nutrient mixture appropiate for culturing an wide variety of cells in
laboratory like osteoblasts. A list of the components that integrate HAM’S F-10 is listed in
appendix A.
Cells in culture containers were placed at a density of 5X106/container with 4 ml
HAMS F-10 medium containing 10% FCS. One day prior to experimentation cells were
removed by trypsin treatment. 2 ml trypsin was applied to the cells in the culture containers
and then incubated 2-3 min in the incubator. 10 ml HAM’S F-10 was added to the culture
container and then the cells together with the medium were aspired with a pipette and placed
in a falcon tube to centrifugation. Centrifugation was carried out in maximal speed of 500g or
36
lower for 10 min. After centrifugation, medium was removed and cells were placed in
coverslips No. 1 (60 x 24 mm) in a density of 3000-5000 cells/cm2 with HAM’S F-10
medium containing 10% FCS.
During experimentation the coverslips were placed into the electro-chamber and filled
with Ham’s F10 medium without FCS. Cells were always 1-4 days old, and experiments were
carried out in a 37 oC room. Since in room there is not 5% CO2 necessary to help to maintain
a constant pH in the solution, 10 mM Hepes were added to the Ham’s F10 medium. Hepes is
a buffer commonly used in laboratory for stabilizing the changes of pH in cell culture
solutions such as Ham’s F10 medium. The preparation of Ham’s F10 medium with 10 mM is
indicated in appendix A.
2.103. Electric Field Chamber
The electro-chamber was based on the model of Albert Harris (127) (figure 14). The
chamber had two containers on both sides, these were filled with 0.9% saline solution and the
electrical current was introduced to these containers by means of platinum wires. To apply the
electric field to the cells, agar bridges were used. One end was placed in one container and the
other end was placed on one side of the coverslip (see figure 14). In the middle of the
chamber there was a block of polycarbonate that served to insulate the two margins of the
coverslip from each other where the agar bridges were placed. The function of this block was
to reduce the culture medium to a thin strip between the cells and bottom of the block, and
thereby producing a high resistance and confining the electric current across the cells. A pair
of sections from the polycarbonate (100 µm) was attached to the bottom of this block on both
sides so as to leave a 14 mm wide channel between them. Block is 1 cm wide. Thereby the
two parallel pieces of polycarbonate in the block had the requisite to build a bridge of 100 µm
high, 1 cm wide and 14 mm large. In both sides of the block a platinum wire is attached,
which was connected to a Fluke 75 digital multimeter to control the electric field strength
across the cells. Thereby the electric field can be measured in volts per centimeter and
adjusted to give the desired total strength across the cells. The voltage is generated by means
of a bio-rad power supply model 200/2.0 voltage-regulated. Cells were observed in real time
under the bridge.
The use of agar bridges permits the flow of current but block convection or toxic
substances from the wires (128), as well as joule heating (129), and electrolysis and electroosmosis (130). Agar bridges were prepared with 150 mg agar salt; 15 ml saline solution and
boiled in 100 oC for 5 min. The agar solution was put into a mold until it set forming the
bridge, which was like hard gelatin. The saline solution was prepared with 9 g NaCl and 1000
ml sterile water.
2.104. Phase Contrast Microscopy
The human eye is sensitive only to the colors of the visible spectrum (variations in
light frequency) or to differing levels of light intensity (variations in wave amplitude) (121).
The relationship between high and low light intensity in an image is interpreted as contrast by
the human eye. Then contrast could be defined as a difference in light intensity.
Phase contrast microscopy is an optical technique that is utilized to produce high
contrast images of transparent specimens such as living cells, microorganisms, organelles, and
thin tissue slices among others. Phase contrast technique employs an optical mechanism to
translate minute variations in phase into corresponding changes in amplitude, which can be
37
visualized as differences in image contrast (121); thereby it provides an excellent method of
improving contrast in unstained biological specimens without significant loss in resolution.
Phase contrast microscopy is widely utilized to examine dynamic events in living cells
because one of its major advantages is that living cells are examined in their natural state
without being killed, fixed, and stained. Therefore, as a result the dynamic of biological
processes in live cells can be observed and recorded in high contrast with sharp clarity of
minute specimen detail and in real time. Since these advantages offered by phase contrast
microscopy, this technique was utilized to visualize the dynamic behavior of bone cells when
subjected to an electric field.
Figure 14. Diagram of the electro-chamber used to subject the cells to different electric fields. In the
middle of the chamber there was a block of polycarbonate that served to insulate the two margins of the
coverslip from one another where the agar bridges were placed. The block had a bridge in the bottom (100 µm
high, 14 mm wide and 1 cm large) where the cells were placed and the electric field controlled.
The formation of an image in the microscope relies on a complex interplay between
two critical optical phenomena: diffraction and interference (121). When light from the
microscope lamp passes through a specimen as a living cell, some of the light passes around
and through the specimen, but does not interact with it. Light is undisturbed in its path. Such
light is called direct light or undeviated light. The background light (often called the
surround) passing around the specimen is also undeviated light. On the other hand, some of
the light passing through the specimen is deviated, which becomes scattered and diffracted
into divergent waves by tiny details and features present in the specimen. Light is disturbed in
its path. Such light is called diffracted. The light diffracted by the specimen is not reduced in
amplitude as it is in a light-absorbing object, but is slowed by the specimen because of the
specimen's refractive index or thickness (or both). This diffracted light, arrives at the image
plane out of phase with the direct light that has passed undeviated. Thus, the net effect is to
transform the relative phase difference introduced by the specimen into a difference in
amplitude (intensity) of the light emerging from the image plane.
38
In the optical microscope, image formation occurs at the intermediate image plane at
the diaphragm of the eyepiece. The direct or undeviated light is projected by the objective and
spread evenly across the entire intermediate image plane. The light diffracted by the specimen
is also captured by the objective lens and brought to focus at various localized places on the
same intermediate image plane. Thus, the image produced by the objective lens is conjugate
with the specimen, meaning that each image point is geometrically related to a corresponding
point in the specimen. It follows that each point in the specimen is therefore represented by a
corresponding point in the image. At the intermediate image plane the diffracted light causes
interference with the undeviated light. Interference is the result of two waves when are added
together. The resulting wave has an amplitude value that is either increased through
constructive interference, or diminished through destructive interference. Then the light
diffracted waves can interfere either constructively or destructively with the undeviated light
to produce a resulting wave that has either increased or decreased amplitude. If the crests of
one of the waves coincide with the crests of the other, the amplitudes are determined by the
arithmetic sum of the amplitudes from the two original waves. For example, if the amplitudes
of both waves are equal, the resulting amplitude is doubled. Such additive interference is
called constructive. In other case if the crests of one wave coincide with the troughs of the
other wave (180 degrees out of phase), the resulting amplitude is decreased or may even be
completely cancelled. Such interference is called destructive. Interference produces therefore
changes in amplitude and thereby changes in intensity, resulting in more or less dark areas.
These patterns of light and dark are recognized as the image of the specimen. Since human
eyes are sensitive to variations in intensity, the image then becomes a more or less faithful
reconstitution of the original specimen (121). Thus, phase contrast microscopy makes
possible to watch the dynamic behavior of the cells in “real time”.
The behavior of osteoblasts during electric field application was examined using a
Leitz Diavert inverted phase contrast microscope equipped with a Nikon 40X objective.
During microscopic observations all cell responses were filmed with a CCD camera (Xillix
microImager MI1400) placed in the microscope and interfaced to a computer (figure 15).
Images were captured using the IMAGE PRO program (version 4.5 for windows). Control of
capture was carried out manually; all images were saved in TIF format for a subsequent
analysis.
Figure 15. Diagram of the system used to analyze the behavior of osteoblasts when subjected to DC
ELFs.
39
2.105. Fluorescence Microscopy
Fluorescence is the property of some atoms and molecules to absorb light at a
particular wavelength and to subsequently emit light of longer wavelength after a brief
interval, termed the fluorescence lifetime (131). The category of molecules capable of
undergoing electronic transitions that ultimately result in fluorescence are known as
fluorescent probes, fluorochromes, or simply dyes. Fluorochromes that are conjugated to a
larger macromolecule (such as a nucleic acid, lipid, enzyme, or protein) through adsorption or
covalent bonds are termed fluorophores (132). In general, fluorophores are divided into two
broad classes, termed intrinsic and extrinsic. Intrinsic fluorophores, such as aromatic amino
acids and green fluorescent protein, are those that occur naturally. Extrinsic fluorophores are
synthetic dyes or modified biochemicals that are added to a specimen to produce fluorescence
with specific spectral properties. Fluorophores are extremely valuable in biological
applications, because when a fluorophore is designed to bind selectively to a particular
component of the cell, it enables researchers to detect that specific component with exquisite
sensitivity and selectivity. Fluorescence microscopy is the method utilized to detect the
changes of fluorescence emitted by fluorophores (131, 132).
The atom is the source of all forms of electromagnetic radiation, whether visible or
invisible. Higher-energy forms of radiation, such as gamma waves and X-rays, are produced
by events that occur to disrupt the nuclear stability of the atom. Radiation having lower
energy, such as ultraviolet, visible, and infrared light, as well as radio and microwaves,
originate from the electron clouds that surround the nucleus or the interaction of one atom
with another. These forms of radiation occur due to fact that electrons moving in orbits
around the nucleus of an atom are arranged in different energy levels within their probability
distribution functions. Many of the electrons can absorb additional energy from external
sources of electromagnetic radiation that results in their promotion to an inherently unstable
higher energy level. Eventually, the excited electron loses the extra energy by emitting
electromagnetic radiation of lower energy and, in doing so, falls back into its original and
stable energy level (131).
The fluorescence process is governed by three important events (131): 1) Absorption
of energy by the fluorochrome. Excitation of the fluorochrome is carried out by an incoming
photon of energy that is supplied by an external light source. The photon is absorbed by the
fluorochrome and then collides with its electrons, exciting and elevating them to higher
energy levels, creating thus an excited state; 2) Transition or relaxation in the excited state.
The excited electrons lose some energy and return to the so-called lowest excited singlet state;
3) Emission of a longer wavelength photon. From the lowest excited singlet state, the
electrons drop back to the ground state with simultaneous emission of fluorescent light,
returning thus the fluorochrome to its ground state.
When electrons go from the excited state to the ground state, there is a loss of energy.
As a result, the emission spectrum is shifted to longer wavelengths than the excitation
spectrum (wavelength varies inversely to radiation energy). Therefore the emitted light is
always of longer wavelengths than the excitation light. This phenomenon is known as Stokes'
Law or Stokes' shift (131). The greater the Stokes' shift, the easier it is to separate excitation
light from emission light. The emission intensity peak is usually lower than the excitation
peak; and the emission curve is often a mirror image of the excitation curve, but shifted to
longer wavelengths. To achieve maximum fluorescence intensity, the fluorochrome is usually
excited at the wavelength at the peak of the excitation curve, and the emission is selected at
the peak wavelength of the emission curve. Appropriate filters control the selections of
40
excitation wavelengths and emission wavelengths. The basic task of the fluorescence
microscope is to permit excitation light to irradiate the specimen and then to separate the
much weaker re-radiating fluorescent light from the brighter excitation light. Thus, only the
emission light reaches the eye or other detector. The resulting fluorescing areas shine against
a dark background with sufficient contrast to permit detection. The darker the background of
the non-fluorescing material, the more efficient the instrument.
The fluorescence process is cyclical; the same fluorescent dye can be repeatedly
excited and detected. Thus the fact that a single fluorescent dye can generate many thousands
of detectable photons is fundamental to the high sensitivity of fluorescence detection
techniques (131).
The detection of fluorescence requires 4 essential elements: the source of excitation,
the fluorescent dye, wavelength filters to isolate emission photons from excitation photons,
and a detector that registers emission photons and produces a recordable output, in this case, a
photographic image. In this work fluorescence microscopy was used to evaluate cellular
calcium response, changes in membrane potential, and the dynamic behavior of actin
cytoskeleton. The system was based on a fluorescence inverse microscope (DIAPHOT,
Nikon) equipped with a Nikon 40X fluorescent objective. The source of excitation,
fluorescent dyes, filters, as well as detectors were adapted to the microscope in every
fluorescent measurement. Calcium imaging is the first technique described here.
2.105a. Calcium Imaging
The ion calcium, Ca2+, is one of the most important intracellular messengers in the
cells, which can transmit the information received in the cell membrane surface to specific
targets within the cell. Previous reports have demonstrated that some cells respond with a
cytoplasmic calcium increase when subjected to electric fields (99, 107). To evaluate if the
electric field was causing a calcium increase in the cytoplasm of osteoblasts, intracellular
calcium activity in cells under field exposure was measured using fluorescence microscopy.
The fluorescent dye used for analyzing intracellular calcium activity was fura 2, which
enters to the cell and binds to the intracellular calcium ions (131). When fura 2 is excited
emits a fluorescence that is proportional to the amount of bound ions (calcium concentration).
Fura 2 is loaded into the cells by means of Acetoxymethyl (AM) ester. AM ester is a noninvasive technique that is by far the most popular method for loading fluorescent ion
indicators (132). The carboxylate groups of indicators for Ca2+ and other cations are
derivatized as acetoxymethyl esters, rendering the indicators permeant to membranes and
insensitive to ions. Once inside the cell, these derivatized indicators are hydrolyzed by
ubiquitous intracellular esterases, releasing the calcium indicator. Molecular probes provides
the complex Fura 2-AM; therefore it is only necessary to generate an aqueous solution for its
permeation into the cell. In this case dimethyl sulfoxide (DMSO) was used.
The fluorescence excitation spectra of fura 2 shifts to shorter wavelengths as calcium
concentration increases, much as the absorption spectra do. The maximal absorption peaks are
335 and 363 nm at maximal and minimal calcium concentration respectively (figure 16). The
fluorescence emission peak of both calcium-bound and calcium-free forms of fura 2 is 505
nm (131, 132). Therefore fluorescence emission is independent of calcium concentration.
41
Figure 16. Fluorescence excitation (excited at 340 nm) and emission (detected at 510 nm) spectra of
calcium-saturated (A) and calcium-free (B) fura 2. Graphic was taken of reference 131.
Fluorescent dyes that show an excitation or emission spectral shift upon ion binding
can be calibrated using a ratio of the fluorescence intensities measured at two different
wavelengths. Measuring the ratio of intensities has profound implications. The ratio
measurement is independent of dye concentration and optical path length, parameters that
cannot be controlled within a cell. Degradation of the dye due to prolonged exposure to the
excitation source and variations in the excitation intensity are also compensated for by the
ratio technique. The result is therefore the cancellation of artifactual variations in the
fluorescence signal that might otherwise be misinterpreted as changes in ion concentration
(131). For ratiometric measurements of fura 2, excitations at 340 nm and 380 nm have been
usually preferred (132). At these wavelengths the fluorescence intensities have an opposite
ion-sensitive response. At 340 nm the fluorescence intensity of Ca2+-bound fura 2 increases.
At 380 nm the fluorescence intensity of Ca2+-bound fura 2 decreases. Thus the ratio of 340
nm/380 nm due to the change of intracellular ion concentration is unambiguously identified.
The ratioing fluorescence intensities are detected usually at ~510 nm. From this ratio, the
level of intracellular calcium can be estimated using dissociation constants (Kd) that are
derived from calibration curves corresponding to a series of precisely manipulated ion
concentrations (131). By using the ratio of fluorescence intensities produced by excitation at
two wavelengths, factors such as uneven dye distribution, leakage, photobleaching, and
problems associated with measuring calcium in cells of unequal thickness are minimized
because they should affect both measurements to the same extent.
Once the indicator has been calibrated with solutions of known calcium
concentrations, the following equation (method of Grynkiewicz) can be used to relate the
intensity ratios to calcium levels (132).
[Ca2+] = Kd Q [(R-Rmin)/(Rmax-R)]
(15)
Where R represents the fluorescence intensity ratio Fλ1/Fλ2, in which Fλ1 is the
fluorescence intensity of the excitation wave length in 340 nm; and Fλ2 is the fluorescence
intensity of the excitation wave length in 380 nm. Ratios corresponding to the titration end
points are denoted by the subscripts indicating the minimum and maximum calcium
concentration. Q is the ratio of Fmin to Fmax at λ2 (~380 nm). Kd is the calcium dissociation
constant of the indicator. Calibrating fura indicators requires making measurements for the
completely ion-free and ion-saturated indicator (to determine the values for Fmin, Fmax, Rmin
and Rmax) and for the indicator in the presence of known calcium concentrations (to determine
Kd).
42
To evaluate calcium responses in osteoblasts, 300 micromolar, µM, Fura 2 was
prepared with dimethyl sulfoxide, DMSO, as a stock solution. Cells in coverslips were loaded
with 3 µM fura 2 AM for 60 min at room temperature or 37 oC. After loading, cells were
washed twice with PBS. After washing the coverslip was mounted in the electro-chamber and
normal 10 mM Hepes medium was applied. Fluorescence emission was collected from a
group of 4 ± 1 cells. Excitation wavelength was alternated between 340 nm and 380 nm and
fluorescence intensity (F340 and F380 respectively) was monitored at 505 ± 10 nm. Calcium
concentration [Ca2+]i was expressed as the ratio of F340/380, an indicator of Ca2+ activity
(132).
Experiments were carried out in a temperature controlled room at 37 oC, or in a
temperature uncontrolled room. When cells were in the temperature uncontrolled room,
medium (37 oC) was kept hot prior to stimulation. Calcium was visualized for a maximum
period of 10 min. 4 ± 1 cells were selected to visualization. Cells were 1-4 days old and
cultured in a density of 5000 cells/cm2 for every experiment.
The system used for calcium measurements consisted of the fluorescence inverse
microscope, which was connected to a Xenon lamp with a motorized monochromator
(Applied Imaging, UK) for excitation of the fluorescent dye. The fluorescence emissions were
recorded with a double camera system (Photonic Science) interfaced also to microscope; both
monochromator and camera were interfaced to a computer (figure 17). The whole system is
controlled by Quanticell 700m Ver.2.20 software (Applied Imaging, UK). The
monochromator is controlled by the computer system and allows switching between user
configurable wavelengths; the video signal is captured with a frame grabber system that
supports dual excitation wavelength experiments. Images captured are showed in pixels with
possible 256 different intensities. Furthermore images captured are averaged by the system to
eliminate intensity variations due to variation of the light source or scattered light. Digital
dates are saved in a 128 video RAM during the measurement at the same time they are
showed at monitor. Thus a “real time“ measurement could be observed.
Figure 17. Diagram of the system used to monitor intracellular calcium activity.
2.105b. Membrane Potential Measurements
The process of measuring the changes in membrane potential was very similar to
calcium measurements; the system used was exactly the same: monochromator and camera
interfaced to a computer and controlled by Quanticell 700m Ver.2.20 software. The only
exception is that membrane potential changes do not require a ratiometric measurement. The
43
fluorescent dye used for membrane potential measurements does not show an excitation or
emission spectral shift upon ion binding. Therefore the fluorescence detected is directly
related to the change of potential.
The membrane potential dye used was DiBAC4 [3]. DiBAC4 [3] is a bis-barbituric acid
oxonol with maximal peak excitation at approximately 490 nm and a fluorescence emission at
520 nm wavelengths. The dye enters depolarized cells where it binds to intracellular or
membrane proteins and exhibits enhanced fluorescence and red spectral shifts. Increased
depolarization results in more influx of the anionic dye and thus an increase in fluorescence.
Conversely, hyperpolarization is indicated by a decrease in fluorescence (131). DiBAC4 [3]
has been reported as a membrane potential indicator in bone cells previously (133).
To evaluate changes of membrane potential in osteoblasts, 10 mM DiBAC4 [3] was
prepared with DMSO as stock solution. Cells on coverslips were loaded with 1 µM DiBAC4
(3) for 30 min at 37 oC. After loading cells were washed twice with PBS and the coverslip
was mounted in the electro-chamber with normal 10 mM Hepes medium.
2.105c. Visualization of Actin Filament Dynamic
Visualization of actin filaments was carried out with the green fluorescent protein,
GFP. The GFP from jellyfish aequorea victoria is a photoprotein that emits light when is
excited by blue light produced by aequorin photoprotein (123). The cloning of the wild-type
GFP gene and the subsequent expression in heterologous systems established GFP as a novel
genetic reporter system. Thus the use of GFP when expressed as fusions to many proteins (in
either eukaryotic or prokaryotic cells) provides a “fluorescent tag” on the protein, which
allows its localization in living cells when is illuminated by blue or UV light. Lightstimulated GFP fluorescence is species-independent and does not require any cofactors,
substrates or additional gene products from A. Victoria. Its biggest absorbance peak is at 395
nm with a smaller peak at 475 nm. Excitation at either of the two wavelengths results in
emission of green light at 508 nm (134). Several GFP chromophore variants have been
developed to improve the utility of GFP.
The GFP-Calponin was used to visualize stress fibers in cells exposed to electric
fields. The basic Calponin, CaP h1, is a protein that has been reported to incorporate along
into the stress fibers (135). Thereby the fusion of GFP with CaP h1 proved an indirect way to
visualize stress fibers without perturbation of the actin-myosin complex.
To incorporate GFP-h1 calponin into osteoblasts, the transfection was performed with
Effectene (Qiagen, Hilgen, Germany). Effectene Transfection Reagent is an innovative nonliposomal lipid formulation that is used in conjunction with a special DNA-condensing
enhancer and optimized buffer to achieve high transfection efficiencies (136). The enhancer
first condenses the DNA molecules and Effectene Reagent subsequently coats them with
cationic lipids (see figure 18), providing a particularly efficient way of transferring DNA into
eukaryotic cells. Effectene Reagent offers significant advantages over many liposome
reagents and other transfection methods:
•
•
•
•
•
Fast and easy transfection
No transfection-complex removal needed for most cell lines
Transfection in the presence of serum
Ideal for primary cells or sensitive cell lines
High transfection efficiencies and minimal cytotoxicity
44
•
Low DNA requirement
Transfection was performed according to the manufacture’s instructions. Briefly, 0,5
µg EGFP-CaP h1 plasmid was mixed with 4 µl enhancer and 145 µl transfection buffer and
incubated for 10 min at room temperature. 25 µl Effectene were added to the plasmid solution
and incubated for 10 min. After 10 min, the resulting complexes were mixed with 3 ml
growth medium and added directly to the cells. The cells were then incubated for 24 hours at
37 oC and 5% CO2. Stress fibers could be observed in 70 % of cells after time. Cell density
was between 0.5-1 X 106/container. Fluorescent cells were passaged to coverslips by trypsin
process.
The system used for analyzing the dynamic behavior of cytoskeletal filaments
consisted of the fluorescence inverse microscope connected to a Xenon lamp and a CCD
camera (Xillix microImager MI1400) that was interfaced to a computer. A filter was
incorporated between lamp and microscope to permit an exact fluorescence excitation at 475
nm. Fluorescence emission was limited with a filter to a range of 505-560 nm. Analysis and
capture of the images were carried out with the IMAGE PRO program (version 4.5 for
windows). Control of capture of images was carried out manually and was done in specific
time intervals (normally every 5 minutes). All images captured were saved in TIF format.
Figure 18. Model of Effectene Principle. The Effectene procedure has two steps. DNA is first mixed
with Enhancer and a buffer that provides optimal salt conditions for efficient DNA condensation. Effectene
Reagent is then added and the mixture is incubated to allow Effectene–DNA complexes to form. The complexes
are mixed with growth medium (which can contain serum and antibiotics), and added directly to the cells. The
cells are then incubated until harvested and analyzed for gene expression. Picture taken of reference 136.
45
2.106. Traction Force Microscopy
Traction force microscopy is a new technique recently developed that allows the
evaluation of the mechanical forces carried out by the cells (125). Cells in culture elaborate
secreted matrix components into an assembled matrix known as extracellular matrix that is
adsorbed to the surface on which the cells are growing (121). Specialized regions of the cell
membrane called focal adhesions mediate adhesion of cells to the extracellular matrix through
transmembrane proteins called integrins, which bind simultaneously the extracellular matrix
on the outside and actin cytoskeleton inside (figure 19). Traction forces are the forces
generated against cell-substratum adhesive contacts by the cytoskeleton-extracellular matrix
linkages. The force generated is a contractile force that is the tensional force generated within
contractile microfilaments, which pulls inward on the surface membrane. In turn, the
substratum locally exerts an equal and opposite traction force on the cell via the same
attachments, with magnitude depending on the susceptibility of the attachments to disruption.
The effective inward-directed forces must essentially be in balance with the traction force
provided by dynamic cell substratum attachments in order to maintain a cellular force balance
(137, 138). Thus the actual traction exerted by the cell on its substratum is directly related to
the intracellularly generated contractile force.
Figure 19. In traction force microscopy cells are cultivated in flexible gels that contain fluorescent
beads. Focal adhesions mediate adhesion of contractile filaments to the extracellular matrix through
transmembrane proteins called integrins. Forces generated against cell-substratum adhesive contacts by the
cytoskeleton-extracellular matrix linkages result in the displacement of the fluorescent beads (double arrows).
The motion of these beads allows calculating the traction forces and their spatial distribution in the cells.
The stability of cytoskeletal structure and cell shape results from the cell’s ability to
bring the internal tensional forces into balance. A change in the cellular force balance can
result in integrated changes in cell, cytoskeletal and nuclear form and influence a number of
cell functions (137, 138). Therefore the knowledge of forces governing the behavior of the
cells is critical to understand their mechanical interactions. This makes traction force
microscopy an excellent tool for the detection of the mechanical response of the cells when
subjected to DC electric fields.
In traction force microscopy, living cells are cultured in flexible polyacrylamide gels
that contain fluorescent microbeads directly beneath the surface of the gel (139). The
displacement of these beads can be tracked in the vicinity of attached cells, and the resulting
46
displacement field can be measured to determine the mechanical strain induced in the gel by
the overlying cells (figure 19). Mathematical algorithms allow calculating the traction forces
that arise from cell-substrate mechanical interaction as well as their spatial distribution
beneath individual cells (125).
2.106a. Preparation and Characterization of Flexible Substrates
Flexible substrates were prepared with thin sheets of collagen-coated polyacrylamide
gels using a method previously described (125, 140). The first step was the preparation of
coverslips where the polyacrylamide sheets should be placed:
1. - Coverslips are passed through inner flame of Bunsen burner.
2. - 1 drop of 0.1 NAOH is applied to coverslips.
3. - Coverslips are placed under fume hood for air-drying.
4. - 1 drop of 3-aminopropyltrimethoxysilane is applied.
5. - After 5 min, coverslips are washed 3 times during 5 min with distilled water.
6. - Coverslips are incubated with phosphate buffer solution containing 0.5 %
glutaraldehyde for 30 min under fume hood.
7. - Coverslips are washed 3 times during 10 min with distilled water.
8. - Coverslips are placed under fume hood for drying.
9. - After drying coverslips are stored.
The polyacrylamide substrates were made with the mixture of acrylamide (Bio-Rad,
USA), bis-acrylamide (Bio-Rad, USA), and distilled H2O. The rigidity or flexibility of the
substrate is determined by the percentage of bis-acrylamide concentration. In this work the
final concentration was 8% acrylamide and 0.04 to 0.06% of bis-acrylamide. Fluorescent
latex beads of 0.2 µm and 0.5 µm were added to the acrylamide mixture in volume ratio of
2%. Polymerization of the mixture was carried out by addition of 0.05 % ammonium
persulfate (Bio-Rad) and 0.005 % tetramethyl ethylenediamine (Bio-Rad). 14 µl of this
solution were immediately placed onto the surface of a coverslip and the droplet was flattened
using a circular coverslip of 16 mm diameter (like representation in figure 17). After
polymerization the circular coverslip was removed. Coverslips are washed 2 times during 15
min with HEPES solution.
To provide a physiological surface for cell culture, a saturating density of type I
collagen was covalently attached to the base surface of the acrylamide gel (140). 200 µl of
freshly-made Sulfo-SANPAH were applied to coverslips (5 mg Sulfo-SANPAH in 10 ml
HEPES), and then coverslips were exposed to UV light at distances of 40 cm during 10 min.
Sulfo-SANPAH solution was retired of coverslip and process was repeated again. Coverslips
are washed with 50 mM HEPES for 3-4 min. Gels in coverslips are covered with 0.2 mg/ml
collagen type I (400 µl in 5 ml HEPES solution), and then they are incubated overnight at 4
o
C. Gels are washed 3 times during 5 min with PBS, and then sterilized under UV light.
Finally coverslips with flexible gels are kept in fridge until their use.
A method based on the Hertz theory (141) was used to determine the flexibility of the
polyacrylamide substrata. An atomic force microscope (AFM) with a small ball at the top of
the cantilever was used to press on the polyacrylamide sheets (142). The total indentation
caused by the ball that was fixed to the cantilever was between 0.5 µm and 1.5 µm. (Maximal
displacements caused by the cells are about 1 µm on soft substrates). Balls were used at the
tip of the cantilever with a radius between 2-10 µm. Young's modulus was calculated as Υ =
3 (1-ν2) F / 4 d3/2r1/2 , where F is the force exerted on the sheet, d is the indentation of the
47
substrate, r is the radius of the steel ball, and ν is the Poisson ratio of polyacrylamide, which
was assumed to be 0.3 (143). Values for the used sheets were about 4,400 N/m2 (s.d. < 1%)
for the 0.04% and 6,800 N/m2 (s.d. < 1%) for 0.06% bis-acrylamide substrates.
2.106b. Calculation of Traction Forces
The mechanical properties of the gel allow the cell to deform the substratum (144). To
address the question if applying an electric field to the cell causes a change in the traction
force vector map, only differential traction vectors were calculated; thus the change in traction
force that occurred in the cell during electric field application was the only dependent on it.
The image with no electric field was compared with the tensioned cell image with the electric
field applied. The resulting vectors only showed the change in force between these two states
and not the total traction force generated by the cell. This made it easier to correlate the
reaction of the cell to the applied electric field.
To calculate the deformations of the substrate, a computer program based on pattern
recognition using a cross-correlation algorithm was used (125). Thus deformations of the
substrate were determined as a matrix of vectors, by comparing the fluorescent light patterns
caused by the embedded beads, in the cell during rest and in cell subjected to the electric field.
Briefly, the first step was to record digitized images of the fluorescent beads embedded in the
substratum (8 bits per pixel, 512 by 384 pixels, 0.303 µm per pixel) with the cell during rest
(image 1), and with the cell subjected to the electric field (image 2). The tracing of the cell
nucleus and of the lateral cell boundary are drawn manually for both images using phase
contrast images and a custom interactive program (in this case IMAGE PRO, version 4.5 for
windows). The digitized images are superimposed on the tracing of the cell and then the
images are divided into small square areas. Every square is a determined pattern. The
computer program then tries to match the bead pattern in each small square of image 2 (cell
subjected to the electric field) against the pattern in different regions of the image 1 (cell in
rest). Deformation vectors (arrows) are then drawn from the position in the image 1 to the
position in image 2 with the best match. The degree of match is scored with a normalized
cross-correlation equation, which yield a value of 1 for a perfect match and 0 for no
similarity. No vector is assigned if this value falls below a user-defined threshold. The size of
the square, the distance for the pattern search, and the threshold for positive identification are
determined empirically. Coordinates defining the deformation field and cell boundary are also
input into a supercomputer and analyzed with a maximum likelihood algorithm, thus
computer generates the traction vectors throughout the cell (125). Average compressive stress
is calculated by averaging the absolute values of stress vectors projected onto specified
directions. The projection was obtained by multiplying the magnitude of the vector with the
cosine of the angle between the vector and the direction of projection. Finally, the images are
represented by drawing a small arrow with its base at the center of each square. This arrow
has the direction of the best-fit traction vector and a length proportional to the magnitude of
the traction. For the calculation of the traction force vectors, the LIBTRC software developed
by Dembo (125) for a Linux platform (Red Hat Linux 7.2) was used.
2.107. Analysis of the Calcium Role
Intracellular Ca2+ increase can result from Ca2+ release from intracellular stores or
from the opening of Ca2+ channels located on the membrane that permit Ca2+ influx into the
cells (145-149). Both possibilities were investigated in this work. Ca2+ channels located on the
membrane can be opened by voltage or stress. Those activated by voltage are known as
48
voltage-sensitive calcium channels, VSCC; and those activated by stress are known as stresssensitive calcium channels, SSCC.
To determine the path and the possible Ca2+ role in cells under electric field exposure,
a set of biochemical experiments with thapsigargin, manganese, cadmium, lanthanum,
nifedipine and nitrendipine were carried out to block the calcium response and determine the
possible transduction pathway. All experiments were carried out at 37 oC. Ca2+ activity was
visualized for a maximum period of 10 min. 4 ± 1 cells were selected for visualization. Cells
were 1-4 days old and cultured at a density of 5000 cells/cm2.
2.107a. Thapsigargin
The tumor promoter thapsigargin is recognized to selectively inhibit the Ca2+-ATPase
activity of the endoplasmic reticulum, impeding the reabsorption of calcium and thereby
depleting the Ca2+ content of the endoplasmic reticulum by an endogenous Ca2+ leak (145,
146). Practically calcium is released from the endoplasmic reticulum through normal drift. All
calcium is released from the endoplasmic reticulum within 30 min after thapsigargin
treatment, and thus calcium is not available to be released when inositol triphosphate
receptors in endoplasmic reticulum are activated.
Thapsigargin was prepared with dimethyl sulfoxide (DMSO) in a 500 µM stock
solution. Cells were mounted in the electro-chamber and placed for observation as described
for previous experiment (37 oC). Medium with 5 µM thapsigargin was applied for 40 min.
After 40 min, the electric field was applied without removing the medium with thapsigargin.
Cell responses were observed during the next 3 hours.
To visualize Ca2+ activity under thapsigargin effect, cells were loaded with fura 2 as
described and then cells were washed twice with PBS. After washing, cells were incubated
with medium with 5 µM thapsigargin for 40 min. Medium was not removed and the electric
field was applied. Calcium activity was measured as described previously.
2.107b. Manganese
The affinity of Manganese (Mn2+) for fura 2 is 42 times greater than the affinity of
Ca2+ (131). Mn2+ quenches the Fura 2 fluorescence causing a fluorescence decrease.
Fluorescence quenching can be defined as a bimolecular process that reduces the fluorescence
quantum yield without changing the fluorescence emission spectrum (132). Since manganese
ions may use the same entry pathway as Ca2+, extracellular Mn2+ can act as an indicator for
the activity of calcium influx into the cell (147). To study Mn2+ influx and the effects of this
cation on Ca2+ entry, the fluorescence has to be independent on concentration of intracellular
Ca2+ but decreases when Mn2+ quenches fura 2. Since the absorption of fura 2 excited at 360
nm is not affected depending on calcium concentration (section 2.105a), the decline of
fluorescence excited at 360 can be used to monitor manganese influx into the cells (148).
50 mM Manganese was prepared with sterile water as a stock solution. After fura
loading cells were incubated with 2 mM Mn2+ 10 min prior to stimulation. Then medium was
not removed and the electric field was applied. Fluorescent traces were observed with an
excitation wavelength of 360 nm.
49
2.107c. Cadmium
Cadmium (Cd2+) is a divalent cation that is known to be a potent calcium channel
blocker (149). It enters cells primarily by passing through all classes of voltage-sensitive
calcium channels, VSCC, and thereby inhibits influx of calcium (150). It has been reported
that Cd2+ in low concentrations (< 10µM) inhibit L and N type voltage-dependent calcium
channel and in higher doses block all Ca2+ channels (151).
10 mM Cd2+ stock solution was prepared with sterile water. Cells were incubated with
different Cd2+ concentration 10 min prior to experiment. After 10 min, Cd2+ was not removed
and the electric field was applied. Ca2+ activity was measured as described previously.
2.107d. Lanthanum
Lanthanum (La3+) is a trivalent cation that can be used to displace extracellular bound
calcium and inhibit calcium transport across biological membranes (152). La3+ blocks all Ca2+
entry into cells, including that entering through the passive membrane leak pathway (152).
10 mM La3+ was prepared with sterile water as a stock solution. Cells were incubated
with 100 µM La3+ 10 min prior to experiment. Medium with La3+ was not removed and the
electric field was applied. Ca2+ activity was measured as described previously.
2.107e. Nifedipine and Nitrendipine
Nifedipine and nitrendipine are two 1,4-dihydropyridines. 1,4-dihydropyridines act by
selectively blocking L-type VSCCs in the plasma membrane (122, 153). Both drugs were
dissolved in DMSO to produce a 10 mM stock solution. Cells were incubated with 10 µM for
10 min prior to stimulation. Since nifedipine is sensitive to light, cells were focused, light was
turned off and only a very low red light remained. Medium was removed and medium with
nifedipine was applied. Red light was turned off and after 10 min of incubation cells were
subjected to stimulation. During the experiment the monitor was covered to prevent light
changes. Nitrendipine is not sensitive to light. Ca2+ activity was measured as described
previously.
2.108. Actin Filament Disruption
To investigate the role of actin filaments in the physical behavior showed by the cells
when subjected to DC ELFs, Latrunculin A and Cytochalasin B, two potent inhibitors of actin
filament polymerization were used.
2.108a. Latrunculin A
Latrunculin A (Lat A) is a potent drug that causes actin depolymerization and cell
rounding (154-156). It binds actin monomers and thereby inhibits actin polymerization. Thus
if filaments turn over, newly freed monomers will be bound by Lat A and thereby be
incapable of reassembly to create new filaments synthesis. Practically cell rounding is reached
after 50-60 minutes with Lat A treatment.
250 µM Lat A was prepared with DMSO as a stock solution. Cells on coverslips were
mounted in the electro-chamber with normal Hepes medium and fixed to microscope.
Medium was removed and new medium with latrunculin A in a 100 nM concentration was
50
applied. After 30 min of incubation, without washing out the medium, cells were subjected to
the electric field. Whole actin depolymerization was visualized as cell rounding; this was
obtained with more time of incubation (60-70 minutes). When cell rounding was evident,
medium with Lat A was removed and then cells were washed twice with PBS. After washing,
normal Hepes medium was applied and then cells were subjected to the electric field.
To evaluate the possible role of actin cytoskeleton in the intracellular calcium activity
of osteoblasts subjected to electric fields, experiments with Lat A and fura 2 were carried out.
Cells were incubated with fura 2 as previously mentioned. Cells were mounted in the electrochamber, focused and fixed to the microscope. Medium was removed carefully and new
medium with Lat A (100 nM) was applied. After 70 min, cell rounding was very evident in
the cells and then cells were subjected to the electric field without removing the medium. Ca2+
activity was measured as described previously.
2.108b. Cytochalasin B
Cytochalasin B, CB, disrupts microfilament organization and caps the barbed end of
actin filaments and accordingly inhibits further elongation from the barbed end (157, 158).
5 mM CB was diluted in DMSO as a stock solution. Coverslips with the cells were
mounted in the electro-chamber, which was fixed to microscope. Medium was removed and
new medium with 2 µM CB was applied. After 30 min of incubation, without washing out the
medium, cells were subjected to the electric field.
2.109. Myosin Light Chain Kinase Inhibition
Myosin II in combination with actin filaments are responsible for generating the
cytoskeletal contractility and the traction forces in the cells (138). Myosin II is activated by
the enzyme myosin light chain kinase, MLCK, in the presence of calcium/calmodulin (159).
Inhibition of myosin II was carried out to investigate its role in the physical response showed
by the cells when exposed to DC electric fields. Inhibition of myosin II activation by
Ca2+/calmodulin-dependent MLCK was carried out with two structurally different inhibitors
of MLCK (ML-7 and Wortmannin).
2.109a. ML-7
ML-7 is a synthetic napthalenesulphonyl compound that is structurally unrelated to
ATP but can bind specifically at or near the ATP-binding site of MLCK, blocking ATP
binding to MLCK (159-161). Therefore, it inhibits the transfer of phosphate from MLCK to
its substrate, myosin light chain, which is required for myosin II activation.
5 mM ML-7 was prepared with DMSO as a stock solution. Cells were mounted in the
electro-chamber and then incubated for 60 min with different ML-7 concentrations. After 60
min of incubation the electric field was applied without removing the medium.
2.109b. Wortmannin
Wortmannin is well known as potent inhibitor of MLCK that has been shown to
reduce MLC phosphorylation (162-164). Wortmannin acts at or near to the catalytic domain
site of MLCK. The binding site of Wortmannin exist at, or near to the ATP binding site of
MLCK, thus it acts as a non-competitive inhibitor with respect to ATP (162).
51
1 mM Wortmannin was prepared with DMSO as a stock solution. Cells were mounted
in the electro-chamber and then incubated for 60 min with different Wortmannin
concentrations. After 60 min of incubation, the electric field was applied without removing
the medium.
2.110. pH Measurements
The pH of the medium was measured by a pH meter (Inolab pH level 1, wtw). Prior to
the experiment, the pH meter was calibrated with a standard solution to 7.2. Prior to the
application of a DC ELF, the electrode was inserted at one side of the chamber and pH was
measured. Then the electrode was changed to the other side and pH was measured again. At
both sides pH was the same. When the DC ELF was applied, pH was measured at both sides
of the bridge in the electro-chamber. Changes in Volt per pH change were calculated.
52
III
RESULTS
INTRODUCTION
This chapter describes the cellular responses observed in osteoblasts when exposed to
DC electric fields. To describe the effects caused in the cells by the application of a DC
electric field, (DC ELF), this chapter was divided in two parts. The first part describes only
the physical behavior of the cells under DC ELF exposure. Experiments with phase contrast,
fluorescence, and traction force microscopy were coordinated to characterize the physical
behavior undertaken by the cells. The second part relates to the biochemical experiments that
were carried out to identify the possible signaling mechanism in the cells exerted by the
electric field.
53
3.1. BEHAVIOR OF OSTEOBLASTS UNDER ELECTRIC FIELD EXPOSURE
In previous reports osteoblasts have responded in distinct ways when were subjected
to DC ELFs (5-7). Hence, the first objective of this work was to characterize the effects of
different DC ELF strengths on the physical behavior of the cells. The changes in the physical
behavior were observed with phase contrast, fluorescence, and traction force microscopy,
thereby a better characterization could be carried out. Cells were observed in a group of
approximately 10 cells/experiment, which were 1-4 days old. Field strengths were chosen
according to previous works with osteoblasts and other cells under DC ELF exposure (5, 127,
165, 166), thus a range between 1 to 20 V/cm was selected.
To determine the physical changes caused by DC ELFs in osteoblasts, the changes in
cell shape and cytoskeletal architecture were observed. The cytoskeleton is a three
dimensional network of filamentous proteins that fills the space between organelles and gives
shape and structure to cells. Besides this it also provides the cells with the ability to move.
Three main protein systems constitute the cytoskeleton, these are: Microfilaments,
Intermediate filaments and Microtubules. All three constituents are dynamic structures, they
constantly change shape through cycles of polymerization and depolymerization. It has been
suggested that microfilaments play the principal role in determining the shape and structure in
cells, as well as in providing force and motility (121, 167, 168). Therefore the effects of ELFs
in cytoskeletal architecture only on microfilaments were observed.
The first step before the application of DC ELFs was to test if the electro-chamber
could have an effect in the cells; the electro-chamber was used with normal culture medium to
host the cells without electric field. Cells were observed for 24 hours and in all cases cells
looked viable and no specific cell shape could be identified.
The first experiments with DC ELFs were carried out with the highest field strength
and then subsequently it was decreased. The initial cellular response adopted by the cells was
a pronounced retraction of their lamellar extensions exposed directly to the field lines and
subsequently a total retraction. As ELF strengths were gradually decreased, the retraction in
the cells were less pronounced. When the field strengths were in a range of 7-12 V/cm,
following the retraction phase, cells demonstrated a highly directional re-extension of both
lamellar long sides exposed perpendicularly to the direction of the electric field lines,
therefore causing each osteoblasts to become elongated perpendicularly to the direction of the
electric field polarity. Retraction was even less pronounced with field strengths lower than 7
V/cm and decreasing gradually with lower V/cm. When osteoblasts were exposed to DC ELF
strengths lower than 7 V/cm, cells underwent a gradual outward spreading of their long sides
exposed perpendicularly to the direction of the electric field lines. At the same time, cells
contracted their broad part directly exposed to the direction of the field lines, which also
caused an alignment or orientation perpendicular to the direction of the field lines of each
osteoblast. All cellular responses adopted by the cells always seemed to be proportional to
time; they decreased in speed when voltage decreased. No difference was found according to
age of the cell.
Osteoblasts showed different behavior depending on DC ELF strengths; the cellular
responses (retraction, retraction-elongation, and orientation or alignment) were classified into
3 different field strength ranges and are summarized in table 1. The next sections describe
every cellular response in detail.
54
Range (Volts/cm)
>12
7-12
<7
Cellular Response
Retraction
Retraction and Elongation
Alignment, Orientation
Table 1. Cellular responses under different DC electric field strengths.
3.101. Retraction
The first experimental conditions involved exposing osteoblasts to the highest DC
ELF strength. The first response observed in osteoblasts was a clear and pronounced
retraction (figure 20). When cells were subjected to ranges greater than 12 V/cm, they started
to retract their lamellar extensions that were exposed directly to the electric field lines
(cathode and anode). Subsequently cells underwent a total retraction of the whole
cytoskeleton in response to the applied field strength. The retraction phase did not begin
immediately in the cells. It became more pronounced with longer exposure time (table 2).
Furthermore, retraction seemed to be always proportional to time and it increased in speed
when voltage increased (figure 21). In other words, with greater electric fields, cells
underwent faster retraction. For example, with 15 V/cm, cells started to respond with
retraction after 5 minutes of field exposure, with 20 V/cm, after only 2 minutes.
Strength
12 V/cm
15 V/cm
18 V/cm
20 V/cm
First visible
retraction in
some cells
9 min
5 min
3 min
2 min
Maximal time of
Clear
retraction in all exposure to the electric
cells
field
40 min
120 min
25 min
70 min
10 min
40 min
5 min
15 min
No. of
Experiments
10
30
7
7
Table 2. Correlation Time-Field strength in cells that underwent only retraction when subjected to DC
ELFs. First column shows the field strength applied. The second column shows the time when cells started to
retract. The third column shows the time for a visible retraction in all cells. The fourth column shows the
maximal time of exposure for the cells. Last column shows the number of experiments that were made at every
field strength.
A
B
C
Figure 20. Cells underwent retraction when they were subjected to 15 V/cm ELFs. Pictures show the
same cells before (A), during (B), and after (C) electric field application. Image B was taken after 30 min of
exposure to the field. Image C shows the cells 90 min after the field was removed. Cells underwent a clear
retraction at the sides exposed directly to the field lines. The arrows indicate the direction of the retraction. The
signs indicate the orientation of the electric field polarity. Images are 80 µm large.
55
Figure 21. Retraction increased in speed when voltage increased. Graphic depicts results from
experiments with 12, 15, 18 and 20 V/cm
The behavior of microfilament architecture was evaluated under DC ELF exposure.
Microfilaments are linear assemblages of the protein actin and are better known as actin
filaments. Actin filaments group to form stress fibers, which are responsible for maintaining
the structure and shape in the cells (figure 22A).
To evaluate the role of actin cytoskeleton during the retraction phase, cells were
transfected with GFP-h1 CaP as indicated in material and methods (section 2.105c), and then
subjected to DC ELFs as above mentioned. Cells underwent a shortening of their stress fibers
after field application (7 different experiments). The stress fibers that shortened were those
exposed directly to the electric field lines (figure 22). The beginning of stress fiber shortening
was correlated with the retraction visualized in cells with phase contrast microscopy. These
observations showed that the pronounced retraction of the cells under DC ELFs of 12 V/cm or
greater magnitude was carried out by the shortening of stress fibers at those sides directly
exposed to the electric field lines (see figure 20 and 22), which caused a visible retraction of
the cells in those sides.
A
B
C
Figure. 22. Cell transfected with GFP-h1 CaP were subjected to 15 V/cm. The same cell in rest (A),
under 10 min (B) and under 30 min (C) of field exposure. Cell retracted their stress fibers directly exposed to the
electric field lines. Every images is 25 µm large.
The recuperation (re-spreading) of the cells after their retraction depended on time of
exposure to the field. When the electric field was turned off cells were left to rest in the
electro-chamber for a minimum period of 120 min. Cells began to re-spread their retracted
sides until they recovered a normal shape (figure 20C). The re-spreading began to be visible
during the first 20 min, which was taken as a recuperation signal. The coverslip, where the
cells were attached was removed from the chamber after 120 min and put into a normal
culture plate with growth medium for 24 hours. In all cases cells looked normal and they were
tested with the same DC ELF strength again. Cells responded always in the same manner
56
again. On the other hand, if the cells did not begin to re-spread during the first 20 min, they
were not removed from the electro-chamber. New medium was applied after 120 min. Cells
stayed at rest in the electro-chamber for 10-24 hours. In all cases the cells died. Recuperation
or re-spreading seemed to be always proportional to time of exposure and field strength
(figure 23). Longer time of exposure than those indicated in table 2 caused damage in the
cells and as consequent they did not recuperate. Furthermore, with longer time of exposure or
greater electric fields, cells needed a longer time for full recuperation.
Figure 23. Maximum time of exposure to the electric field. After this time cells did not re-spread.
Retraction was less pronounced when DC ELF strengths were decreased. In 3 out of 7
experiments with 12 V/cm DC ELFs, some cells underwent an elongation following the
retraction phase. In the remaining 4 experiments cells underwent only retraction. With DC
ELF intensities of 15 V/cm or greater, cells never underwent this elongation phase. However,
with DC ELFs of 11 V/cm, in all experiments cells underwent an elongation. This suggested
12 V/cm was the threshold between the retraction and retraction-elongation phases (table 1).
The process retraction-elongation is described in detail in the next section.
3.102. Retraction and Elongation
DC ELFs of 12 V/cm caused some cells to become elongated after the retraction
phase. This observation prompted that subsequent field strengths were gradually decreased to
evaluate their effect on the behavior of the cells. When osteoblasts were subjected to DC
ELFs of 7 to 12 V/cm, they aligned and elongated perpendicularly to the direction of the
electric field lines, which caused a pronounced orientation perpendicular to the electric field
polarity. This realignment process was carried out gradually. Cells started to retract their
lamellar extensions directly exposed to the electric field lines after 5 (+/- 1) min of field
exposure. No retraction or morphological changes were visualized prior to this time. The
retraction only became evident after 25-30 min, and following the retraction phase, cells
began a highly directional re-extension of both their lamellar long sides exposed
perpendicularly to the direction of the electric field polarity, which caused each osteoblast to
become perpendicularly elongated to the direction of the electric field lines (figure 24).
Elongation was more evident and pronounced when the cells were subjected to lower field
strengths and greater time of exposure (table 3, figure 25). Like retraction phase, the
elongation phase seemed always to be proportional to the field strength; it occurred faster
with higher DC ELFs (figure 25). As previously mentioned, cells did not exhibit any
elongation with electric fields greater than 12 V/cm; they only had retraction. The elongation
57
phase was only possible with DC ELFs lower than 12 V/cm. In the elongation phase, cells
remained aligned with an active elongation at each end as long as the electric field continued.
After the electric field was turned off, cells began to spread their retracted sides and the loss
of the orientation was evident.
A
B
C
Figure 24. Cells underwent retraction-elongation when subjected to 11 V/cm DC ELFs. Picture A
shows the cells at rest. Picture B and C show the same cells after 40 min (retraction phase) and 90 min
(elongation phase) of field exposure respectively. Cells underwent first a retraction of the sides exposed directly
to the field lines (arrows in B). Following the retraction, cells elongated perpendicularly to the direction of field
polarity (arrows in C). Signs show the electric field polarity. Images are 80 µm large.
Range
11 V/cm
10 V/cm
9 V/cm
7 V/cm
Time for a clear
elongation in all cells
120 minutes
150 minutes
180 minutes
200 minutes
Maximal time of exposure
to the field
200 minutes
240 minutes
300 minutes
360 minutes
No. of Experiments
10
30
8
8
Table 3. Correlation Time-Field strength in cells that underwent retraction-elongation when subjected to
different DC ELFs. First column shows the field strength applied. The second column indicates the time for a
clear alignment in all cells. The third column shows the maximum time of exposure for the cells. Last column
shows the number of experiments that were made at each field strength.
Figure 25. Cellular alignment decreased in speed when voltage decreased.
Visualization of actin filaments with GFP-h1 CaP in elongated cells showed that all
stress fibers exposed directly to the direction of electric field polarity were disrupted, and only
those exposed perpendicularly to the field lines remained, elongated, and aligned resulting in
an alignment perpendicular of the stress fibers to the direction of the field polarity (figure 26).
58
These observations suggested that stress fibers at the sides exposed directly to the field lines
were completely depolymerized after their shortening (see figure 22). In other words, DC
ELFs between 7-12 V/cm caused that cells lost their stress fibers exposed directly to the field
lines, which in turn generated an elongated shape in the cells (5 different experiments).
A
B
Figure 26. Stress fibers visualized in an elongated cell. Picture A shows an aligned cell with phase
contrast microscopy. Picture B shows the same cell transfected with GFP-h1 CaP. All visible stress fibers
exposed directly to the electric field lines were disrupted. Picture was taken after 150 min of electric field
exposure. Field strength was 10 V/cm. Both images are 15 µm large.
Re-spreading of the cells after the elongation phase was proportional to time of
exposure and field strength (figure 27). The process of evaluating the re-spreading of the cells
was the same as that undertaken with retraction. Cells were kept at rest for 120 min after the
field was turned off. If cells started to re-spread during the first 20 min, after 120 min they
were removed from the electro-chamber and put into a normal culture plate with growth
medium. The next day cells were stimulated again. In all cases cells responded to the electric
field in an identical manner. However, if the cells did not start to re-spread during the first 20
min, they were not removed of the electro-chamber and new medium was applied. Cells were
kept at rest, but in all cases cells died.
Figure 27. Maximum time of exposure to different electric field strengths for cells that underwent the
retraction-elongation process. After this time cells died.
59
In these experiments it was very clear that cellular retraction was less evident when the
field strength decreased. Retraction in cells appeared later if the DC ELF strengths decreased.
This made difficult the identification of retraction phase in cells exposed to DC ELFs of 7
V/cm or lower intensity, as cells started to orientate themselves at the same time. Under these
field strengths, cells underwent only a light retraction of their sides directly exposed to the
field lines, and at the same time they underwent an outward spreading of their long sides
perpendicular to the electric field lines. This cellular response was classified as the alignment
or orientation phase (table1). This process is described in detail in the next section.
3.103. Alignment and Orientation
Retraction phase was less pronounced and evident in accordance with decreases of DC
ELF intensities. When osteoblasts were subjected to DC ELFs lower than 7 V/cm, they
underwent an alignment perpendicular to the direction of the electric field lines, but without
an evident retraction. Cells underwent an outward spreading of their long sides exposed
perpendicularly to the direction of the electric field lines, and at the same time they contracted
their broad part exposed directly to the electric field lines (figure 28). This contractionspreading process was observed as a light movement in the cells. This cellular response could
be assumed to be the same as that seen in the retraction-elongation phase, but it differed
slightly because retraction was less evident and sometimes the cells changed their position
before they orientated perpendicularly to the field lines. It may be possible to say that cells
underwent a gradual “rotation” to achieve their orientation. In the phase of retractionelongation cells never turned or changed position. They underwent an evident retraction of
their sides exposed directly to the electric field lines and then elongated perpendicularly to the
field polarity. Therefore both processes were different. Alignment or orientation process
became evident in a few cells after 60 min of stimulation and visible in all cells after 120 min.
A clear and pronounced orientation perpendicular to the field lines was evident in all cells
after 200 minutes of exposure to the field (figure 29). The orientation process seemed to be
inversely proportional to the strength and time of exposure to the field (table 4, figure 29).
Lower electric fields caused slower orientation processes.
Figure 28. Pictures demonstrate an alignment process in cells subjected to 6 V/cm for 240 minutes.
Picture A shows the cells at rest; B after 30 min of exposure; C after 60 min; D after 100 min; E after 150 min
and; F after 240 min. A clear cellular orientation to the electric field polarity (signs) was evident in all cells.
Images are 80 µm large.
60
Range
6 V/cm
5 V/cm
4 V/cm
Time for a clear
orientation in all cells
240 minutes
300 minutes
330 minutes
Maximal time of exposure
to electric fields
390 minutes
420 minutes
450 minutes
No. of Experiments
10
6
10
Table 4. Correlation Time-Field strength in cells that underwent an alignment process when subjected to
different DC ELFs. First column shows the field strength applied. The second column indicates the time for a
clear orientation in all cells. The third column shows the maximum time of exposure for the cells. Last column
shows the number of experiments that were made at each field strength.
Figure 29. Orientation process decreased in speed when voltage decreased.
In the orientation process, Cells transfected with GFP-h1 CaP showed the same results
as with the retraction-elongation phase. Cells lost all stress fibers exposed directly to the
electric field polarity, and only those perpendicular to the field lines remained, which
elongated and aligned perpendicularly to the direction of field polarity (figure 30). Stress
fibers usually traversed the entire length of the cell, which could be the reason that each cell
become perpendicularly orientated to the direction of the field lines.
A
B
Figure 30. Visualization of stress fibers in an orientated cell. Picture A shows an orientated cell with
phase contrast microscopy; B shows the same cell transfected with GFP-h1 CaP. All stress fibers exposed
directly to the field polarity were disrupted. Images were taken after 240 min of electric field exposure (6 V/cm).
Every images is 15 µm large.
61
As seen in all other cases, cells were able to remain for only a specific time under field
exposure. Their recuperation or re-spreading was dependent on the field strength and time of
exposure to the field (figure 31). When the field was turned off, cells began to spread in all
directions but more evident at the sides where they were exposed directly to the electric field
lines, exactly in front of cathode and anode. After 120 minutes cells were removed of the
chamber and put into a normal culture plate with growth medium. They were again stimulated
24 hours later. In all cases cells responded to the electric field in the same manner. If the cells
did not start to re-spread during the first 20 min, they were not removed from the electrochamber and new medium was applied. Cells were kept at rest, but in all these cases the cells
died.
Figure 31. Maximum time of exposure to different electric field strengths for cells that underwent the
orientation process. After this time, cells died.
3.104. Summary of Cellular Responses caused by DC Electric Fields
The application of a DC ELF to osteoblasts caused 3 different effects: retraction,
retraction-elongation, or alignment. The responses showed by the cells were dependent on the
field strength (table 1), and suggested to be a process of adaptation to a contractile force
caused by the application of a DC ELF. Cells first responded with a retraction of their sides
exposed directly to the direction of field polarity. It suggests that the electric field was
exerting a contractile force at both sides of the cell exposed directly to the direction of field
lines, causing thus their retraction at exactly those sides. When the force exerted was very
high (proportional to the field strength), it caused a fast and very pronounced retraction, and
as a consequence cells were not able to tolerate a long time of exposure to the field. It might
cause irreversible damages and cells could not recuperate. This hypothesis suggested that it
was harder for cells to adapt to high forces, and as a consequence cells remained in a retracted
state. On the other hand, if the force exerted was very low, then the effect on the cells was low
and slow. Thereby the process of adaptation was also progressive and slow, and as a
consequence retraction was almost not visualized. The result was therefore that cells were
able to accept a longer time of exposure to the field. However, in other case, if the force
exerted was somewhere between low and high, then the effect on the cells was not
pronounced and neither low. As a consequence, what was firstly visualized was the effect
caused by the application of the DC ELF (retraction), followed by the adaptation of the cells
to the tension exerted (elongation), which caused the retraction-elongation phase. Therefore,
the responses showed by osteoblasts under DC ELF exposure seemed to coincide with the
hypothesis of an adaptation process.
62
The DC ELF strengths where both retraction followed by an elongation occurred, and
as consequence resulting in the alignment or orientation of the cells, were found to be
between 7 and 12 V/cm. These field strengths represent the ideal values to investigate the
process of adaptation of osteoblasts to DC ELFs. Therefore all of the following experiments
were carried out in the range of 7 to 12 V/cm. Normally 10 V/cm, as the middle point was the
field strength chosen.
To evaluate such hypothesis of an adaptation process, it was necessary to differentiate
the responses of the cells of secondary effects, such as temperature and changes of pH in the
medium. The second step was to evaluate if a DC ELF really was causing a contraction force
in the cells. Actually there are not still techniques available that can measure the force
distribution at the whole cell membrane. However, traction force microscopy (TFM) is a good
method to evaluate indirectly a force distribution across cells. Thus, to evaluate if a DC ELF
was exerting a contractile force, experiments with TFM were carried out. The next sections
describe the experiments conducted to measure the temperature and pH, followed by the
experiments with TFM.
3.105. Measurement of Temperature in Medium
To test if the electric field increased the temperature of the medium, a thermometer
was inserted into the chamber. Temperature was registered at both sides of the chamber
bridge over a 3 hours period (usual time for a single experiment). Temperature of the medium
was never higher than the temperature in the room; it remained 1 oC lower (36 oC). It was not
possible to insert the thermometer under the bridge where the cells were, but to confirm that
temperature under the bridge was the same, infrared measurements were conducted (figure
32). No significant change of temperature was detected. These observations showed that an
electric field did not increase the temperature of the medium and therefore the cellular
responses could be accounted for the electric field only.
3.106. Measurements of pH in Medium, Electrophoresis and Electro-osmosis
An electric field should redistribute charged macromolecules that are free to move
laterally in the plasma membrane and extracellular medium. Thus negatively charged
macromolecules tend to move to the positive side of the cell and positively charged
macromolecules to the negative side, causing the electrophoresis effect. Electrophoresis in
cells was demonstrated experimentally in the seventies by Poo and Jaffe (101, 120). Since cell
membrane surfaces have a net negative charge in account their lipids (see section 1.302), at
the same time that electrophoresis occurs, positive particles bound to the surface of cell
membrane will move when the electric field is applied, resulting in an electro-osmotic flow of
fluid towards the cathode side of the membrane and parallel to the surface to the cell (100,
102). Therefore the movement of the charged macromolecules is under the influence of both
the electro-osmotic fluid drag and electrophoretic forces on its own charge.
If the application of a DC ELF causes a redistribution of particles in the medium, then
the pH should change at both sides of the cells, in this case, on both sides of the bridge. To
evaluate if the pH changed with the application of a DC ELF, a pH electrode was inserted into
both sides of the bridge during DC ELF exposure. It was very evident that the pH changed
and its change was dependent on field strength. A relation of 0.15 ± 0.02 pH units per volt
was found (table 5). Whereas pH on the negative side was decreased, pH on the positive side
was increased (figure 33). After the electric field was removed, medium recovered its normal
pH immediately (in about 20 seconds, but this delay could be due to the speed of the
63
equipment). The pH measurements were conducted at 36 oC and at room temperature (20 oC),
and no significant change was visualized. These observations showed that an electric field
produced a redistribution of the particles in the medium causing changes in the pH.
Unfortunately the pH under the bridge could not be measured, but if there was a proportional
change on both sides of the bridge, in the middle point under the bridge, the pH should be
constant.
Figure 32. Infrared measurements of temperature under the bridge in the electro-chamber.
Measurement was carried out at 15 V/cm. Picture A shows the temperature without electric field. No change was
visualized after 2 (B), 15 (C) and 30 (D) minutes of field application.
Volts/cm ± 5 mV
0
1
3
5
7
9
11
13
15
17
pH positive side
7,2
7,49
7,8
8,1
8,41
8,74
9,06
9,38
9,75
10,07
pH negative side
7,2
6,88
6,57
6,25
5,92
5,59
5,26
4,92
4,56
4,22
Total/2
7,2
7,18
7,18
7,17
7,16
7,16
7,16
7,15
7,15
7,14
Table 5. pH units on both sides of the bridge were dependent on the field strength. Whereas pH on the
anode side increased with voltage increased, pH on the cathode side decreased. Values were found at a relation
of about 0.15 ± 0.02 per volt. Errors could be due to voltmeter and power supply precision.
64
Figure 33. Relation between pH and electric field strength.
3.107. Traction Force Microscopy
Cells in culture secrete an elaborate matrix composed of several components into an
assembled matrix known as extracellular matrix that is adsorbed to the surface on which the
cells are growing (121). Specialized regions of the cell membrane called focal adhesions
mediate adhesion of cells to the extracellular matrix. At these sites, bundles of actin filaments
(stress fibers) are anchored to transmembrane proteins called integrins through a multimolecular complex of proteins. Thereby integrins bind simultaneously to the extracellular
matrix on the outside and actin cytoskeleton inside.
Cells generate tension within contractile microfilaments (stress fibers) in their
cytoskeleton and the extracellular matrix resists the deformations caused by the changes in
contractility. Thus the force generated within the cell by the cytoskeleton is expressed outside
the cell as a traction force in the substratum.
Traction force microscopy, TFM, is a technique used for the detection of cellular
traction forces. TFM provides direct quantitative information about the detailed magnitude,
65
direction, and location of cell-substrate mechanical interactions. In the present study, TFM
was used to evaluate if the application of DC ELFs were exerting changes in cytoskeleton
contractility of osteoblasts that could be the responsible of generating cell shape changes.
Traction forces in osteoblasts under DC ELF exposure were determined by a method
described in detail in the section of material and methods. Briefly, cells were plated on
collagen-coated flexible polyacrylamide substrates with fluorescent beads (figure 34).
Substrate deformations were a result of the forces exerted by the cells during cellular
movements. The traction force measured was the change between the traction force exerted
by the cells at rest (no electric field) and the traction force exerted by the cell under DC ELF
exposure. Thus, the change in traction forces caused by the application of the field was
calculated. Figure 35 shows a traction image computed from an experiment with 10 V/cm.
The arrows give the direction of the traction vectors and their length is proportional to the
magnitude of the traction. This image was computed comparing the substrate deformation
during rest and after 30 s of DC ELF application. It was evident that the traction force
changes computed were exactly at the sides exposed directly to the field lines, where the
retraction with phase contrast and fluorescence microscopy was evident (figure 20 and 22).
Theorically the application of a DC ELF should have only a magnitude at those sides exposed
directly to the field lines (Eq. 3, section 1.401). Therefore, the increase in traction force
seemed to have a relation with the distribution of transmembrane potential caused at the
membrane by the application of a DC ELF. These observations suggest that the retraction of
the cells could possibly be caused by an increase in contractility in the cytoskeleton when
subjected to a DC ELF. In other words, a DC ELF was exerting a contractile force on both
sides of the cells.
Figure 34. Fluorescent beads embedded in a flexible polyacrylamide substrate. Bead displacements are
produced by substrate deformations that are product of the traction forces exerted by the cells (125). Image is 80
µm large.
The global traction force exerted by the whole cell during DC ELF exposure can also
be determined using the software developed by Dembo (125). Figure 36 shows the global
traction force computed in the experiment mentioned in figure 35. Positive forces are assigned
to increases in cell contractility (fluorescent beads move inwards in the substrate). Negative
forces are assigned to decreases in cell contractility (fluorescent beads move outwards). In
66
cells under DC ELF exposure traction forces are positive when the cells retract, and negative
when the cells spread.
Figure 35. The application of a 10 V/cm DC ELF causes an increase in traction force in osteoblasts.
Traction force vectors were calculated comparing the differences in traction force between a cell at rest and a
cell after 30 s of field application. Arrows show the direction of traction forces. Arrow length is proportional to
traction force magnitude. Scales are showed at the top in the picture.
M63elf05100166
40,0E+0
Force [N/m²]
38,0E+0
36,0E+0
34,0E+0
32,0E+0
30,0E+0
28,0E+0
0
10 V Electric field on
5
Tim e [m in]
Figure 36. Global traction force computed for a cell at rest (blue) and under 10 V/cm field exposure
(red). Traction force was calculated firstly in the cell at rest, then cell was kept at rest without perturbation for 5
min, subsequently the electric field was applied. After 30 s of field application image number 2 was computed.
Traction force magnitude is in N/m2.
To examine clearly the force distribution caused in the cells by the application of a DC
ELF, traction force magnitudes of each pixel in the cell area were represented in different
colors (144). It was observed that the highest increases in traction forces were concentrated at
the membrane zone, preferentially at the sides exposed directly to the direction of field
67
polarity (figure 37b). The lowest or no traction force changes were found in the middle of the
cell and at the sides exposed perpendicularly to the field lines (see figure 37). Interestingly,
the increase in traction force also took place at the same sides where cells underwent the
retraction, and where only a theorical perturbation would be imposed by the electric field.
Traction force vectors were computed in 24 cells subjected to 10 V/cm DC ELFs (SD=18 s).
Two cells did not show significant changes after field application; 10 cases were computed
after 30 s of field application and 14 cases after 10 s. In all 22 cells visualized, the traction
vector changes caused by application of the field showed a pattern similar to figure 37, an
increase in traction force at the anode- and cathode-facing membrane. These observations
suggested that DC ELFs were causing a contractile force in the cells up the first seconds of
application, before any morphological change could be visualized. The force imposed pulls
inward on the cell surface at both sides directly exposed to the field lines.
Figure 37. traction forces computed in a cell subjected to a 10 V/cm DC ELF. Picture A represents the
increase of traction force vectors caused by the application of the electric field. Picture B represents the traction
force magnitudes in different colors for the same cell. It was evident that the highest increase in traction force
was concentrated at the membrane zone at the sides exposed to the field lines. Magnitude in color is represented
in dyn/cm2. Arrows in picture A show the direction of traction forces. Arrow length is proportional to traction
force magnitude. Scales are showed at the top in the picture.
Analysis of the traction force vectors during an alignment process (figure 38) showed
clearly that cells underwent first an increase in traction force at the sides exposed directly to
the field lines as described previously. The contractile force pulling inward on the cell was of
about 5 to 30 % of the total traction force exerted by the cell and visualized within 10 to 30 s
of field application. The force imposed remained without change in the direction but
increased in some cellular regions in the following time. Then after about 5 minutes cells
underwent a decrease in their tractional forces. The traction vectors showing a decrease in
force were visualized preferentially at the sides exposed perpendicularly to the direction of the
field polarity (figure 38). The decrease in force was more evident with an increase in time of
exposure. All these changes in traction forces were distinguished in the cells before any
visible morphological change was observed with phase contrast microscopy. These
observations showed that the direction of the tractional forces had a correlation with the
68
morphological changes underwent by the cells. An increase in traction forces was observed
where the cell underwent the retraction. Cells never spreaded out at the sides that had an
increase in traction force. A decrease in traction forces was observed only where cells
underwent an elongation. Therefore, these observations suggested that before the cells
underwent some morphological change, they had clearly defined the layout to follow by the
orientation of their tractional forces; thus the retraction-elongation process responsible for
causing the cells to re-orientate perpendicularly to the field lines, was only the result of the
adaptation to the tension imposed.
Figure 38. Traction force distributions during an alignment process in cells subjected to 10 V/cm. After
field application cells underwent an increase in traction forces at their sides exposed directly to the field lines.
The increase in traction force was related to the retraction of the cells at those sides. Subsequently cells
underwent a decrease in traction forces at the sides exposed perpendicularly to the field lines. The decrease in
traction force was related to the elongation of the cells at those sides. Arrows show the traction vector direction
and the length is proportional to the magnitude of the traction. The scale vector represents 10 N/m2. The bar
scale represents 10 µm. Signs show the electric field polarity.
Throughout all of the conducted experiments, when cells were subjected to the field
not once did a cell show a decrease in traction force at the sides exposed directly to the field
lines; cells always had an increase. Cells under field exposure never spreaded out at the sides
exposed directly to the direction of field polarity. Thus cells spreaded out only at the sides
exposed perpendicularly to the field lines, where the theorical tension imposed should be zero
(Equation 3).
69
3.107a. Experiment Control of Traction Force Microscopy
To confirm that DC ELFs did not cause displacements in the fluorescent beads or
changes in the flexible gel and therefore mistakes in the measurements of traction forces,
coverslips with the polyacrylamide substrates and the embedded fluorescent beads were tested
without cells. The coverslips were mounted into the chamber with normal HEPES medium
and then the electric field was applied. A significant change in the polyacrylamide substrates
was never observed.
To evaluate if DC ELFs were directly responsible for maintaining the increase of
traction forces in the cells, DC ELFs were removed from the cells after 2 minutes of
application. Two minutes was chosen because at this time cells showed a clear and defined
increase in traction forces. In all cases cells underwent a decrease of their tractional forces
after the field was removed, which indicated that the presence of the field was necessary to
maintain the increase in cell contractility.
Experiments with TFM demonstrated that a DC ELF caused a contractile force on the
cells. The contractile force was visualized as an increase in the traction forces at the
membrane zones exposed directly to the field lines. The increase in force was visualized
before the retraction of the cells at those sides. Subsequently cells underwent a decrease in
traction forces at the membrane zones exposed perpendicularly to the field lines. The decrease
in force was visualized before the elongation of the cells at those sides. These results
suggested that cells were under a contractile force with a defined layout before any
morphological change took place.
Experiments with TFM also demonstrated that the contractile force was dependent on
field application, as increase in force disappeared immediately after the DC ELF was
removed. Thus the next step was to evaluate if the process of elongation was an automatic
process after cells underwent the retraction.
3.107. Alignment Was Not Programmed in the Cells
To evaluate if the elongation process in the cells was programmed after they
underwent the retraction, the electric field was turned off immediately after the cells started
with their outward spreading perpendicular to the direction of field polarity (figure 39). In all
cases (5 experiments), cells respreaded out in all directions and more evident at those sides
exposed directly to the field lines, where the retraction had taken place. Therefore, cells had to
stay under field exposure to achieve their re-orientation perpendicular to the direction of the
field lines.
3.108. Summary of the Behavior of Osteoblasts Under Electric Field Exposure
Osteoblasts responded with retraction, retraction-elongation, or alignment when they
were subjected to DC ELFs. The cellular responses were dependent on field strength and time
of exposure. TFM showed that osteoblasts under DC ELF exposure underwent an increase in
cytoskeleton contractility immediately after the application of the field. The change in
contractility, measured as an increase in the traction forces of the cells, was visualized after
10-30 s of field application, before any morphological change could be observed. The
increase in contractility took place at the membrane zones exposed directly to the cathodeand anode-facing membrane. Thus it seemed to be in connection with the retraction
underwent by the cells later at those sides. This observation suggested that only those zones
exposed directly to the field lines were under a tensional force with a direction pulling
70
inwards on the cell, which could cause the retraction of the cell at those sides. On the other
hand, cells also underwent a decrease in traction force when they were under field exposure.
The traction vectors showing a loss of force were visualized at the sides exposed
perpendicularly to the cathode- and anode-facing membrane, exactly where the cells
underwent an elongation later. Thus, cells retracted where an increase in traction force was
visualized, and elongated where a decrease in traction force took place. Therefore the
direction of the tractional forces had clearly defined the re-orientation of the cells
perpendicular to the direction of field lines, before any morphological change could be
observed.
A
B
C
Figure 39. The elongation of the cells was not programmed after the retraction phase. Normal cells (A)
were subjected to 10 V/cm. After 50 min of field application, when retraction was evident (yellow arrows in B)
and cells started with the elongation process (blue arrows in B), the DC ELF was removed. Cells re-spreaded in
all directions and principally in the retracted sides (arrows in C). Images are 40 µm large.
These findings suggested that the response of osteoblasts to a DC ELF is a process of
adaptation to a contractile force caused by the application of the field. Cells changed their
shape to adapt to the tension imposed and with it to minimize the perturbation. This
hypothesis also suggested that for cells it was harder to adapt to high forces and as a
consequence cells remained retracted. Experiments with magnitudes greater than 12 V/cm
demonstrated that cells did not elongate after retraction phase had been evident. However,
cells elongated and orientated with DC ELFs lower than 12 V/cm. The question now is: how
can a DC ELF cause an increase in cytoskeletal contractility? In other words, what is the
pathway or mechanism of interaction between osteoblasts and DC ELFs that caused the
increase in cytoskeletal contractility, and therefore the re-orientation of the cells perpendicular
to the direction of field lines? A set of biochemical experiments were undertaken to answer
this question. These experiments are described in the next section of this work.
71
3.2. BIOCHEMICAL EVALUATION OF OSTEOBLASTS UNDER ELECTRIC
FIELD EXPOSURE
Experiments with phase contrast, fluorescence, and traction force microscopy showed
that osteoblasts retracted and elongated in response to an increase in cytoskeletal contractility
caused by the application of a DC ELF. To investigate the mechanism(s) of interaction
between DC ELFs and osteoblasts that could cause the increase of contractility, as well as
their retraction, and elongation, biochemical experiments were carried out. Various questions
were opened to determine what biochemical evaluations should be conducted; these questions
and their possible answers are listed and described below:
1) Since contractile forces are directly related to stress fibers, then cells without stress fibers
should not be able to develop changes in contractility. One would expect that if the
contractile force generated by the cytoskeleton is responsible for causing the retractionelongation process, then if blocked, this process should be inhibited. To evaluate this
hypothesis, two drugs, cytochalasin B and latrunculin A were used to disrupt stress fiber
organization (actin filament polymerization) in the cells when subjected to DC ELFs.
2) Since a DC ELF is causing a contractile force at both sides of the cell, then it should
produce the same effect on both sides of the cell membrane. However the anode facing
cell membrane should be more hyperpolarized, and cathode facing cell membrane should
be more depolarized. Thus to evaluate if a DC ELF was really causing a different
polarization effect at both cell membrane sides, visualization of membrane potential
changes were carried out in osteoblasts.
3) Cellular signaling mechanisms are designed to transmit information from the cell
membrane surface to specific targets within the cell. Often the information is transmitted
by means of intracellular messengers, of which the ion calcium, Ca2+, is one of the most
important. Intracellular Ca2+ increase can result from Ca2+ release from intracellular stores
or from the opening of Ca2+ channels located on the membrane that permit Ca2+ influx in
the cells. Contraction and shape changes in cells occur through two processes amply
reported to be controlled by Ca2+ (169-172). To evaluate if Ca2+ activity was required for
the cells to respond to DC ELFs, measurements of intracellular Ca2+ were carried out.
Both possibilities, Ca2+ release and Ca2+ influx were investigated in this work.
4) Actin filaments in combination with myosin II, form the so called stress fibers, which are
responsible for generating the traction forces in the cells (138). Myosin II is activated by
the enzyme myosin light chain kinase, MLCK. To evaluate if activation of MLCK caused
the increase in traction force in the cells under ELF exposure, drugs were used to inhibit
MLCK activation.
The results found from these experiments permited an evaluation of the mechanism(s)
of interaction between DC ELFs and osteoblasts that caused the alignment of the cells
perpendicular to the field lines.
3.201. Alignment Is Dependent on Actin Polymerization
Stress fibers are responsible for generating the contractile force in the cytoskeleton.
The perpendicular elongation of cells under field exposure has also been reported to only be
dependent on stress fibers (173). To address the question that the re-orientation perpendicular
of osteoblasts to the field lines was solely dependent on the stress fibers, cells without stress
fibers should not generate cytoskeletal contractility. Cytochalasin B, CB, and latrunculin A,
72
Lat A, were used to modulate actin polymerization. Both drugs are potent inhibitors of actin
filament polymerization (154-158). CB disrupts microfilament organization and caps the
barbed end of actin filament and accordingly inhibits further elongation from the barbed end
(158). Lat A binds actin monomers and thereby inhibits actin polymerization (154). Thus if
filaments turn over, newly freed monomers will be bound by Lat A and thus be incapable of
reassembly and new filaments synthesis monomers will be incapable of assembly. Both drugs
were able to inhibit the elongation of the cells in concentrations of 2 µM CB (12 experiments)
and 100 nM Lat A (20 experiments). In all cases cells never elongated under the presence of
either CB or Lat A, but cells elongated after the drugs were washed out.
Stress fibers are the support to maintain the stability of cytoskeletal structure and cell
shape; the loss of stress fibers by application of the drugs caused cell rounding as previously
reported (156). Thereby, cell growth could be analyzed in rounded osteoblasts exposed to DC
ELFs. Cells were treated with Lat A for periods of 60-70 min until cell rounding was evident
(figure 40A). Lat A was washed out and normal HEPES medium was applied. The washing
also caused detachment of some cells from the substrate under the bridge by suction through a
pipette; thereby a better cell rounding was observed in some cells. Cell rounding was also an
excellent model to test the effect of a DC ELF in a spherical cell (76). The theorical model
states that the direct target of the steady field on a single spherical cell is the cell membrane
since it is nonconductive (section 1.401). Thus assuming the membrane is nonconductive, or
its resistivity is much higher than those of the internal and external media, the transmembrane
potential change caused by the application of a DC ELF in a spherical cell, may be calculated
simply using equation 3 given in section 1.401.
Vind = 1.5 EappRcosθ
Where Vind is the induced transmembrane voltage, Eappl is the external DC ELF, R is
the cell radius and θ is the polar angle measured from the center of the cell with respect to a
point of interest on the cell membrane.
The equation shows that the application of a DC ELF causes a maximum and
minimum perturbation at both opposite sides of the membrane. Thus, in rounded cells the
difference between both magnitudes should be evident in the cell behavior. Figure 40 shows
the growth out of rounded osteoblasts under a 10 V/cm DC ELF. Cells only stretched out on
the sides where the theorical magnitude of the perturbation was zero, perpendicular to
direction of the electric field lines (90 degrees). A rounded cell never underwent retraction,
only elongation and was always evident in the direction perpendicular to the field lines. Cells
never stretched out along to the axis of the field lines, which suggest that the external DC ELF
caused a perturbation that inhibited the formation of new actin filaments and stress fibers at
those sides exposed directly to the direction of field lines. Interestingly those sides were
exactly the same where intact cells underwent the retraction; therefore, this perturbation
seems to be no only responsible of blocking stress fiber formation, but also to disrupt it.
73
Figure 40. Rounded osteoblasts subjected to a 10 V/cm DC electric field. Cells were treated with 100
nM Lat A to disrupt their stress fibers prior to stimulation. Stress fiber disruption was evident through cell
rounding (A). After Lat A was removed and normal medium applied, cells were exposed to the field. Registers
were taken after 20 min (B), 60 min (C), 90 min (D), 120 min (E), and after 180 min (F) of field exposure. Cells
only stretched out at the sides perpendicular to the electric field lines where the theorical tension imposed is zero.
Images are 200 µm large.
When the DC ELF was turned off, cells began to spread out in all directions but more
evident at the sides that were exposed directly to the direction of field polarity. This
observation suggested that the permanent application of the field was necessary to cause the
elongation of the cells at the sides exposed perpendicularly to the field lines. After the field
was turned off, cells were kept at rest for 120 minutes and then were removed from the
electro-chamber and put into a normal culture plate with growth medium. Cells were tested
the following day with a normal DC ELF stimulation. Cells responded well in all cases. These
observations showed that cells did not undergo any damage by the application of Lat A and
DC ELF.
3.202. Membrane Potential Changes
It has been demonstrated experimentally that a DC ELF can significantly alter the
transmembrane potential in the cells (98, 99). The anode-facing side of the cells became more
hyperpolarized and cathode-facing side of the cells became more depolarized. The use of
optical dyes to detect changes in the membrane potential has been very useful since changes
in specific membrane zones can be visualized. To investigate if the electric field caused
changes in the membrane potential, experiments with the dye DiBaC4 [3] were carried out (10
experiments). DiBAC4 [3] has shown to be a good indicator of membrane potential changes in
bone cells (133). Osteoblasts subjected to a DC ELF underwent a change in their membrane
potential (figure 41). The anode-facing side of the cell became more hyperpolarized which
was showed as a fluorescence decrease. The cathode-facing side of the cell became more
depolarized, here depicted as a fluorescence increase.
74
Figure 41. Membrane potential distribution in a cell subjected to 12 V/cm ELF. The anode side was
hyperpolarized (A) and the cathode side was depolarized (B). The curves at the right side represent the average
of hyperpolarization and depolarisation of the closed zones in A and B at the left side respectively (Y axis). The
DC ELF was applied for about 5 minutes. Line marked with on shows the beginning of application of the DC
ELF. Image is 40 µm large.
It was evident that the application of a DC ELF caused a different polarization effect
on both sides of the cell membrane. Since cells are not symmetrical in shape and structure, it
is difficult to set an evaluation of how big the change in voltage at every part of the cell was.
Cells are a three-dimensional structure; figure 41 represents only a two-dimension figure and
not the really cell shape. The registers visualized in the graphic in figure 41 were only the
average of the marked zones in the cells. The registers only showed that the polarization was
differently distributed in two opposite sides of the cell; anode side was hyperpolarized and
cathode side was depolarized.
3.203. Calcium Measurements
A large number of studies suggest that calcium plays a role in the control of direction,
locomotion and modulation of shape in the cells as a second messenger (171, 174, 175).
Previous reports have demonstrated that some cells respond with a cytoplasmic calcium
increase when subjected to ELFs (99, 107, 176-178). To evaluate if the electric field was
causing a calcium increase in the cytoplasm of osteoblasts, calcium in cells under field
exposure was measured using the calcium-sensitive fluorescent dye fura 2 AM. Fluorescent
images of the cells showed a cytoplasmic calcium increase when subjected to DC ELFs
(figure 42). To identify the type of calcium present in cells under field exposure, calcium
waves of single cells were evaluated. It was evident that a DC ELF produced a temporal and
sometimes oscillant calcium waves in the cells, however no spikes, sparks or any other
calcium signal (figure 43). The calcium waves stayed for several seconds or minutes in the
cells. To investigate if calcium waves were produced directly by the electric field, on-off
experiments were carried out. Experiments with 10- (figure 44A), 5- (figure 44B) and 1(figure 44C) min on-off were evaluated. It was evident that calcium was present only in on
periods and it disappeared during off periods. This observation suggested that the permanent
application of a DC ELF was responsible for generating the cytoplasmic calcium increase and
maintaining it for several seconds or minutes in the cells.
75
Figure 42. Cytoplasmic calcium increases in osteoblasts under DC ELF exposure. The fluorescence
changes are related to changes of intracellular calcium concentration in the cytoplasm. Changes in color are
derived from an increase in the intracellular calcium concentration of the cells. Therefore colors are proportional
to the calcium concentration. Blue color is the rest level. Brightest color represents the maximal calcium
concentration reached by the cells. Images are 60 µm large. 10 V/cm were applied.
Figure 43. The application of a 10 V/cm DC ELF caused temporal and sometimes oscillant calcium
waves in osteoblasts. Graphics show the calcium response of single cells, which are shown in the black picture.
Every script represents a single cell. Colors in graphics were used simply to differentiate each calcium response.
Ratio (Y-axis) represents the changes in calcium concentration (for details see section 2.105a); X-axis represents
the time (seconds). Line marked with on shows the beginning of application of the DC ELF. Image with cells is
90 µm large.
76
Figure 44. Visualization of calcium activity in cells under 15 V/cm on-off DC ELFs. Cells responded
with calcium increase only during on periods. Graphics represent different experiments: in experiment 1 on-off
periods were 10 min (A), in experiment 2 on-off periods were 5 min (B), and in experiment 3 on-off periods
were 1 min (C). Lines separate on of off periods. First line is always on activation. Y-axis represents the changes
in calcium concentration (Ratio); X-axis represents the time (seconds).
3.203a. Characterization of Calcium Response (Peak Value)
It was evident that cells under DC ELF exposure responded with a cytoplasmic
calcium increase. However cells did not respond immediately after field application, cells had
a delay between beginning of stimulation and the calcium increase (see figure 43). Since cells
responded sometimes with multiple oscillations, the characterization of the calcium response
was carried out only analyzing the first calcium wave under a 10 V/cm DC ELF. It was
measured at the time when calcium reached its peak value (maximal calcium concentration,
figure 45), which is expressed in values of ratio percentage (see methods, section 2.105a).
Table 6 and figure 46 show an analysis of time and ratio for 22 cells. Cells were analyzed in 5
different experiments and taken from different cell cultures that were 1-4 days old. The delay
average between beginning of stimulation and calcium peak value was measured with an
average of 84.59 s; peak value (ratio) was measured with an increase average of 31.3 %. It
means cells had a delay of 84.59 s to reach an increase of 31.3% of their basal calcium
concentration. This delay and ratio percentage were taken like basis for the next experiments.
Figure 45. Delay between the maximal calcium increase and the beginning of stimulation. Graphics
show 2 single cells subjected to a 10 V/cm DC ELF. First line represents the ELF activation; second line
represents when peak value was reached. Delay between both lines was measured for analysis. Y-axis represents
the changes in calcium concentration (Ratio); X-axis represents the time (seconds).
77
Cell
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
Time Activation (Seconds)
22,6
35,97
34,02
16,95
47,37
24,54
41,61
43,17
34
84,3
121,51
66,97
84,33
73,61
95,4
92,71
37,22
227,98
223,18
154,83
150,08
148,72
Ratio increase (%)
36
24
22
70
14
44
30
40
75
17
16
35
15
18
62
21
18
23
57
6
15
31
Table 6. 22 cells were analyzed under 10 V/cm DC ELFs. Analysis was carried out measuring the delay
between the beginning of stimulation and maximum calcium increase, which was indicated as a ratio percentage.
Since not all cells responded at the same time with a calcium increase (see graphics),
calcium responses were analyzed in steps of 50 seconds. Thus it was identified the number of
cells that reached their peak value in the first 50 seconds, in 100 seconds, etc. It was
compared the Peak value in cell number percentage (figure 47); This mean that was compared
the percentage of cells that reached their maximum calcium concentration in steps of 50
seconds. It was evident that most cells (45.4%) had their maximal calcium increase in the first
50 seconds and then cell number percentage was decreasing as time went by.
In summary, the application of a DC ELF caused an increase of Ca2+ in osteoblasts.
Most cells responded in the first 50 seconds of field application. Intracellular Ca2+ increase
could result from Ca2+ release from intracellular stores or from the opening of Ca2+ channels
located on the membrane that permitted Ca2+ influx into the cells. To evaluate the pathway
responsible for causing an increase of Ca2+, experiments with manganese and thapsigargin
were carried out.
78
Figure 46. Analysis of Ca2+ response in 22 single cells. First graphic shows the delay between beginning
of stimulation and maximum calcium increase in every cell. Second graphic shows the ratio increase in every
cell.
Figure 47. Percentage of cells responding with an increase of Ca2+ in steps of 50 seconds. Most cells
responded in the first 50 seconds. 10 V/cm were applied.
79
3.203b. Manganese
To evaluate if the Ca2+ visualized in the cells was dependent on the calcium ions
transported across the membrane, manganese (Mn2+) was used as a substitute for calcium in
the extracellular medium. Mn2+ ions may use the same Ca2+ entry pathways and thus act as an
indicator for the activity of calcium influx into osteoblasts (147). The affinity of Mn2+ for fura
2 is 42 times greater than the affinity of Ca2+ for fura 2 (131). Mn2+ quenches the fura
fluorescence causing a fluorescence decrease. Thus a loss of fluorescence directly indicates
the permeation of the ion. Figure 48 shows an experiment with Mn2+. Cells were loaded Fura
2 like normal calcium experiment; then 10 min prior to stimulation cells were incubated with
2 mM Mn2+ and subsequently the electric field was applied. Fluorescence decrease of fura 2
was evident when the cells were subjected to DC ELFs (figure 48). Therefore, these results
indicated that Mn2+ ions flowed into the cell and quenched fura 2 causing the fluorescence
decrease. Since Mn2+ ions use the same Ca2+ entry pathways and besides are not present in
intact cells, these results showed that the application of a DC ELF can induce a calcium influx
into the cells.
Figure 48. Manganese influx into the cells quenches fura 2. Quenching is visualized as a loss of
fluorescence intensity. First line indicates beginning of stimulation. Second line indicates the beginning of
fluorescence decrease. 10 V/cm were applied. Y-axis represents the changes in fluorescence intensity; X-axis
represents the time (seconds). Fluorescence traces were observed with an excitation wavelength of 360 nm.
The influx of Mn2+ into osteoblasts did not occur immediately after field application.
To compare the delay of Mn2+ influx and the normal Ca2+ response of the cells, 19 cells in 5
different experiments with Mn2+ were evaluated (table 7, figure 49). Delay was examined by
evaluating when fura 2 started to decrease its fluorescence, which indicated the presence of
Mn2+ (figure 46). Delay average was 88.89 seconds, almost at the same time like that seen
previously with normal calcium response.
Cell
1
2
3
4
5
6
7
8
9
10
11
12
13
Influx Time (seconds)
62,99
116,99
51,99
68,99
66,99
56,99
86,99
44,99
221,01
50,99
111,02
101,99
41,99
80
14
15
16
17
18
19
82,99
38
19
204
128
133
Table 7. Delay of Mn2+ influx into the cells. 10 V/cm were applied.
Figure 49. Analysis of Mn2+ influx in 19 single cells. Graphic shows the delay between beginning of
stimulation and quenching of fura 2. Cells were stimulated with 10V/cm.
Since quenching of fura 2 was not visualized at the same time in osteoblasts, Mn2+
influx was also analyzed in steps of 50 seconds (figure 50). The highest percentage of cells
fell now in cells responding slower than 50 seconds (see figure 49), which may indicate that
the most calcium influx into the cells was after 50 seconds of field application and not in the
first 50 seconds. To confirm such observation, it was blocked the intracellular calcium release
with thapsigargin; thus only extracellular calcium should be present in cells. Therefore all
calcium increase should be provided by a calcium influx into the cells. Experiments with
thapsigargin are described in the next section.
Figure 50. Percentage of osteoblasts showing a Mn2+ influx when subjected to 10 V/cm.
81
3.203c. Thapsigargin
In the control group, the largest cell population responds with an increase of Ca2+ in
the first 50 seconds of field application (figure 47). However the largest cell population
responding with fura quenching (Mn2+) was after 50 seconds of field application. The delay
average for both is almost at the same time. This means that two different cell populations
may be present in osteoblasts responding with an increase of Ca2+ when subjected to DC
ELFs. The first population with a Ca2+ response generated intracellularly; the second
population generated by a Ca2+ influx from the extracellular medium. Intracellular calcium is
released from intracellular stores by opening of a Ca2+ -permeable channel in the endoplasmic
reticulum in response to the second messenger inositol [1,4,5] triphosphate [IP3] (145, 179).
To evaluate if the increase of cytoplasmic calcium showed by the first cell population was
dependent on the calcium release from intracellular stores (endoplasmic reticulum), cells were
treated with thapsigargin after fura 2 incubation and subjected to the electric field.
Thapsigargin causes a release of calcium into the cytoplasm, mainly through blocking the
Ca2+-ATPase responsible for pumping calcium back into the endoplasmic reticulum and thus
impeding the re-absorption of calcium (122). Therefore depletion of the Ca2+ content of the
endoplasmic reticulum is carried out by an endogenous Ca2+ leak (145, 146). Practically all
calcium is released from the endoplasmic reticulum within 30 min after thapsigargin
treatment, and thus the IP3 sensitive pools are not available for release when IP3 is raised.
Osteoblasts were treated with 5 µM thapsigargin for 30-40 min after fura 2 incubation
and prior to field application. Cells did not respond with Ca2+ increase when they were
subjected to the field. Only 6 of 35 cells responded with an increase of Ca2+ (5 different
experiments). In these 6 cells, Ca2+ increase was not visualized in the first 50 seconds of
exposure to the field. The fastest calcium response in cells treated with thapsigargin was 86,8
seconds (table 8, figure 51). These results suggested that the increase of Ca2+ visualized in the
first cell population could be caused by Ca2+ release of intracellular stores. However, Ca2+
influx could be also dependent on calcium release. This last observation was taken as only 6
of 35 cells were observed with an increase of Ca2+.
Figure 51. Calcium responses in osteoblasts under thapsigargin incubation. Cells were incubated for 40
min with 5 µM thapsigargin after fura 2 incubation; then a 10 V/cm DC ELF was applied. Only 6 of 35 cells
responded with an increase of calcium. First line in the picture shows the beginning of stimulation. Second line
shows when the peak value was reached. Y-axis represents the changes in calcium concentration (Ratio); X-axis
represents the time (seconds).
Cell
1
2
3
4
5
6
Time Activation (Seconds)
143,58
144,71
108
278,4
86,8
104,8
Fluorescence increase (%)
24
15
10
14
19
11
Table 8. Calcium increases in 6 of 35 single osteoblasts under 5 µM thapsigargin. 10 V/cm were
applied.
82
3.203d. Summary of Calcium Response in Osteoblasts
Osteoblasts responded with an increase of Ca2+ when they were subjected to DC ELFs.
However the increase of Ca2+ was not immediately visualized after field application, calcium
increased progressively until reaching its maximal concentration or peak value. The delay
between beginning of stimulation and calcium peak value was measured with an average of
84.59 s; peak value (ratio) was measured with an increase average of 31.3 %. Analysis of Ca2+
increases in single cells, as well as experiments with manganese and thapsigargin showed that
Ca2+ activity was a contribution of both, Ca2+ release from intracellular stores and Ca2+ influx
from the opening of Ca2+ channels located on the membrane. Analysis in cell number
percentage showed that about 50% of osteoblasts responded in the first 50 s of field
application, and the other 50% was distributed between 50 to 250 s. Osteoblasts that
responded with an increase of Ca2+ in the first 50 s were called group one and named “cells
with fast response”. Osteoblasts that responded with an increase of Ca2+ after 50 s were called
group two and named “cells with slow response”. Manganese experiments showed that Ca2+
influx from extracellular medium was carried out principally after 50 s of field application.
Thapsigargin experiments showed that Ca2+ increases in cells with “fast response” could be
caused by the calcium release from intracellular stores. Furthermore calcium release could
contribute to Ca2+ influx across cell membrane.
To address the question of how a DC ELF can activate calcium release from
intracellular stores and how it can stimulate calcium influx across the membrane, the first
evaluation was if the integrity of the cytoskeleton could play a role that caused Ca2+ release
from intracellular stores. This question arose since an increase in contractility in the cells was
measured with traction force microscopy during the first few seconds of field application; and
sine an increase of Ca2+ was visualized in cells after an average of 85 s. Therefore, it was
investigated if the contractility could play a role in causing the increase of Ca2+ in osteoblasts.
Ca2+ responses were monitored in cells that were not able to increase their contractility, in
cells without stress fibers. On the other hand, to evaluate calcium influx across the membrane,
a set of experiments with calcium blockers were undertaken. Cadmium, Cd2+, Lanthanum,
La3+, Nifedipine, Nif, and Nitrendipine, Nit, were used as calcium channel blockers at the cell
membrane.
83
3.203e. Calcium and Latrunculin
Actin filaments have been shown to play a fundamental role in cell contraction in
muscle and nonmuscle cells (180). Furthermore in combination with myosin II, actin
filaments are responsible for generating the traction forces in cells. Traction force microscopy
showed that the application of a DC ELF caused an increase in traction forces in osteoblasts.
Since this increase in contractility was visualized in the cells before an increase of Ca2+ in
cytoplasm, and as actin filaments are also responsible for generating the traction forces, it was
investigated if the integrity of the cytoskeleton could play a role in causing the increase of
Ca2+ in osteoblasts. Therefore Ca2+ activity was monitored in rounded cells (cells with
depolymerized actin filaments). After fura 2 incubation cells were incubated with Lat A. Cells
were kept at rest until they lost their actin filaments (about 70 min and visualized as cell
rounding). The medium with Lat A was not removed and then the DC ELF was applied.
Calcium responses were analyzed like previous experiments (figure 52). Only 11 of 28
rounded osteoblasts responded with Ca2+ increase when they were subjected to the field (4
different experiments). Osteoblasts also showed a delay between stimulation and calcium
responses. The average time to reach the calcium peak value increased to 189.3 s (table 9,
figure 53); the fastest delay was found at 74 s. These observations suggested that the integrity
of the cytoskeleton played a role in producing the increase of Ca2+ in osteoblasts in the first 50
s of field exposure, but it was not able to block the increase of Ca2+ after 50 s. It also
suggested that the increase of contractility could be responsible for causing Ca2+ release of
intracellular stores in osteoblasts subjected to DC ELFs.
Figure 52. Ca2+ increase in cells with depolymerized actin filaments and subjected to 10V/cm. The first
picture (black) shows the cells at rest and after 70 min of 100 nM Lat A incubation. Loss of actin filaments was
evident as cell rounding. Graphics show Ca2+ increases in every rounded cell (noted by scripts). Colors in
graphics were used simply to differentiate each calcium response. Y-axis represents the changes in calcium
concentration (Ratio); X-axis represents the time (seconds). Line marked with on shows the beginning of
application of the DC ELF. Images with cells are 90 µm large.
84
Cell
1
2
3
4
5
6
7
8
9
10
11
Time activation
85,81
84,33
127,23
264,82
267,76
186,41
215
110
74
75
592
Table 9. Time required for a Ca2+ response in cells incubated with 100 nM Lat A and subjected to 10
V/cm.
Figure 53. Analysis of Ca2+ response in 11 of 28 single rounded osteoblasts subjected to 10 V/cm.
Rounded cells that responded with an increase of cytoplasmic Ca2+ when subjected to
DC ELFs were also analyzed in steps of 50 s. It was identified the percentage of cells that
reached their maximum calcium concentration in steps of 50 seconds. It was evident that
osteoblasts did not undergo an increase of Ca2+ in the first 50 s of field exposure (figure 54).
These results could indicate that the integrity of the cytoskeleton was necessary to cause the
increase of Ca2+ in the first 50 s.
Figure 54. Percentage of rounded cells that responded with an increase of Ca2+ when subjected to 10
V/cm.
85
The Ca2+-increase delay in cells with depolymerized actin filaments fell in the second
cell population, after 50 s of field exposure. It indicated that the integrity of the cytoskeleton
was necessary to cause Ca2+ release from intracellular stores. However calcium influx from
the extracellular medium in osteoblasts was not dependent on the integrity of the
cytoskeleton. Interestingly the total percentage of cells responding with an increase of Ca2+
was reduced fewer than 50 % (only 11 of 28 cells). These results seemed to coincide with
those of the thapsigargin experiments, where only 6 of 36 cells responded with an increase of
Ca2+, and besides it was also after 50 s. Taken together, these observations indicated that Ca2+
release was necessary to cause Ca2+ influx too.
3.203f. Cadmium
Extracellular calcium influx across the membrane can be activated by different
mechanisms (121, 122, 145, 146, 149, 172): voltage-operated channels, receptor-operated
channels, mechanically-operated channels, and the store-operated channels, which are opened
following the depletion of internal calcium stores. The results obtained previously indicate
that Ca2+ is first released from intracellular stores and then extracellular calcium enters in the
cells. The use of drugs for blocking the Ca2+ influx should reduce the second cell population,
cells with a slow response. Sets of experiments with different Ca2+ blockers were carried out
to confirm such hypothesis. The first blocker used was cadmium, Cd2+. It is a divalent cation
that is known to be a potent calcium channel blocker (149). Cd2+ enters cells primarily by
passing through all classes of voltage-operated calcium channels and thereby inhibits influx of
calcium (150). It has been reported that Cd2+ in low concentrations (< 10µM) inhibit only L
and N type voltage-dependent calcium channel, and in higher doses block all Ca2+ channels
(151). To evaluate if calcium ions passed across voltage-operated calcium channels in
osteoblasts subjected to DC ELFs, cells were incubated with different Cd2+ concentration after
fura 2 loading and subjected to stimulation.
3.203fa. 10 µM Cadmium
The effect of Cd2+ in the calcium responses of osteoblasts was firstly evaluated with
the minimum concentration (10 µM). This concentration has been reported to inhibit only L
and N type calcium channels in cells (151). 23 cells were evaluated in 4 different experiments
(table 10, figure 55). Osteoblasts also showed a delay between beginning of stimulation and
calcium response. The delay between beginning of stimulation and calcium peak value was of
81.97 s in average. Calcium concentration reached a value of 30.1% (ratio) in average of its
rest concentration.
Cell
1
2
3
4
5
6
7
8
9
10
11
12
Time Activation (seconds)
99,63
159,28
31,46
44,33
64,57
34,22
152,84
155,94
167,93
149,46
136,5
43,26
Ratio increase (%)
17
20
25
15
15
6
28
29
35
43
35
19
86
13
14
15
16
17
18
19
20
21
22
23
46,02
28,5
69,97
33,15
47,86
15,87
19,57
18,35
23,31
23,31
320
20
66
34
69
59
41
8
20
23
25
40
Table 10. Time required for Ca2+ response in 23 single cells incubated with 10 µM Cd2+ and subjected
to 10 V/cm. Delay was measured between beginning of stimulation and maximum ratio increase.
Figure 55. Analysis of Ca2+ responses in 23 single cells incubated with 10 µM Cd2+ and subjected to 10
V/cm. First graphic shows the delay in cells to reach their calcium peak value after beginning of stimulation.
Second graphic shows the ratio increase per cell.
Since calcium responses were evident in osteoblasts incubated with 10 µM Cd2+ and
subjected to DC ELFs, cell number percentage was also analyzed in steps of 50 s (figure 56).
Most cells (56.5%) reached the calcium peak value during the first 50 s of field exposure.
87
Figure 56. Analysis of percentage of cells incubated with 10 µM Cd2+ that responded with an increase
of Ca when subjected to 10 V/cm.
2+
The percentage of osteoblasts incubated with 10 µM Cd2+ that underwent an increase
of Ca in the first 50 s increased if compared with control experiments (characterization of
calcium response, section 3.203a). The percentage of cells increased of 45.4 to 56.5 %. Since
the number of cells responding in the first 50 s was higher, this increase therefore produced a
shortening in the delay between beginning of stimulation and calcium peak value. This
observation could also indicate that 10 µM Cd2+ had an effect only in cells that responded after
50 s.
2+
3.203fb. 25 µM Cadmium
It was evident that 10µM Cd2+ did not block all calcium influx into the cells. Thus
Cd2+ concentration was increased to 25 µM. During this stage 3 different experiments were
carried out and 13 cells were evaluated (table 11, figure 57). The delay between beginning of
stimulation and the peak value was found with an average of 57.7 s. This delay shortened
when compared with the delay of cells incubated with 10 µM Cd2+. Ratio increased with an
average of 28.1% in the cells.
Cell
1
2
3
4
5
6
7
8
9
10
11
12
13
Time Activation (seconds)
87,43
24,78
36,7
24,83
86,35
30,22
155,34
127,76
49,49
37,13
13,13
38,05
38,94
Ratio increase (%)
17
11
15
34
20
11
26
18
37
45
39
57
37
Table 11. Time required for Ca2+ response in cells incubated with 25 µM Cd2+ and subjected to 10
V/cm. Delay was measured between beginning of stimulation and maximum ratio increase.
88
Figure 57. Analysis of Ca2+ responses in 13 single cells incubated with 25 µM Cd2+ and subjected to 10
V/cm. First graphic shows the delay in cells to reach their calcium peak value after beginning of stimulation.
Second graphic shows the ratio increase per cell.
The number of osteoblasts incubated with 25 µM Cd2+ and responding with an
increase of Ca2+ was also evaluated in steps of 50 s (figure 58). Most cells (71.4%) reached
the calcium peak value in the first 50 s of field exposure.
Figure 58. Analysis of percentage of cells incubated with 25 µM Cd2+ that responded with an increase
of Ca when subjected to 10 V/cm.
2+
89
The percentage of osteoblasts incubated with 25 µM Cd2+ that underwent an increase
of Ca in the first 50 s increased from 56.5 to 71.4 % when compared with those incubated
with 10 µM Cd2+. This increase produced a shortening in the delay between beginning of
stimulation and calcium peak value (57.7 s), as the number of cells responding in the first 50 s
was much higher. This observation could also indicate that 25 µM Cd2+ had an effect only in
cells that responded after 50 s.
2+
3.203fc. 50 µM Cadmium
To evaluate if higher Cd2+ concentrations could block calcium influx into the cells,
and if it could reduce even more the percentage of cells responding slower than 50 seconds,
Cd2+ concentration was increased to 50 µM Cd2+. During this stage 3 different experiments
were also carried out and 11 cells were evaluated (table 12, figure 59). The delay between
beginning of stimulation and the peak value was found with an average of 36.47 s. This delay
shortened when compared with the delay of cells incubated with 25 µM Cd2+. Ratio increased
with an average of 19.36% in the cells.
Cell
1
2
3
4
5
6
7
8
9
10
11
Time Activation (seconds)
54,12
69,75
28,97
19,43
15,11
21,58
42,09
32,38
12,95
43,22
61,61
Fluorescence increase (%)
21
17
17
30
13
12
15
11
8
35
34
Table 12. Time required for Ca2+ response in cells incubated with 50 µM Cd2+ and subjected to 10
V/cm. Delay was measured between beginning of stimulation and maximum ratio increase.
90
Figure 59. Analysis of Ca2+ responses in 11 single cells incubated with 50 µM Cd2+ and subjected to 10
V/cm. First graphic shows the delay in cells to reach their calcium peak value after beginning of stimulation.
Second graphic shows the ratio increase per cell.
The percentage of osteoblasts incubated with 50 µM Cd2+ and responding with an
increase of Ca2+ was also evaluated in steps of 50 s (figure 60). Most cells (72.7%) reached
the calcium peak value in the first 50 s of field exposure. The percentage of osteoblasts
responding slower than 50 s decreased again.
Figure 60. Analysis of percentage of osteoblasts incubated with 50 µM Cd2+ that responded with an
increase of Ca2+ when subjected to 10 V/cm.
Osteoblasts incubated with 50 µM Cd2+ had an increase of Ca2+ in cytoplasm.
However the percentage of cells responding slower than 50 s reduced even more if compared
with those incubated with 25 µM Cd2+. Inclusive, in these experiments there was not a single
cell responding after 100 seconds of field exposure. Since the number of cells incubated with
50 µM Cd2+ and responding in the first 50 s increased to 72.7 %, the delay between beginning
of stimulation and calcium peak value had again a shortening. It was found with an average of
36.47 s. These observations confirmed that calcium influx was activated in cells normally
after 50 s of field application, and therefore Cd2+ had its effect only after 50 s.
91
3.203fd. 100 µM Cadmium
To confirm that Cd2+ had its effect in osteoblasts only after 50 seconds of field
exposure, Cd2+ concentration was increased to 100 µM. During this stage 3 different
experiments were carried out and 19 cells in total were evaluated (table 13, figure 61). The
delay between beginning of stimulation and calcium peak value was found with an average of
26.11 s. This delay also shortened when compared with the delay of cells incubated with 50
µM Cd2+. Ratio had an average of 35.26%.
Cell
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
Time Activation (seconds)
12,95
37,78
35,62
23,75
22,67
17,27
20,51
8,64
8,64
19,43
19,43
20,94
18,9
9,99
9,99
24,94
45,97
69,97
69,01
Fluorescence increase (%)
34
33
35
37
30
41
37
33
35
114
83
8
17
14
41
28
17
13
20
Table 13. Time required for Ca2+ response in cells incubated with 100 µM Cd2+ and subjected to 10
V/cm. Delay was measured between beginning of stimulation and maximum ratio increase.
92
Figure 61. Analysis of Ca2+ responses in 19 single cells incubated with 100 µM Cd2+ and subjected to
10 V/cm. First graphic shows the delay in cells to reach their calcium peak value after beginning of stimulation.
Second graphic shows the ratio increase per cell.
The percentage of osteoblasts incubated with 100 µM Cd2+ and responding with an
increase of Ca2+ was also evaluated in steps of 50 s (figure 62). Most cells (89.5%) reached
the calcium peak value in the first 50 s of field exposure. The percentage of osteoblasts
responding slower than 50 seconds was decreased again.
Figure 62. Analysis of percentage of osteoblasts incubated with 50 µM Cd2+ that responded with an
increase of Ca2+ when subjected to 10 V/cm.
The percentage of osteoblasts incubated with 100 µM Cd2+ and responding with an
increase of Ca2+ after 50 s reduced even more (of 27.3 to 10.5%) when compared with those
incubated with 50 µM Cd2+. Since percentage of osteoblasts responding after 50 s reduced,
therefore the percentage of osteoblasts responding before 50 s increased (of 72.7 to 89.5%); as
a consequence it produced a shortening in the delay between beginning of stimulation and
calcium peak value. Average delay reduced of 36.47 s to 26.11 s. These experiments
demonstrated that Ca2+ increase in osteoblasts after 50 s of field exposure is principally
caused by calcium influx across the membrane.
93
3.203fe. Summary of Cadmium Experiments
The percentage of osteoblasts incubated with Cd2+ that responded with an increase of
Ca2+ after 50 s of field exposure decreased according to Cd2+ concentration; higher
concentrations of Cd2+, lower percentage of osteoblasts (figure 63). These results
demonstrated that Cd2+ had its effect only in cells with a slow response. Since slow calcium
responses were assigned to a calcium influx across the membrane, Cd2+ had its effect blocking
the calcium influx. These results are consistent, as Cd2+ is known to be a potent calcium
channel blocker that inhibits influx of calcium from extracellular medium (149-151).
Furthermore these observations help to confirm that there are two different cell populations in
osteoblasts subjected to DC ELFs. Population number one with a fast calcium response (faster
than 50 s), which could be caused by Ca2+ release from intracellular stores; population number
two with a slow response (slower than 50 s), which could be caused by the calcium influx
from extracellular medium. As a consequence for the use of Cd2+, the percentage of
osteoblasts responding after 50 s was decreased, thereby the delay between beginning of
stimulation and calcium peak value was also decreased (figure 64). Shortening in the delay
seemed reasonable, as the percentage of cells responding faster than 50 s was higher, thus
average time was faster.
Figure 63. Effect of Cd2+ on the percentage of osteoblasts that responded with an increase of Ca2+ after
50 s of field exposure. It was evident that higher concentrations of Cd2+ reduced the percentage of cells.
Figure 64. Effect of Cd2+ in the delay between beginning of stimulation and peak value of Ca2+. Higher
concentrations of Cd2+ decreased the delay.
94
3.203g. Nifedipine and Nitrendipine
It has been reported that the major route of Cd2+ ions to flow into cells is through
voltage-operated calcium channels, also known as voltage-sensitive calcium channels
(VSCC), or dihydropyridine-sensitive calcium channels (L-type calcium channel) (150). It is
amply accepted that L-type VSCCs are present in osteoblasts (147, 181). To evaluate if Ca2+
ions flow into osteoblasts across L-type VSCCs when subjected to DC ELFs, and besides to
investigate whether it was the motif that caused to Cd2+ to reduce the percentage of cells with
a slow calcium response, experiments blocking L-type VSCCs were carried out. Cells were
incubated with 10 µM nifedipine and 10 µM nitrendipine after fura 2 loading. Nifedipine, Nif,
and nitrendipine, Nit, are two 1,4-dihydropyridines that act by selectively blocking L-type
VSCC in the plasma membrane (121, 122, 153).
The effect of Nif was examined in 24 cells in 5 different experiments (table 14 and
figure 65). Osteoblasts incubated with Nif responded also with an increase of Ca2+. The delay
between beginning of stimulation and calcium peak value was found with an average of 72.61
s; ratio had an average of 45.9% in cells.
Cell
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
Time Activation (seconds)
97,72
101,44
96,48
27,67
49,26
371,34
222,37
12
31
33
49,66
59,38
70,18
66,94
68,02
62,62
45,94
55,13
34,9
29,39
11
15,59
50,53
80,88
Ratio increase (%)
12
12
14
42
22
47
23
80
32
67
80
60
45
62
39
82
57
49
42
42
55
64
45
30
Table 14. Time required for Ca2+ response in cells incubated with 10 µM Nif and subjected to 10 V/cm.
Delay was measured between beginning of stimulation and maximum ratio increase.
95
Figure 63. Analysis of Ca2+ responses in 24 single cells incubated with 10 µM Nif and subjected to 10
5V/cm. First graphic shows the delay in cells to reach their calcium peak value after beginning of stimulation.
Second graphic shows the ratio increase per cell.
To evaluate and compare the effects of Nif with the effects of Cd2+ in osteoblasts, the
percentage of cells responding with an increase of Ca2+ was also evaluated in steps of 50 s
(figure 66). Nifedipine did not have an effect in cells responding in the first 50 s, but its effect
was evident in cells responding slower than 50 s. Its effect was similar to that caused by 10
µM Cd2+. Interestingly 10 µM Cd2+ has been reported to block L calcium currents (151),
which suggest that L-type VSCCs were opened in osteoblasts after 50 s of field exposure.
96
Figure 66. Analysis of percentage of osteoblasts incubated with 10 µM Nif that responded with an
increase of Ca2+ when subjected to 10 V/cm.
Nitrendipine, Nit, was also evaluated in a 10 µM concentration. 20 cells were
examined in 2 different experiments (table 15, figure 67). Osteoblasts responded with calcium
increases when were subjected to the field. However, the delay between beginning of
stimulation and calcium peak value increased to 129.62 s. This result was very contradictory,
as Cd2+ and Nif reduced the delay but Nit increased it. Ratio had an average of 30.85 % in
cells.
Cell
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
Time Activation (seconds)
82,95
109,99
70,95
61,99
111,98
57,99
83
71,99
127,98
77,99
200
144,94
60,95
24
234,96
242
208,97
206,97
224,96
188,01
Fluorescence increase (%)
26
47
40
45
12
35
58
39
15
43
18
66
21
29
16
17
23
24
16
27
Table 15. Time required for Ca2+ response in cells incubated with 10 µM Nit and subjected to 10 V/cm.
Delay was measured between beginning of stimulation and maximum ratio increase.
97
Figure 67. Analysis of Ca2+ responses in 20 single cells incubated with 10 µM Nit and subjected to 10
V/cm. First graphic shows the delay in cells to reach their calcium peak value after beginning of stimulation.
Second graphic shows the ratio increase per cell.
The percentage of osteoblasts incubated with 10µM Nit and responding with an
increase of Ca2+ was also evaluated in steps of 50 s (figure 68). Nit had its effect only in cells
that responded in the first 50 s of field exposure, in cells with a fast response. The percentage
of osteoblasts responding in the first 50 s was reduced drastically (see figure). These
observations were different when compared with those observed in cells under Cd2+ and Nif
incubation. The results were a little contradictory because Nit has been reported to block Ltype VSCC (153), exactly as Cd2+ and Nif.
Figure 68. Analysis of percentage of osteoblasts incubated with 10 µM Nit that responded with an
increase of Ca2+ when subjected to 10 V/cm.
98
3.203h. Lanthanum
The results found with Cd2+ and Nif were consistent and demonstrated that their
effects were blocking the calcium influx from extracellular medium. However the results
found with Nit were a little contradictory. Nit decreased the percentage of cells with a fast
calcium response. To confirm the results of Cd2+ and Nif, Lanthanum, La3+, a general blocker
of calcium transport across cell membrane was used to block all calcium influx in osteoblasts.
La3+ is a trivalent cation that can be used to displace extracellulary bound calcium and inhibit
calcium transport across biological membranes (152). La3+ blocks all Ca2+ entry into cells
including that entering through the passive membrane leak pathway (152). The use of La3+ to
quench extracellularly bound Ca2+ makes it possible to determine if Ca2+ influx in osteoblasts
was carried out principally after 50 s of field exposure. To achieve this purpose, 3 different
experiments were carried out and 18 cells in total were evaluated (table 16, figure 69). Cells
were incubated with 100 µM La3+ after fura 2 loading and subjected to a DC ELF. Osteoblasts
responded with an increase of Ca2+ when they were stimulated. The delay between beginning
of stimulation and calcium peak value was found with an average of 88.82 s; ratio was
measured with an average of 65.55 % in cells.
Cell
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
Time Activation (seconds)
20
13
20
39,99
30
64,95
22,96
26
153,06
61,99
66,95
34,95
184
166
213
204
204
73,99
Fluorescence increase (%)
79
92
91
81
95
87
57
61
29
26
30
43
54
61
51
71
54
64
Table 16. Time required for Ca2+ response in cells incubated with 100 µM La3+ and subjected to 10
V/cm. Delay was measured between beginning of stimulation and maximum ratio increase.
The percentage of osteoblasts incubated with 100 µM La3+ and responding with an
increase of Ca2+ was also evaluated in steps of 50 s (figure 70). Most osteoblasts (44.4 %)
responded in the first 50 s of field exposure, exactly as seen with Cd2+ and Nif experiments.
This result helped to confirm that an increase of Ca2+ in osteoblasts after 50 s of field
exposure was principally caused by calcium influx across L-type VSCCs at the membrane.
Therefore an increase of Ca2+ in osteoblasts before 50 s was probably due to calcium release
from intracellular stores.
99
Figure 69. Analysis of Ca2+ responses in 18 single cells incubated with 100 µM La3+ and subjected to 10
V/cm. First graphic shows the delay in cells to reach their calcium peak value after beginning of stimulation.
Second graphic shows the ratio increase per cell.
Figure 70. Analysis of percentage of osteoblasts incubated with 100 µM La3+ that responded with an
increase of Ca2+ when subjected to 10 V/cm.
100
3.203i. Summary of Calcium
Osteoblasts subjected to 10 V/cm DC ELFs responded with a cytoplasmic calcium
increase. The increase of Ca2+ was not observed immediately after field application, calcium
increased progressively until reaching its maximum concentration or peak value. The delay
between beginning of stimulation and calcium peak value was measured with an average of
84.59 s; peak value was measured with an increase average of 31.3 %. Experiments with
thapsigargin and manganese indicated that osteoblasts responded with an increase of Ca2+ in
two different ways: with Ca2+ release from intracellular stores, and with Ca2+ influx across the
membrane. Ca2+ release from intracellular stores was called the fast calcium response because
it was visualized preferentially in the first 50 s of field application. Ca2+ influx across the
membrane was called the slow calcium response because it was visualized preferentially after
50 s of field application. Experiments with thapsigargin and latrunculin also demonstrated
that Ca2+ release from intracellular stores might cause a calcium influx in osteoblasts.
Since Cd2+, La3+, and Nif have their effect blocking the calcium influx across the
membrane, their effect was only evident in osteoblasts responding with a slow calcium
response. Since percentage of cells responding with a slow calcium response was reduced by
the use of these drugs, the delay between the beginning of stimulation and calcium peak value
was also decreased. This shortening in the delay seemed reasonable, as the percentage of cells
responding faster than 50 s was higher, thus average time was faster.
Interestingly the delay between the beginning of stimulation and calcium peak value
showed that the electric field was not directly responsible for generating the increase of Ca2+
in the cytoplasm of cells, which suggested the existence of an intermediate mechanism.
Experiments with latrunculin suggested that this intermediate mechanism could be the
cytoskeletal contractility caused by the application of the field. Traction force microscopy
showed that an increase in traction force was visualized prior to an increase of Ca2+. However
a question appears, what is the reason for this increase of Ca2+? To address this question two
sets of experiments were carried out:
a) Cytoskeletal contractility dependent on an increase of Ca2+ was inhibited.
b) The behavior of osteoblasts under field exposure was analyzed by blocking all
calcium responses.
3.204. Myosin Light Chain Kinase
Apart from the well-characterized role of conventional myosin in muscle contraction,
members of the myosin superfamily are also involved in a number of cellular functions,
including membrane trafficking, cytokinesis, organelle transport, and signal transduction
(182, 183). The best-studied example is myosin II, the cytoplasmic analogue of smooth and
skeletal muscle myosin (159). Myosin II in association with actin filaments are responsible
for a variety of cellular movements in eukaryotic cells, such as changes in cell shape and
migration (180). Myosin II is composed of a pair of heavy chains, MHCs, a pair of essential
light chains, MLCs, and a pair of regulatory light chains, RLCs. The activity of myosin II is
regulated by phosphorylation on conserved sites (at serine 19) of its RLC. Phosphorylation
stimulates the actin-activated ATPase of myosin II and promotes the assembly of myosin II
into actin filaments. This interaction is observed in a force-productive manner leading to the
generation of tractional forces in cells (138). At present, two well-characterized kinases have
shown to catalyze the in vitro phosphorylation of the RLC at serine 19: The Ca2+/calmodulindependent myosin light chain kinase, MLCK, and Rho kinase (159, 184). The only known
101
physiological substrate for MLCK is the RLC of myosin II. MLCK is regulated by
calmodulin and requires an increase of cytoplasmic calcium (161). Rho kinase is not regulated
by Ca2+ and therefore it does not require cytoplasmic calcium (184).
When osteoblasts were subjected to DC ELFs they underwent an increase in traction
forces. The increase in traction forces was visualized prior to an increase of Ca2+ in
cytoplasm, and this was monitored prior to retraction in osteoblasts. Since an increase of Ca2+
in cytoplasm may active a myosin II-based contraction that may help to break adhesive
contacts by direct application of physical stress as reported (185, 186), myosin II-based
contraction by Ca2+/calmodulin-dependent MLCK was blocked with Wortmannin and ML-7
in osteoblasts under field exposure. Wortmannin and ML-7 are two structurally different
MLCK inhibitors that have been reported to inhibit the activation of myosin II by Ca2+/Camdependent on MLCK pathway in several cell types (160-163). The use of both drugs allowed
confirmation if an increase of Ca2+ led to the activation of MLCK and hence a myosin IIbased contraction, which could be involved in the retraction phase and therefore in the reorientation of osteoblasts. Cells were incubated with different concentrations of both drugs
and subjected to 10 V/cm DC ELFs (table17). During this stage 7 experiments with
Wortmannin and 7 experiments with ML-7 were carried out. The concentrations used were in
the range reported to cause the inhibition of myosin II-dependent contraction (160-163). Both
drugs were not able to block neither the retraction nor the re-orientation of the cells under DC
ELF exposure (figure 71). Thus the inhibition of MLCK did not cause any effect in the
alignment of osteoblasts. This observation also showed that the increase of Ca2+ visualized in
osteoblasts under field exposure was involved in other mechanism different to contraction and
traction forces in the cells.
MLCK inhibitor
ML-7
ML-7
ML-7
ML-7
Wortmannin
Wortmannin
Wortmannin
Concentrations
500 nM
1 µM
5 µM
10 µM
10 nM
100 nM
1 µM
Response
Alignment
Alignment
Alignment
Alignment
Alignment
Alignment
Alignment
Table 17. Re-orientation of osteoblasts incubated with ML-7 or Wortmannin and subjected to 10 V/cm.
Figure 71. Re-orientation perpendicular to the field gradient of osteoblasts incubated with 10µM ML-7
(A), or 1 µM Wortmannin (B). Pictures were taken after 180 min of exposure to a 10 V/cm DC ELF. Both drugs
were not able to block the alignment of the cells. Images are 200 µm large. Signs show the electric field polarity.
102
3.205. Alignment Is Dependent on Ca2+
Osteoblasts under DC ELF exposure underwent an increase in traction forces. The
increase in contractility was visualized prior to an increase of Ca2+ in the cytoplasm; Ca2+
increase was monitored prior to retraction in the cells. Experiments with Wortmannin and
ML-7 showed that the increase of Ca2+ was not involved in generating a myosin II-based
contraction, and that MLCK was not involved in the process of re-orientation of osteoblasts.
The increase of Ca2+ was principally caused by calcium release from intracellular stores.
Therefore to evaluate the role of Ca2+ release during the re-orientation of osteoblasts under
field exposure, cells were pretreated 30 min with 5 µM thapsigargin before electrical
stimulation. To achieve this purpose, 5 different experiments were carried out and 48 cells in
total were observed during 180 min under field exposure. In all cases osteoblasts did not
undergo the alignment or re-orientation process (figure 72). Cells remained without shape
changes for about 40-50 min and then underwent a strong retraction until finally disintegrated.
These observations suggested that an increase of Ca2+ caused by the release from intracellular
stores could have a regulatory role in cell shape as previously reported in other works (169,
170).
A
B
C
Figure 72. Osteoblasts incubated with 5 µM thapsigargin and subjected to a 10 V/cm DC ELF. Cells did
not undergo the re-orientation perpendicular to the field lines. Images are 200 µm large.
Since osteoblasts pretreated with thapsigargin did not undergo a retraction in the first
10 min of field exposure like a normal experiment, it suggested that an increase of Ca2+ could
be involved in the breakage of cell-substratum attachments, or activating proteins that
regulate cytoskeletal organization as previously reported (169-171, 187). However, since
molecular experiments are necessary to identify the existence of target molecules in
osteoblasts, it was very difficult to confirm these theories in this work. To date, it is not
known if osteoblasts contain the same kind of molecules like those cells mentioned in
previous references. Therefore these hypotheses remain to be evaluated in future work.
103
IV
DISCUSSION
AND
CONCLUSION
INTRODUCTION
This chapter presents the analysis of cellular responses underwent by osteoblasts under
DC ELF exposure; its first objective is to give a summary of the cellular responses observed,
and then to present an analysis and discussion of each response. Discussion was carried out
analyzing and comparing the results of this work with those previously presented by other
research groups. A theorical analysis of how a DC ELF may act at the cellular level is also
considered here. At the end of this chapter, together with the conclusion of the results, it
describes the new contributions of this research work. Finally it opens new perspectives for
further research.
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4.1. Summary of the Behavior of Osteoblasts Under DC ELF Exposure
Osteoblasts under DC ELF exposure underwent retraction, retraction-elongation,
and alignment as response to the field imposed. The changes in cell shape were dependent on
the field strength: DC ELFs greater than 12 V/cm caused only a retraction in the cells; with
DC ELFs of 7 to 12 V/cm retraction was followed by an elongation of the cells: DC ELFs
lower than 7 V/cm caused a gradual orientation of the cells. In the retraction phase,
osteoblasts started to retract their lamellar extensions that were exposed directly to the
direction of field polarity (cathode and anode), and then subsequently underwent a total
retraction of the whole cytoskeleton, causing thus a pronounced retraction of osteoblasts.
Retraction increased in speed and intensity with increased voltage, and was less pronounced
and evident with decreased voltage. In the retraction-elongation process, following the
retraction phase osteoblasts began with a highly directional re-extension of both their lamellar
long sides exposed perpendicularly to the direction of the field polarity, which caused each
osteoblast to become elongated perpendicularly to the direction of the electric field lines. The
phase of elongation in the cells was more evident and more pronounced when the cells were
subjected to longer times of exposure. In the alignment phase osteoblasts underwent a gradual
outward spreading of their long sides exposed perpendicularly to the field lines and at the
same time that contracted their broad part exposed directly in front of the field polarity. This
movement in the cells ended when each osteoblast was orientated perpendicularly to the field
lines, and then only an active elongation at each end of the cells was visualized as long as the
electric field continued.
To analyze the changes in cell shape underwent by osteoblasts by the application of
DC ELFs, experiments with phase contrast, fluorescence and traction force microscopy were
coordinated to evaluate each response. The changes in cell shape seemed to be the result of a
process of 5 steps. This process is represented in figure 73 and described below.
1.
The first effect caused in osteoblasts by the application of a DC ELF was an increase
in cytoskeletal contractility. Traction force microscopy showed that osteoblasts
underwent an increase in their traction forces after 10-30 s of field application
(problems with the recording speed did not permit to show traction force changes in
the first 10 s). The increase in traction forces was preferentially visualized at the
membrane zones directly exposed to the cathode- and anode-facing membrane, where
exactly a retraction was visualized later. The force exerted was maintained without
change until about 5 min of field exposure.
2.
The second effect visualized in osteoblasts was an increase of Ca2+ in the cytoplasm.
The increase of Ca2+ was not observed immediately after field application, calcium
increased progressively until reaching a maximum concentration or peak value; it
remained for several seconds or minutes, and then decreased progressively with some
lower oscillations until reaching its normal rest level. The delay between beginning of
stimulation and calcium peak value was measured with an average of 84.59 s;
calcium peak value was measured with an increase average of 31.3 % from its rest
level.
3.
The third effect visualized in osteoblasts was the retraction of their sides exposed
directly to the direction of field polarity. Dynamic visualization of the cytoskeleton in
osteoblasts showed that all stress fibers exposed directly in front of the field polarity
were disrupted. The beginning of stress fiber disruption was correlated with the
beginning of retraction visualized in the cells at the same time with phase contrast
105
microscopy. Osteoblasts were not able to re-organize stress fibers at the sides exposed
directly to the direction of field polarity again. However, in retraction-elongation and
alignment phases, stress fibers re-organized at the sides exposed perpendicularly to
the field lines, which aligned and elongated causing thus the re-orientation
perpendicular of osteoblasts to the field lines.
4.
The fourth effect detected in osteoblasts was a change in the direction of traction
force vectors. The contractile force exerted by the electric field during the first
seconds remained without change in the direction until approximately 5 min, and then
a decrease in traction force was visualized in the whole cell. The loss in traction force
was correlated at the same time with the retraction of the cells observed with phase
contrast microscopy (described previously in the third step). The retraction was
evident only at the sides exposed directly to the direction of field polarity, where the
increase in traction force had been visualized. With longer time of exposure, the
traction vectors showing a loss of force were visualized at both sides exposed
perpendicularly to the field lines, exactly where the cells underwent an elongation
phase later.
5.
The fifth effect visualized was an elongation of osteoblasts. After cells underwent a
loss in traction force at both sides exposed perpendicularly to the field lines, cells
began to undergo an active re-extension of their both lamellar long sides exposed
perpendicularly to the field lines, exactly where the loss in traction force had been
visualized previously. The re-extension of both long sides of osteoblasts caused each
osteoblast to become elongated perpendicularly to the direction of field polarity.
Time of exposure to the electric field was dependent on field strength; cells were not
able to accept long time of exposure when exposed to high field strengths. Time of exposure
increased when field strength decreased. It suggested that it was more difficult for cells to
adapt to high tension forces, and as consequence cells were not able to remain long time of
exposure. However, when the field strength was decreased to lower values, then it was not so
difficult for cells to adapt to the tensional force, as consequence cells accepted longer time of
exposure. Furthermore, as the tensional force was not so high, cells should try to counteract
this perturbation imposed. Therefore and since cells only elongated at the sides where
theorically the force imposed was zero (at the sides exposed perpendicularly to the field lines,
see Equation 3), it is suggested that the elongation of the cells after their retraction was a
response of adaptation. On the other hand, if the exerted tensional force decreased to very low
levels, the effect in the cells was low and slow. As consequence the effect in cells (retraction)
was almost not visualized. Furthermore, cells adapted slowly and progressively to counteract
thus the tension imposed. Thereby cells underwent a gradual re-orientation perpendicular to
the field lines. These observations suggest that the changes in cell shape were a process of
adaptation to a tensional force caused by the application of a DC ELF. To confirm this
hypothesis, in the next section every step identified will be analyzed and discussed. The
results are also compared with those previously presented by other works.
4.2. Analysis of Results
Osteoblasts under DC ELF exposure underwent a 5 step process to re-orientate
perpendicularly to the field lines and thereby adapt to the imposed tension upon them. Several
questions were opened to analyze this process of 5 steps. What mechanism caused the
observed retraction in osteoblasts? What mechanism was responsible for causing an increase
of Ca2+ in the cytoplasm, and what was the reason? What cellular elements were responsible
106
for generating the alignment observed in osteoblasts? What was the reason this alignment of
osteoblasts under DC ELF exposure? The answers to these questions are analyzed and
discussed in the following section.
Figure 73. Osteoblasts under DC ELF exposure aligned perpendicularly to the field lines in a process of
5 steps. Cells underwent firstly a cytoskeletal contraction, which was followed by an increase of Ca2+ in the
cytoplasm. Subsequently cells retracted their sides exposed directly to the direction of field polarity, and then
underwent a change in the direction of their tractional forces, which was followed by the elongation of the cells.
4.201. Cytoskeletal Contractility Defined Retraction of Osteoblasts
In this study it was demonstrated that the application of a DC ELF caused a
contractile force in osteoblasts. The contractile force was visualized as an increase in the
traction forces at the membrane zones directly exposed to the cathode- and anode-facing
membrane (section 3.107 in results). The increase in traction force was measured after 10-30 s
of field application (problems with the recording speed did not permit to show traction force
changes in the first 10 s). The results observed suggest that only those zones exposed directly
107
to the direction of field polarity were under a tensional force with a direction pulling inward
on the cell. The direction of the pulling force was maintained without change until about 5
minutes of field exposure, and then a decrease in traction force was visualized in the whole
cell. The loss in traction force was correlated at the same time with the retraction of the cells
observed with phase contrast microscopy. The retraction was evident and pronounced only at
the sides exposed directly to the direction of field polarity, where the increase in traction force
had been previously visualized. Therefore, these observations suggest that the direction of the
tractional forces had clearly defined the layout of retraction of osteoblasts before it could be
observed. Furthermore, as the increase in force disappeared immediately after the DC ELF
was removed, it showed that the contractile force was the only responsible for the retraction
of osteoblasts. Cells respreaded at the sides directly exposed to the direction of field polarity
after the electric field was removed, exactly where the retraction had taken place. How could
the contractile force cause the retraction in osteoblasts?
The maintenance of cell shape requires tension over the whole cell. Cells generate
tension within contractile microfilaments in their cytoskeleton and the extracellular matrix
resists the deformations caused by the changes in contractility. A mechanical equilibrium is
the result from the combination of forces exerted on the adhesive contacts and the forces
generated within the cytoskeleton (138). The stability of cell shape and cytoskeletal structure
results therefore from the cell’s ability to bring the internal tensional forces into equilibrium
(137). A change in the tension exerted over the whole cell will cause a change in the internal
force balance that can result in integrated changes in cell, cytoskeletal and nuclear form, and
thus influence a number of cell functions (137, 138). The forces generated within the cell by
the cytoskeleton are expressed outside the cell as traction forces in the substratum. A change
in traction forces is therefore also a change in cytoskeletal contractility.
The tensional force generated by a DC ELF had to cause a change in the internal force
balance in osteoblasts and therefore it could be the reason for retraction. The pulling force
exerted into the cell by the field should have caused an increase in tension against the cellsubstratum attachment and an increase in the cytoskeletal contractility at those sides exposed
directly to the field lines (137, 168, 185, 188). The weakening or severing of the cytoskeletonextracellular matrix linkage is at least partially driven by contractile forces (168, 186, 187). It
is likely that cells undergo an increase in tension until they reach a threshold that causes the
cell-substratum detachment (189). Thereby, an increase in tension has been frequently
observed prior to retraction in cells (188). On the other hand, an increase in traction force also
causes membrane tension. The membrane tension is also directly related to the process of
extension or retraction in the cells (190, 191). An increased membrane tension causes
retraction of cellular protrusions. Low membrane tension favors the cellular protrusions.
Thereby membrane tension can modulate the cell shape and motility as reported (190-193).
Regardless of what major role tension plays, it is evident that a tension caused by an increase
in traction forces should favor the retraction in the cells. Thus, the caused tensional force in
the cell-substratum attachment or membrane of osteoblasts by the application of a DC ELF
could stimulate their retraction. The generated contractile force could pull on filaments
connected to integrin adhesion receptors that were linked in turn to extracellular matrix
ligands. This input of force could potentially accelerate bond disruption, either at the
extracellular receptor-ligand site or at an intracellular receptor-cytoskeleton site, depending on
which connection was most labile (185, 186). Thus, the generated contractile force might help
to break adhesive contacts by direct application of physical stress and favors the retraction of
those sides where was increased, at those exposed directly to the direction of field polarity.
108
4.202. Contractile Forces Caused Increases of Ca2+ in Cytoplasm
Osteoblasts responded with an increase of Ca2+ when were subjected to DC ELFs.
However the increase of Ca2+ was not visualized immediately after field application, calcium
increased progressively until reaching maximum concentration or peak value. The delay
between beginning of stimulation and calcium peak value was measured with an average of
84.59 s; peak value (ratio) was measured with an average of 31.3 %. Analysis of Ca2+
responses in single cells, as well as experiments with manganese and thapsigargin showed
that Ca2+ responses were a contribution of both, Ca2+ release from intracellular stores and Ca2+
influx from the opening of Ca2+ channels located at the membrane. Ca2+ release from
intracellular stores was a fast calcium response because it was visualized preferentially in the
first 50 s of field application. Ca2+ influx across the membrane was a slow calcium response
because it was visualized preferentially after 50 s of field application. The use of Cd2+, La3+,
and Nif had their effect by blocking the calcium ion influx across the membrane, but no
blocking Ca2+ release from intracellular stores. As a consequence these drugs had also an
effect on the delay between beginning of stimulation and calcium peak value; the drugs
decreased the delay. The shortening in the delay was caused principally because the
percentage of cells responding faster than 50 s was higher, therefore average time was
shortened. Interestingly, the delay showed also that the electric field was not directly
responsible for generating the increase of Ca2+ in the cytoplasm. It suggested the existence of
an intermediate mechanism. What mechanism could be the responsible for causing an
increase of Ca2+ in cytoplasm?
Measurements of Ca2+ in rounded osteoblasts (without stress fibers) suggested that the
mechanism responsible for causing Ca2+ release from intracellular stores could be the
contractile force generated by the field. Previous works have reported that mechanical
stimulation can activate Ca2+ release from intracellular stores even in the complete absence of
external calcium (194). Furthermore, the cytoskeleton has also been reported to modulate
Ca2+ release from intracellular stores (195). These observations suggest a modulatory role of
the cytoskeleton to cause calcium release from intracellular stores. In this study, visualization
of calcium in rounded osteoblasts showed that an increase of Ca2+ was not carried out at the
same time as seen with intact cells. Rounded osteoblasts responded with an increase of Ca2+,
but it was after a delay of 189 s on average, much later than the 85 s showed by intact cells.
Furthermore, this average of 189 s was calculated only counting the number of cells that
responded with an increase of Ca2+, and not the total number of cells observed in the
experiments (see results, section 3.203e). These findings demonstrated that the integrity of the
cytoskeleton was necessary to cause the fast calcium response, the response caused by Ca2+
release from intracellular stores. Furthermore, since stress fibers are a support to maintain the
stability and form of the cells, osteoblasts without stress fibers could not generate tension due
to the lack of contractile filaments. Thus, the contractility seemed to be a step necessary to
generate Ca2+ release of intracellular stores and to cause the fast calcium response. The
evidence that the contractile force was always visualized prior to an increase of Ca2+ in
cytoplasm supported this hypothesis. An increase in traction force was detected in the cells
during the first 10 s of field exposure; the fastest calcium response was detected after 17 s of
field application (see table 6).
The tensional force generated by a DC ELF could also be responsible for causing
calcium influx into osteoblasts. An increase in traction forces had to generate membrane
tension, at least at the sides where cell-substratum attachments were pulled inward for the
contractile force, at those sides exposed directly to the direction of field polarity. The increase
of tension at the membrane could cause the opening of some stretch-activated calcium
109
channel as reported (196), and thus activating the calcium influx across the membrane in
osteoblasts. Stretch-activated channels are ion channels (cylinder-shaped proteins molecules)
that permit chemical diffusion (opened) in response to stress or strain on the cell membrane
(121). Thus, an increase in membrane tension could cause the opening of stretch-activated
channels. Membrane tension may also explain the increase in the delay between beginning of
stimulation and calcium peak value in rounded osteoblasts. The lack of cytoskeletal support in
rounded osteoblasts delayed the increase of tension at the membrane, and therefore, the
calcium influx. On the other hand, it has been demonstrated experimentally that a voltage can
cause movement in the membrane and with it the activation of stretch-activated channels,
where possibly L- and N-type voltage-sensitive calcium channels, VSCCs, are involved (197,
198). The movement in the membrane can be registered milliseconds or seconds after a
change in the transmembrane potential (198). In this study, experiments with DiBAC4 [3]
showed that DC ELFs caused changes in the transmembrane potential of osteoblasts
(hyperpolarization and depolarization). Changes in the transmembrane potential were not
responsible for activating directly VSCCs, as the fastest calcium response was monitored after
17 s of field application and not immediately as expected. However, a possible effect of those
changes in the transmembrane potential was perhaps the production of a movement in the
membrane of osteoblasts, which in combination with membrane tension generated the
activation of stretch-activated channels, where L- and N-type VSCCs were involved. Since
Cd2+ and Nif reduced the percentage of osteoblasts responding with an increase of Ca2+ after
50 s, it was assumed that L and N type VSCCs were involved in the calcium transport across
the membrane. Cd2+ in low concentrations is specific for L and N type VSCCs (150); Nif is
specific for L-type VSCCs (153). Therefore the use of both drugs had their effect only
blocking the calcium ion influx across the membrane.
Calcium influx in osteoblasts could be activated also by Ca2+ release from intracellular
stores. Cells that were pretreated with thapsigargin to inhibit Ca2+ release from intracellular
stores showed that the percentage of osteoblasts responding with an increase of Ca2+ was
reduced drastically to fewer than 10 % (only 6 of 35 cells). In intact osteoblasts about 50% of
cells responded in the first 50 s of field application, the other 50% was distributed between 50
and 250 s (section 3.203d). Since thapsigargin causes the depletion of Ca2+ content of
intracellular stores (122), it inhibits therefore the Ca2+ release. Thus all calcium visualized in
osteoblasts pretreated with thapsigargin was caused only by calcium influx from extracellular
medium. One would expect that the percentage of osteoblasts pretreated with thapsigargin and
responding with an increase of Ca2+ in the cytoplasm should be 50 % like normal
experiments. However, the reduction in the percentage of cells suggested that Ca2+ release
was also necessary to activate calcium influx in osteoblasts. Experiments with latrunculin also
supported this hypothesis. Ca2+ release from intracellular stores was reduced for the lack of
contractility in rounded osteoblasts; as a consequence the percentage of cells responding with
an increase of Ca2+ was also reduced drastically (only 11 of 28 cells). Taken together, these
observations indicated that Ca2+ release from intracellular stores was a necessary step to
activate Ca2+ influx into osteoblasts. Calcium influx was carried out probably by storeoperated calcium channels at the membrane, which are opened following the depletion of
internal calcium stores (122, 147, 172).
In conclusion, an increase of Ca2+ in the cytoplasm was principally caused by Ca2+
release from intracellular stores with a lower contribution of Ca2+ influx by the opening of
Ca2+ channels located at the membrane. The contractile force generated in the cytoskeleton
was probably responsible for causing Ca2+ release from intracellular stores. Calcium influx
was probably carried out by two different pathways: stretch-activated calcium channels and
store-operated calcium channels. The opening of stretch-activated calcium channels was
110
carried out probably by the increase of tension at the membrane, where possibly L- and Ntype VSCCs were involved. The activation of store-operated calcium channels was carried out
probably by depletion of intracellular calcium stores; depletion was caused by Ca2+ release,
which was driven by the contractile force. Despite these observations, it does not explain what
was the reason for an increase of Ca2+ in the cytoplasm of osteoblasts under DC ELF
exposure; the question still remains, what was the role of an increase of Ca2+ in cytoplasm?
4.203. Re-orientation Was Dependent on Calcium
Dynamic visualization of cytoskeleton showed that the shortening of stress fibers,
which is responsible for the retraction of osteoblasts, was not a direct mechanism activated by
DC ELFs. Stress fibers never underwent disruption or reorganization in the first minutes of
field exposure; shortening of stress fibers occurred after 5 (+/- 1) min, after cells had already
undergone an increase of Ca2+ in the cytoplasm. The increase of Ca2+ reached its peak value
after 85 s on average, in much less time than the visible shortening of stress fibers.
Furthermore, Ca2+ remained in the cytoplasm for some minutes. These observations suggested
that calcium was necessary to cause the reorganization of stress fibers in osteoblasts. On the
other hand, depletion of intracellular calcium stores by thapsigargin showed that Ca2+ release
was also necessary to cause the re-orientation of osteoblasts (section 3.205). Cells pretreated
with thapsigargin were not able to re-orientate, which suggested a regulatory role of Ca2+
release in changes of cell shape. A similar finding was reported by McCaig in his experiments
with neomycin, a drug that binds to polyphosphoinositodes and prevents their breakdown by
phospholipase C, and thus blocking the inositol phospholipid second messenger system that is
responsible for activating Ca2+ release from intracellular stores (173). Neomycin blocked the
perpendicular orientation of myoblasts even with a 16-fold increase of external calcium. Thus,
taken together, these findings suggest that calcium release of intracellular stores was a
significant step to cause the re-orientation of the cells. Interestingly, signaling through
phospholipase C has shown to regulate cytoskeletal organization mediating mobilization of
actin modifying proteins, including gelsolin, which can be activated by an increase of Ca2+ in
cytoplasm to interact with actin filaments (169-171). Thus, one probable consequence of the
increased cytoplasmic calcium was the activation of proteins from the gelsolin family.
Gelsolin binds to an actin filament and severs it by breaking the noncovalent bonds between
actin monomers in the polymer; the final products of the severing reaction are two actin
filaments, one of which contains gelsolin bound to the barbed end and blocks actin
polymerization from that end (199). After removing calcium excess in the cytoplasm, the
synthesis of polyphosphoinositodes (PIPs) dissociate the gelsolin from actin filament
complex. Another possible role for the increase of Ca2+ in regulating cell shape was probably
through the activation of the Ca2+-dependent protease calpain, which has been reported to
regulate rear retraction of some cells during migration (138, 187, 200). At high adhesiveness,
the physical forces exerted by the cytoskeleton are insufficient to break cell-substratum
attachments; thus it is likely that a biochemical mechanism also exists to facilitate the release
by weakening the cell-substratum linkage intracellularly (187). Inhibition of calpain in cells
has shown to block rear retraction by stabilizing cytoskeletal structures (201). Thus its
activation appears to regulate the release of cytoskeleton-extracellular matrix linkage by
destabilizing adhesions at the cells. Calpain has been localized into integrins clusters and
appears to cleave many focal adhesion proteins (201-203).
In this study, the shortening or loss of stress fibers was correlated at the same time
with the retraction of the cells and with a loss in the traction forces. These results showed that
the stress fibers lost their tensional force at the cell-substratum attachment and caused the
relaxation at the tensed elastic gel. A similar observation was found by Harris (127). He
111
showed a weakening of the cellular forces caused by a progressive relaxation of the
contractile forces exerted parallel to the axis of the voltage polarity and accompanied by the
selective retraction of the cells at those sides. Although he supposed that the weakening in
contractility might have been caused by the electrophoretic flow of a component involved in
adhesion, here it is suggested that the relaxation of the contractile force was caused by the
loss of stress fibers, as stress fibers are responsible for causing the contractile force in the
cells. In conclusion, an increase of Ca2+ in the cytoplasm of osteoblasts under field exposure
seemed to be more linked to the retraction of the cells, promoting the re-organization of actin
filaments or destabilizing the cell-substratum detachment.
If calcium was necessary to generate the re-organization of stress fibers, then calcium
responses should be different between intact osteoblasts and osteoblasts without stress fibers.
The differences in calcium responses were that the delay between beginning of stimulation
and calcium peak value increased from 85 to 189 s for rounded osteoblasts; and that the
percentage of rounded osteoblasts responding with an increase of Ca2+ was reduced to fewer
than 40 % (11 of 28 cells). As previously described, these observations showed that the
contractility generated by stress fibers was necessary to cause a fast calcium response;
however it also suggested that an increase of Ca2+ could cause the re-organization or
disruption of stress fibers. Since stress fibers were absent in rounded osteoblasts, then calcium
responses were delayed and reduced. Calcium responses in rounded osteoblasts could have
coordinated also the formation of new actin filaments. After cells removed the calcium excess
in the cytoplasm, the new synthesis of PIPs could also regulate the activity of other proteins
such as profiling or ADF/cofillin (171), in such a way as to coordinate actin dynamics and
generally promote actin filament assembly. On the other hand, an increase of Ca2+ in the
cytoplasm could have played also a role during the alignment of osteoblasts. It has been
demonstrated that the presence of polyvalent cations, like Mg2+, Mn2+, and Ca2+ neutralize
filament negative charges condensed around the charged filaments, thus the polyvalent
cations can “cross bridge” adjacent filaments, resulting in aligned bundles, which reinforce
the alignment by maximizing bridging contact areas between adjacent filament sides (204).
Therefore, an increase of Ca2+ in rounded osteoblasts could help not only forming new actin
filaments, but also reinforcing the alignment of osteoblasts.
The reason for an increase of Ca2+ in the cytoplasm of osteoblasts under field exposure
seemed therefore to be necessary for promoting the re-organization of actin filaments. In
rounded osteoblasts, an increase of Ca2+ probably formed new actin filaments. In intact
osteoblasts, an increase of Ca2+ probably caused the destabilization of cell-substratum
attachments and the shortening of stress fibers, promoting thus the retraction of the cells. In
both cases, an increase of Ca2+ probably contributed also to reinforcing the alignment of cells.
What is the reason for the re-organization of stress fibers?
4.204. Alignment Was Only Dependent on Actin
Loss of stress fibers for the use of latrunculin A or cytochalasin B caused osteoblasts
to round, as stress fibers provide structure and shape to the cells. Formation of new stress
fibers in rounded osteoblasts was only possible perpendicular to the field lines. Intact
osteoblasts lost their stress fibers exposed directly to the direction of field polarity, and only
those exposed perpendicularly to the field lines remained and elongated. In both cases,
rounded and intact osteoblasts were not able to polymerize new actin filaments along the axis
of the field polarity, in the direction of the field lines. The only visualized stress fibers
traversed commonly the entire length of the cell, which caused each osteoblast to become
orientated perpendicularly to the field lines as observed with phase contrast microscopy
112
(sections 3.102-3 and 3.201). These observations were consistent with a hypothesis previously
reported (205), suggesting that the ability of cells to change orientation may be related to the
flexibility of stress fibers to undergo re-orientation. Interestingly, other cells like keratocytes
do not re-orientate under DC ELF exposure (188, 206), but neither show abundant
longitudinal actin bundles as osteoblasts (180, 207). Stress fibers in keratocytes are only
observed at the boundary between the cell body and transition zone and are termed boundary
actin bundles. Furthermore, stress fibers are generally thought to prevent cells from
locomotion (180). Keratocytes under DC ELF exposure migrate very quickly (188, 206).
Aligned cells with stress fibers traversing the entire length of the cell (like in this study), have
not been observed to undergo locomotion under DC ELF exposure (127, 205). However, if
the actin bundles in keratocytes have a distinct organization compared with those actin
bundles in cells undergoing re-orientation is a question of further research. In this study, GFP
h1 calponin showed that the actin bundles observed were stress fibers, as h1 calponin has
been reported to incorporate along the stress fibers (135). Therefore the re-orientation of the
osteoblasts perpendicular to the field lines was caused for the flexibility of stress fibers to
undergo re-orientation. Experiments with latruncuilin A and cytochalasin B (section 3.201)
supported this observation. Osteoblasts never elongated under the influence of either drugs,
but did after the drugs were removed. Since latrunculin A and cytochalasin B are specific
inhibitors of actin polymerization, this result confirmed that the re-orientation of osteoblasts
was only dependent on actin and not other cytoskeletal proteins. This result was similar to that
reported by McCaig (173). He showed that the elongation perpendicular of cells under field
exposure was carried out under inhibition of microtubule polymerization. Thus, taken
together, these results demonstrated that the re-orientation of cells perpendicular to the field
lines under DC ELF exposure was only dependent on actin and the flexibility of stress fibers
to undergo re-orientation.
4.205. Elongation Was only Possible in Low Tension Zones
The inability of rounded osteoblasts to form stress fibers along the axis of the field
polarity, suggested that the electric field was causing a perturbation to link the integrinextracellular matrix linkage to actin filaments at the sides directly exposed to the direction of
field polarity. This perturbation seemed be the same that caused the disruption of stress fibers
at the same sides in intact cells, and thereby the retraction of the cells. In a model for a
spherical cell exposed to a DC ELF, the applied tension is zero perpendicular to the field lines
(90 grades), and increases if the angle of exposure is decreased to zero, where it has its
maximal magnitude (equation 3, section 1.401). On the other hand, it has been proposed that
tension generated by myosin II contraction may decrease the affinity of adhesion receptors for
their respective ligands (208). Membrane tension has also been reported to inhibit actin
polymerization (191). It is likely that membrane tension exerts a force against the filaments
and inhibits the polymerization. An increase in tension in rounded osteoblasts along the axis
of the field polarity could probably cause the inhibition of actin polymerization. An increased
membrane tension could also cause retraction of osteoblasts along the axis of the field
polarity. The increase in traction force at the membrane zones exposed along the axis of the
field polarity was in accordance with the possible exerted tension at those sides. Although
there is increasing evidence for the interaction of actin and myosin II in the generation of
traction forces (168, 186), it fails to explain the perturbation caused in rounded osteoblasts, as
stress fibers were absent. Experiments with ML-7 and Wortmannin confirmed that myosin IIbased contraction by Ca2+/calmodulin-dependent MLCK was not required in the reorientation process. ML-7 and Wortmannin are two structurally different inhibitors of Ca2+dependent MLCK, and both were not able to block the re-orientation of the cells when
subjected to the field. Thus, a generation of contraction by the interaction of myosin II with
113
actin, which is regulated by Ca2+/calmodulin-dependent MLCK, did not seem to be involved
in the alignment of osteoblasts. It looked reasonable, as osteoblasts under DC ELF exposure
responded with an increase of Ca2+ after 85 s on average, much later than the increase of
traction force measured. Therefore, the increase in contractility caused in osteoblasts by the
application of a DC ELF was not generated by myosin II-based contraction through
Ca2+/calmodulin-dependent MLCK, and consequently it was not responsible for the increase
in traction forces. It opens the possibility that activation of Rho-kinase could be involved in
the generation of contractility within the cells. Rho-kinase can also regulate the myosin IIbased contraction, directly activating myosin and other by inactivating myosin phosphatase
(164, 184). However such hypothesis fails too, as the generation of tension requires the
complex actin-myosin that was absent in rounded cells. An explanation that describes the
exerted tension by a DC ELF should involve both, rounded and intact osteoblasts. This
observation suggested that the contractile force and the generated tensional force at the cell
had to be independent on the integrity of the cytoskeleton. Therefore, only two possible
candidates could be responsible for generating the tension that was visualized by the increase
in traction forces: the plasma membrane and the extracellular matrix. Components of the
membrane such as ion channels, G proteins, Na+-K+-ATPase or some other component were
not thought to be responsible, as traction force vectors visualized were distributed according
to the theorical model that predicts the transmembrane potential change caused by a DC ELF
(equation 3). It is difficult to believe that some of those membrane components can distribute
uniformly on both sides of the membrane that is exposed to different polarity, and even cause
a pulling force inward the cell. There is evidence that suggest that the membrane could be the
candidate. It has been demonstrated experimentally that external electric fields caused
changes in the transmembrane potential of the cells (98, 99). It also has been shown that
membranes moved with transmembrane potential changes (197, 198). The membrane
movement can be registered milliseconds or seconds after the potential change (198). Thus, if
a DC ELF caused changes in the transmembrane potential, it could also cause a movement in
the membrane and therefore exerts a tension. Furthermore the interactions of the electric field
with the induced charges on the membrane also impose mechanical stresses on the membrane,
as consequence the membrane is deformed, elongated or compressed at the direction of the
field (86, 209, 210). In this work, the theorical change in the transmembrane potential was
directly related with the displacement of the traction vectors. Therefore, these observations
suggested that the contractile force generated by the application of a DC ELF was caused
probably by membrane tension. However, experiments measuring traction forces and
membrane potential changes at the same time are needed to test this hypothesis.
Since an increase in tension along the axis of the field polarity was the cause of
retraction in osteoblasts, and besides inhibited new synthesis of actin filaments, then new
actin polymerization and therefore cellular protrusions should be favored at zones of low
membrane tension. Exactly in those zones of low tension, the cytoskeleton-extracellular
matrix linkages should be not affected. In this case, at the sides exposed perpendicularly to
the field lines, where an increase in traction forces was not visualized. Therefore, the only
pathway possible for the spreading of osteoblasts was that perpendicular to the field lines, and
as consequence osteoblasts elongated at both sides exposed perpendicularly to the field lines.
These observations suggested that membrane tension was responsible for generating the
retraction-elongation process. Osteoblasts retracted where an increase in tension was evident
and elongated where tension was not detected. The induced changes in traction forces in
osteoblasts under DC ELF exposure were in agreement with these observations. Osteoblasts
retracted where an increase in traction force was evident and elongated where an increase in
traction force was not detected.
114
4.3. Conclusion of Results
The work presented here was originally intended to evaluate the physical behavior of
osteoblasts under DC ELF exposure. This study provided experimental evidences that the
application of a DC ELF generated a tensional force on osteoblasts up the first seconds of
field application. The tensional force was measured through an increase in the traction forces
of cells at the membrane zones directly exposed to the cathode- and anode-facing membrane.
The increase in traction forces was correlated directly with the retraction of osteoblasts at the
same sides where the increase in force was present. However the retraction of osteoblasts was
visualized after 5 (+/-1) min of DC ELF exposure, much later than the measured tensional
force. Thus before osteoblasts retracted, the direction of the traction forces had clearly defined
the layout of retraction at the sides exposed directly to the direction of field polarity.
Retraction was therefore only a result to a tensional force.
To carry out the retraction, osteoblasts responded with an increase of Ca2+ in the
cytoplasm. Calcium responses were not activated directly by the DC ELF, but were dependent
on the tensional force caused by the DC ELF and the integrity of cytoskeleton. The increase
of Ca2+ promoted the disruption of stress fibers and also the destabilization of cell-substratum
attachments, thus causing the retraction of osteoblasts. The beginning of retraction was
correlated with a decrease in traction forces in the whole cell. During the retraction phase, the
traction vectors showing a loss of force were visualized at both sides exposed perpendicularly
to the field lines. The zones showing a decrease in traction forces were correlated directly
with an elongation of osteoblasts at those sides. However, the beginning of elongation was
visualized only after 30-40 min of DC ELF exposure, much later than the decrease in force
measured. Thus before osteoblasts elongated, the direction of traction vectors showing a loss
of force had clearly defined the layout of elongation at those sides exposed perpendicularly to
the field lines. Elongation was therefore the result of a decrease in tension.
Retraction and elongation were thus consequence of changes in traction forces. Before
osteoblasts underwent any morphological change, they had clearly defined the layout to
follow by the orientation of their traction forces; the shape changes were only the result of the
changes in traction forces caused by the application of a DC ELF. Furthermore, the
distribution of traction forces generated by the DC ELF in osteoblasts was directly related to
the theorical model that shows the spatial alterations of the transmembrane potential by the
application of an external DC ELF (76). Thus the found results, force exerted-retraction and
no force exerted-extension were also in relation to the distributed theorical tension at the
membrane of osteoblasts. Therefore, the delayed morphological changes were only a process
of adaptation to a constant tensional force generated by a DC ELF at the membrane of
osteoblasts. Cells facilitate this process involving a set of coordinated biochemical
mechanisms, like an increase of Ca2+ in the cytoplasm.
Since osteoblasts under DC ELF exposure were not able to generate new synthesis of
actin filaments along the axis of the field polarity; and since osteoblasts without stress fibers
were also under a tensional force, the cytoskeleton was discarded for being responsible for
generating the tensional force in osteoblasts. It opens the possibility to suggest that either cell
membrane or the extracellular matrix were the objective of DC ELFs for exerting their
tensional force. However more experiments are needed to confirm this hypothesis. Here, it is
concluded that osteoblasts under DC ELF exposure re-orientated perpendicularly to the field
lines to counteract and adapt to a tensional force dependent on the DC ELF. To my
knowledge, no such observation has been reported. Therefore this work opens new
perspectives to further research.
115
4.301. What Is New in This Work?
An alignment perpendicular to the field lines has been reported in several cell types
under DC ELF exposure (115, 127, 166, 173, 176, 205). The mechanism exerted by the
electric field to cause the observed re-orientation of the cells has been hypothesized, but it is
still unknown. Two different theories have been proposed:
1)
Cells assume this re-orientation in order to expose their shortest dimension to the
electric field and thus minimize the caused transmembrane potential changes
(166, 211). In this case cytoskeletal changes should result by modified ion fluxes
caused by the field-induced asymmetric polarization of cells at the cathodal
versus the anodal face.
2)
Lateral electrophoresis and/or electro-osmosis of cell surface proteins or some
membrane components involved in adhesion, causing thus an asymmetric
distribution or a weakness of the adhesions between the cells and their substrate
respectively (127, 212).
The second theory has seemed to be more convincing, as cellular re-orientation was
shown to be a slow process. The delayed initial response of the cells should reflect the time
needed either for the re-organization of cell surface receptors, or for extracellular molecules to
induce a gradient of either receptor or ligand respectively.
It has recently been demonstrated that a DC ELF can induce a gradient of charged
membrane proteins. Membrane receptors like the epidermal growth factor receptor, EGFR,
showed an asymmetric distribution accumulating cathodally (114, 115). The time required for
the formation of the gradient was 10 min for corneal epithelial cells (115) and 5 min for
human keratinocytes (114). Other membrane receptors like fibroblast growth factor receptor
(FGFR), and transforming growth factor receptor (TGFR), showed the same cathodal
asymmetry but in a much slower process (115). Therefore the theory that lateral
electrophoresis and/or electro-osmosis of cell membrane components can cause the reorientation of cells seems to be convincing, however it has not been proved to date.
The present work provided experimental evidences that the application of a DC ELF
generated a contractile force on osteoblasts, which was distributed according to the theorical
model that shows the spatial alterations of transmembrane potential by the application of that
external DC ELF. These results are new, innovative, and add a potential new theory to explain
the mechanism exerted by a DC ELF to cause the re-orientation of the cells:
Cells re-orientate to counteract and adapt to a tensional mechanical force dependent
on the changes of transmembrane potential generated by the application of an external DC
ELF.
These results were discovered principally by the use of a new technology in this field
of work. To my knowledge, no laboratory has reported dynamic changes in traction forces in
cells exposed to DC ELFs using traction force microscopy. Only one previous work, that of
Albert Harris who used wrinkling elastic substrates, has ever been reported to evaluate the
mechanical behavior of cells under DC ELF exposure (127). Though at that time, wrinkling
substrates were not able to provide direct quantitative information about the detailed
magnitude, direction, and location of cell-substrate mechanical interactions like that which is
now possible with traction force microscopy. As consequence, the work of Albert Harris was
116
not taken into consideration and was forgotten. The present work was able to use the
advantages of traction force microscopy and to evaluate the mechanical response of cells
under DC ELF exposure. These new findings make this work pioneering in the area of
electro-mechanotransduction and therefore open new perspectives for further research.
The present work also provided experimental evidence that DC ELFs influenced cell
behavior similar to external mechanical forces. Previous works have reported that cytoskeletal
contractility can activate an increase of Ca2+ in cytoplasm, which may generate shape
changes. However, the stimulation of cells was carried out mechanically through shear flow
(170), or directly by mechanical stimulation (213), but not with DC ELFs like in this work.
The present work demonstrated experimentally that cytoskeletal contractility was necessary to
actively increase Ca2+ in the cytoplasm. To my knowledge, no similar findings have been
reported experimentally in cells exposed to DC ELF. Thus, taken together with the evidence
that a contractile force was generated in cells, it opens also the possibility that DC ELFs could
cause their effects on cells through exerting a mechanical force. As consequence, the
cytoskeleton induced a retraction and an elongation response, causing thus the observed
alignment of osteoblasts. Therefore this work opens a new perspective in this area that is here
called electro-mechanotransduction.
Calcium responses in cells under DC ELF exposure have been extensively reported
(99, 103, 175, 176, 178). The effects of drugs blocking calcium release from intracellular
stores or stopping calcium influx across the membrane have also been evaluated in cells under
DC ELF exposure (107, 173, 177, 206). The experiments carried out here, which involved the
visualization of calcium responses in osteoblasts under DC ELF exposure had only one
similar precedent - the work of Feder and colleagues (103). However, they used tumor mast
cells but not osteoblasts. This has been the only work that reported a similar finding in cells
under DC ELF exposure (103). Tumor cells responded with a delayed increase of Ca2+ as seen
with osteoblasts. However, they were not able to block the total increase of Ca2+ in the
cytoplasm, as they only used drugs to block calcium influx and not calcium release into the
cells. The objective of blocking only calcium influx in tumor cells was because Feder was
interested in determining the initial signal transduction generated by a redistribution of
membrane proteins, but not cell shape changes.
Previous works that have investigated the role of calcium in shape changes of cells
under DC ELF exposure reported only the effect of drugs blocking the alignment of cells
(173, 176), but not an analysis of effects caused by those drugs in single calcium responses.
Since drugs blocked the alignment of cells, little importance was given to identify the source
responsible for an increase of Ca2+ in the cytoplasm; the source was only thought according to
the drug used. Therefore, a complete study analyzing the effects of drugs in single calcium
responses of cells that underwent an alignment under DC ELF exposure has not been carried
out. Hence, this work took note of this importance and tried to identify firstly the mechanism
responsible for causing an increase of Ca2+ in the cytoplasm, and subsequently it evaluated the
effects of drugs in single calcium responses. An increase of Ca2+ in the cytoplasm of
osteoblasts was principally caused by Ca2+ release from intracellular stores with lower
contribution of Ca2+ influx by the opening of Ca2+ channels located at the membrane. Previous
works have reported that both, Ca2+ release and Ca2+ influx are necessary to activate the
alignment of cells under DC ELF exposure; inhibiting either of them blocked the alignment of
cells as was seen in this case. Therefore this work confirms previous observations, but besides
it further contributes to scientific research by providing a full analysis of single calcium
responses, which had not been previously reported.
117
The present work used the innovative technique of green fluorescence protein, GFP, to
evaluate the dynamic behavior of the cytoskeleton in osteoblasts exposed to DC ELFs. GFP is
a technique that became used approximately after year 1994 (123), and in studies involving
actin after 1996 (214). The principal advantage of using GFP is that it allows dynamic
visualization of cellular components in real time. Consequently GFP is now frequently used to
evaluate the dynamic behavior of a variety of cellular components (123). In this work GFP
was used to evaluate the dynamic behavior of stress fibers in osteoblasts under DC ELF
exposure. These new findings involving the use of GFP determined that: retraction of
osteoblasts was caused by the shortening of stress fibers at both sides of the cell exposed
directly to the direction of field polarity and; new synthesis of stress fibers was not possible at
those sides (exposed directly to the field lines). These results had only been suggested in
previous works (173, 205), but a dynamic visualization of them was lacking until now.
In conclusion this work contributed principally with four important and innovative
findings, which open new perspectives for further research.
a)
DC ELFs generate a tensional force on osteoblasts, which is distributed according
to a theorical model that shows the spatial alterations of transmembrane potential
caused by the application of those external DC ELFs.
b)
The influence of external DC ELFs on cell behavior is similar to that of external
mechanical forces. Cytoskeletal contractility is responsible for activating an
increase of Ca2+ in the cytoplasm of osteoblasts.
c)
Calcium responses of osteoblasts under DC ELF exposure are principally caused
by Ca2+ release from intracellular stores with a lesser contribution of Ca2+ influx by
the opening of Ca2+ channels located at the membrane.
d)
The tensional force on osteoblasts generated by the application of a DC ELF
inhibits new synthesis of actin filaments.
4.302. Future Perspectives
This work provided experimental evidence that a DC ELF generated a tensional force
on osteoblasts, as a consequence the cytoskeleton retracted and elongated. The first question
that rose from this was whether this contractile force was exclusively a property of bone cells
or it is also caused in other cells. To answer this question, experiments using traction force
microscopy to evaluate the mechanical behavior of different cells under DC ELF exposure
have to be carried out.
Rounded cells lacking actin filaments were even observed to be under a tensional
force by the application of a DC ELF. This observation discarded the cytoskeleton for being
the immediate target of a DC ELF. However, this observation opened the possibility of
suggesting that tensional force was exerted at the membrane or at the extracellular matrix.
Future research to evaluate both cases is required. Firstly, an analysis of the changes of
traction forces in rounded cells under DC ELF exposure should be conducted. As a second
step, experiments measuring the distribution of traction forces and membrane potential
changes at the same time are needed. The results would provide a direct correlation between
changes of transmembrane potential and of traction forces. A third and final step should be to
evaluate the changes in traction forces manipulating the extracellular matrix. Cells should be
118
cultivated in different extracellular matrixes and be subjected to the same DC ELF. The
observed changes should only be dependent on the extracellular matrix.
The importance of identifying the mechanism exerted by a DC ELF to influence the
alignment of osteoblasts perpendicular to the field lines, includes not only the physiological
significance in determining the involvement of endogenous electric fields in bone tissue, but
using electric fields as a tool to understand shaping tissues in vivo; and using electric fields to
spatially control the engineering of cells and tissues. The implementation of an in vivo system
will provide a physiological basis for determining whether the elongated shape of osteoblasts,
which are placed in the lacunae or harvesian canals in bone (described in the chapter I of this
thesis) was the result of the electric fields generated endogenously in bone tissue. Therefore,
the results will provide a physiological basis for future development of bone pathology
therapies and more effective devices of stimulation.
On the other hand, recently it has been demonstrated in vivo that the manipulation of
wound-induced electric fields in corneal epithelial cells caused the re-orientation of the cells
perpendicular to the electric field lines, and then subsequently their division, thereby
controlling an orientated cell division (215). Therefore, cellular orientation seems to be a
significant mechanism prior to cell division during electric field exposure. Thus,
understanding the controls of cellular orientation should be crucial to manipulate cell division.
The importance in controlling cell division lies in its fundamental role during development,
wound healing and pathology of any tissue and organ.
119
APPENDIX A
Solutions and Reagents Used
Earle’s salt solution 10X
Earles with antibiotics: per 1 ml Earles:
(e.g. Biochrom KG, No. L 1925)
375 IE Streptomycin
500 IE Penicillin
4 µg Amphotericin B
High Growth Enhancement Medium
(Biowhittaker, Cat. No. BESP055)
High Growth Enhancement Medium with 10% FCS, penicillin/Streptomycin, glutamin and
Vitamin C. Per 500 ml:
50 ml FCS
5 ml of a 200 mM L- Glutamin
1 ml Vitamin C- stock solution (5mg/ml water)
25000 IE of a Penicillin/ Streptomycin
Ham’s F-10 Medium
Ham’s F-10 with 10 FCS, per 500ml:
(Biochrom AG Cat No T 071-10 Lot No 282X)
50 ml FCS
1 ml Vitamin C-stock solution (5mg/ml water)
25000 IE Penicillin/Streptomycin
5 ml of a 200 mM L-Glutamin
Components of Ham’s F-10 Medium (mg/L)
L-Arginine HCl 211.00
L-Aspartic Acid 13.00
L-Asparagine H2O 15.00
L-Alanine 8.90
Biotin 0.007
CuSO4 5H2O 0.0025
L-Cysteine HCl H2O 35.00
Choline Chloride 14.00
CaCl2 (anhyd.) 33.20
FeSO4 7H2O 0.83
Folic Acid 1.30
Glycine 7.50
L-Glutamic Acid 14.70
L-Glutamine 146.00
D-Glucose 1802.00
L-Histidine HCl H2O 21.00
Hypoxanthine Na 4.77
L-Isoleucine 4.00
L-Lysine HCl 36.50
L-Leucine 13.00
Linoleic Acid 0.08
Lipoic Acid 0.21
i-Inositol 18.00
MgCl2 (anhyd.) 57.22
L-Methionine 4.50
Niacinamide 0.04
Pyridoxine HCl 0.06
D-Ca Pantothenate 0.50
120
Phenol Red 1.20
Sodium Pyruvate 110.00
KCl 223.60
L-Phenylalanine 5.00
L-Proline 34.50
Putrescine 2HCl 0.16
Riboflavin 0.04
L-Serine 10.50
NaCl 7599.00
Na2HPO4 (anhyd.) 142.00
NaHCO3 1176.00
L-Threonine 12.00
L-Tryptophan 2.00
Thiamine HCl 0.30
Thymidine 0.70
L-Valine 11.70
Vitamin B12 1.40
ZnSO4 7H2O 0.86
L-Tyrosine 2Na 2H2O 7.80
Ham’s F-10 medium with 10 mM Hepes, PH 7.25 (1 liter preparation):
1. 9,89 g HAMS F10 (Instamed)
2. 5 ml L-Glutamin (200mMol)
3. 5 ml Vit C (2mg\ml HAMS F10)
4. 2 ml sodium hydrogen carbonate (NaHCO3) (60g/l stock solution) 2,383g
HEPES Powder
5. 50 000 IE Penicillin/Streptomycin (5ml stock solution)
Reagents used:
• Penicillinß\Streptomycin
(Boehringer Mannheim, Cat. No. 1074440)
• Amphotericin B
(Boehringer Mannheim)
• L- Glutamin- Solution
(e.g. Sigma, Cat. No. G-7513)
• Vitamin. C
(Wako Chemical Industries, Japan)
• FCS
(Boehringer Mannheim Charge number
14713902)
• PBS solution
(Biochrom KG, Cat. No. L 182- 50 9,55 g/ l)
• Ham’s F-10 medium
(Biochrom AG Cat No T 071-10 Lot No 282X)
• Trypsin/EDTA
1ml/100 ml PBS (Sigma, Cat No T4174)
• Lanthanum Chloride, LaCl3
(Sigma)
• Cadmium Chloride, CdCl2
(Sigma)
• NaCL
(Sigma)
• ML-7, 1-(5-iodonaphthalene-1-sulfonyl)-1h-hexahydro-1,4-diazepine,
C15H17IN2O2S.HCL
(Sigma)
• Nifedipine C17H18N2O6
(Sigma)
• Nitrendipine, C18H20N2O6
(Sigma)
• Manganese Chloride, MnCl2
(Sigma)
• Latrunculin A, latrunculia Magnifica
(Calbiochem)
• Cytochalasin B
(Calbiochem)
• Thapsigargin
(Molecular Probes)
• Fura 2AM
(Molecular Probes)
121
•
•
•
•
•
•
•
•
•
•
•
•
DiBAC4(3)
(Molecular Probes)
Wortmannin
(Calbiochem)
DMSO, Dimethyl sulfoxid
(Roth A994.1)
Agarose Salt, Ultra pure agarose, electrophoresis grade. (Life Technologies, Paisley
Scotland)
Transfection kit, Effectene Transfection Reagent (Effectene, Enhancer, Buffer EC)
(1ml)
(Qiagen Cat. No. 301425)
EGFP-CaP h1 plasmid
(provided by Molecular Biology Institute
from Austrian Academy of Sciences, Billrothstrasse 11, A5020 Salzburg Austria)
Coverslips
(No. 1, 60X24 mm Marienfeld, Germany)
Circular coverslips
(No. 1: ∅ 16 mm, Marienfeld, Germany)
Acrylamide and Fluorescence beads
(Fluo-Sphere, Molecular Probes, Leiden,
Netherlands)
Multimeter
(Model 75. Fluke)
Power Supply
(Model 200/2.0 voltage-regulated. Biorad)
pH meter
(Inolab pH level 1, wtw, Germany)
122
REFERENCES
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137
MIGUEL MIRON MENDOZA
Date of Birth
Place of Birth
Nacionality
June 5th 1971
Orizaba, Mexico
Mexican
Education
1992-1995
Center of research and advanced studies of IPN
(CINVESTAV-IPN), Mexico.
MASTER IN SCIENCES
Speciality: Electrical Engineering, Bioelectronic Section
Thesis: System for stimulation, adquisition, display and
processing of electrophysiological signals from cells. Mexico
City. Mexico.
1988-1992
Instituto Tecnologico de Orizaba, Mexico.
ELECTRONIC ENGINEER
1986-1988
Instituto Pluviosilla, Orizaba, Mexico.
HIGH SCHOOL
1983-1986
Esc. Sec. Fed. Ignacio Manuel Altamirano.
Cd. Mendoza, Mexico.
SECUNDARY
1977-1983
Esc. Prim. Enrique C. Rebsamen. Cd. Mendoza, Mexico
PRIMARY
Professional Experience
October 2001 – September 2002
Philipps-Marburg University, Germany
Department of Experimental Orthopaedics and Biomechanics.
Research Assistant: Effects of mechanical forces on bone
cells.
December 1996 - March 1997
Company: Medios y Procedimientos s.a. Mexico
Engineer: Development of measurement and control systems.
November 1996 – March 1997
National Autonomous University of Mexico (UNAM)
Engineer: development of Instrumentation to calcium
measurements in cells.
September 1996 - February 1997. Military University of Mexico (EMI-SDN)
Teacher.
January 1995 - December 1996
Technical University of Mexico (UNITEC)
Teacher.
138