Cell Shape & Integrins:
Determinants of Extracellular Matrix
Regulation of Growth & Survival
by
Christopher S. Chen
B.S., Biochemistry (1990)
Harvard College
M.S., Mechanical Engineering (1993)
Massachusetts Institute of Technology
Submitted to the Division of Health Sciences and Technology
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Medical Engineering
at the
Massachusetts Institute of Technology
@ 1997 Massachusetts Institute of Technology
All rights reserved
Signature redacted
Signature of A uthor ........................................................
Christopher S. Chen
May 7, 1997
Signature redacted
Certified by..............................................................
.......
DonaldE. Jngber
Associate Professor of Pathology
Harvard Medical School
Thesis Supervisor
I,
,
Signature redacted
Accepted by .................................................................
MarthaL. Gr y
Interim Director, Division of Health Sciences and Technology
MAY 2 7 1997
CELL SHAPE & INTEGRINS:
DETERMINANTS OF EXTRACELLULAR MATRIX
REGULATION OF GROWTH & SURVIVAL
by Christopher S. Chen
Submitted to the Division of Health Sciences and Technology
on May 7, 1997 in partial fulfillment of the requirements for the degree of
Doctor of Philosophy in Medical Engineering
ABSTRACT
The local modulation of cell proliferation (growth) and apoptosis
(programmed cell death) that is essential to the development and
maintenance of tissue pattern is regulated in part by binding interactions
between cells and the surrounding insoluble extracellular matrix (ECM).
Evidence suggests that adhesion to ECM regulates cell function by controlling
the shape of a cell. The focus of this study was to investigate the role of cell
shape in the regulation of cell proliferation and apoptosis by adhesion to ECM.
Growth and apoptosis were examined in capillary endothelial cells whose
shapes were controlled by adhesion to substrates microfabricated with
geometric patterns of ECM. Using microcontact printing (gCP) of selfassembled monolayers (SAMs) of alkanethiolates on gold, we manufactured
substrates that contained micrometer-scale islands of ECM such that cells
attached and spread to the size and shape of the engineered islands. In
addition, novel methods were developed to pattern cells on transparent
substrates contoured with 3-dimensional topography and to attach cells
biospecifically to peptide ligands contained within the SAM.
Progressively restricting bovine and human endothelial cell spreading on
ECM-coated SAMs regulated a transition from growth to quiescence to
apoptosis on a single continuum of cell spreading, in the presence of
saturating levels of soluble growth factor. Shape-dependent apoptosis was
observed regardless of the integrins being engaged by the substrate. By
designing the size and spacing of focal adhesion-sized ECM islands to
promote the spreading of cells across multiple islands, cell shape could be
varied using a constant, low total ECM-cell contact area. Cell growth and
apoptosis were found to be regulated directly by spreading of the cell and
nucleus, and not by total ECM in contact with a cell, or by the quantity of focal
adhesion formed per cell. Additional studies exploring the mechanism by
which binding of ECM controls cell and nuclear structure revealed that
coordinated shape transformations are controlled by mechanical forces
transmitted directly through a discrete cytoskeletal lattice to connect integrins
to the nucleus. In sum, cell geometry is a central mediator of ECM regulation
of cell proliferation and apoptosis.
Thesis Supervisor: Donald E. Ingber
Title: Associate Professor of Pathology, Harvard Medical School
2
TABLE OF CONTENTS
CHAPTER I. GENERAL INTRODUCTION
Extracellular m atrix .....................................................................................
ECM signaling: integrins and cell shape .....................................................
Regulation of the cell cycle.........................................................................8
Regulation of apoptosis .............................................................................
Mechanics of cell deformation................................................................12
Engineering ECM environments ...............................................................
Experim ental Design ..................................................................................
CHAPTER II. DESIGN AND FABRICATION OF SUBSTRATES
Preface ..........................................................................................................
Patterned surfaces: Microcontact printing of SAMs to
pattern ECM and cells on flat surfaces..................................................
Contoured surfaces: Controlling cell attachment on
contoured surfaces with self-assembled monolayers
of alkanethiolates on gold....................................................................
Biospecific surfaces: Direct attachment at spreading of cells
to mixed self-assembled monolayers presenting GRGD
and (EG )30 H groups..................................................................................
.
F ig u res ........................................................................................................
5
6
10
13
16
. 19
20
27
37
. 56
CHAPTER III. ECM REGULATION OF GROWTH AND APOPTOSIS
. 96
P reface ..........................................................................................................
death................................................96
life
and
Geometric control of cell
104
A p p en d ix A ...................................................................................................
113
F ig u res ............................................................................................................
CHAPTER IV. MECHANICAL BASIS OF CELL & NUCLEAR
DEFORMATION
15 1
Preface .............................................................................................................
Demonstration of mechanical connections between
integrins, cytoskeletal filaments, and nucleoplasm
that stabilize nuclear structure.............................................................152
170
Ap p en d ix B ....................................................................................................
176
F ig ures ............................................................................................................
CHAPTER V. CONCLUSIONS.............................................................................
192
BIBLIO G RA PH Y .......................................................................................................
201
ACKNOWLEDGMENTS........................................................................................
216
3
Chapter 1 General Introduction
CHAPTER I. GENERAL INTRODUCTION
Emerging half a billion years ago, multicellular organisms have
evolved such structural complexity, from plants to invertebrates to
mammals, that we have only begun to discover the fundamental
mechanisms which govern their anatomical organization.
The concerted
action of soluble growth factors, extracellular matrix (ECM), and mechanical
forces act throughout life to constantly regulate cell shape, proliferation,
differentiation, migration, and apoptosis (programmed cell death) to develop
and maintain the architecture of complex tissues, such as branching capillary
networks. Unlike growth factor concentrations, binding interactions with the
ECM can change dramatically over micrometer distances, establishing the
local differentials in cellular functions required to drive pattern formation
[Ingber, 1985; Ingber and Folkman, 1989]. Cells respond to the ECM through
several "sensing" mechanisms, which include the binding and activation of
integrins, a class of ECM receptors [Tamkun et al., 1986; Hynes, 1992; Clarke
and Brugge, 1995]; the formation of focal adhesion complexes, a subcellular
structure of clustered integrins, cytoskeletal elements, and signaling
molecules [Burridge et al., 1988; Craig and Johnson, 1996]; and the changes in
cell shape which intrinsically modulates many cellular functions [Ingber,
1990; Ingber and Folkman, 1989; Mooney et al., 1992; Singhvi et al., 1994; Watt
et al., 1988; Ingber, 1997]. The working hypothesis of this thesis is that cell
shape mediates the control of cell growth and apoptosis by ECM.
Understanding the mechanism of capillary growth and regression
could pave the way to new therapeutic approaches in several clinical areas.
Because capillaries provides the conduit of nutrients necessary for the growth
4
Chapter 1 General Introduction
and maintenance of the surrounding tissue, the stimulation of angiogenesis
(the formation of new capillaries) could be used therapeutically, for example,
to increase oxygen delivery to failing myocardium. Conversely, inhibition of
angiogenesis and induction of capillary regression has been shown to block
solid tumor growth [Ingber et al., 1990; Holmgren et al., 1994; O'Reilly et al.,
1995]. Thus, uncovering the mechanisms by which the ECM acts to regulate
capillary cell proliferation and apoptosis has enormous clinical implications.
This chapter contains a brief introduction to the ECM and reviews the
following relevant research areas: (1) signal transduction paradigms for cell
adhesion, (2) regulation of proliferation and apoptosis, (3) relationship
between cell mechanics and shape control, and (4) methods for engineering in
vitro ECM environments. I conclude with a description of the experimental
approach I have taken to determine the role of cell shape in ECM-mediated
growth and apoptosis.
Extracellular matrix
ECMs are insoluble macromolecular networks that function as physical
scaffolds to hold cells and tissues together. The enormous range of
mechanical, material, and chemical properties found in ECMs arise from the
hundreds of proteins and polysaccharides that are secreted by cells and
assembled into 3-dimensional matrices. These components can be classified
into: collagens, elastins, proteoglycans, and glycoproteins. Collagens are large
molecules that usually form the backbone structure of the matrix [Nimni,
1983]. They provide the tensile strength of the tissue. The random coiled-coil
structure of elastins adds a rubber-like elastic property to ECM [Aaron and
Gosline, 1981]. Proteoglycans are highly charged molecules that capture water,
producing a volume-filling gel. The force of hydration resulting from
5
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Chapter 1 General Introduction
proteoglycans embedded in ECM provides a compressive strength to the
tissue [Silbert, 1987]. The large glycoproteins act as key players in cell-ECM
interactions, containing binding sites for many ECM molecules as well as
specific cell surface receptors. In this investigation, I plan to focus primarily
on the effects of fibronectin, a 200-230 kD glycoprotein found in nearly all
ECMs. Fibronectin has binding sites for collagens, heparin, and cell surface
integrin receptors, as well as a synergy site that enhances the avidity of
integrin binding [Hynes, 1989; Nagai et al, 1991].
ECM signaling: integrins and cell shape
Binding of ECM to cell surface receptors elicits a variety of cellular
responses which may or may not have signaling pathways common to each
other. These pathways can be broadly categorized to stem from integrin
signaling or cell spreading.
Integrins, a class of heterodimeric receptors, initiate a variety of
signaling cascades upon binding to ECM [Clarke and Brugge, 1995]. Like other
signaling receptors, occupancy and multivalent aggregation are both required
for full activation of the integrins [Miyamoto et al, 1995]. Occupancy alone
appears to promote redistribution of the receptors to preexisting integrin
clusters. Aggregation using soluble antibodies to non-active site regions of
integrins does not appear to play a role in activation. However, aggregation
using the same antibodies adsorbed to solid substrata colocalizes pp125FAK
(focal adhesion kinase) and tensin. Their phosphorylation then attracts a
battery of signal transduction molecules, such as RhoA, Rac, Ras, MEKK,
ERK1, ERK2, JNK, PLC-gamma, PI 3-K, and c-Src [Miyamoto et al, 1995]. If
both receptor occupancy and aggregration are allowed, the structural proteins
vinculin, talin, alpha-actinin, and F-actin associate with the aggregate to form
6
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Chapter 1 General Introduction
a mature focal adhesion [Plopper et al, 1993; McNamee et al, 1993; Miyamoto
et al, 1995]. These focal adhesion complexes mechanically couple the ECM to
the cytoskeleton [Wang et al., 1993; Ezzell et al., 1997], as well as to provide a
site for promoting the recruitment of multiple signaling molecules, primarily
through tyrosine phosphorylation events [Burridge, 1988; Plopper et al, 1995;
Schwartz et al, 1995]. These signaling molecules then initiate several cascades
that have been shown to play a role in cell function (e.g., the MAPK pathway
for proliferation; the rho GTPase pathways for cytoskeletal rearrangements).
Thus integrin signaling can be measured either by level of phosphotyrosine
production or by amount of accumulation of cytoskeletal linker proteins.
Initiating adhesion signals leads to changes in cytoskeletal
polymerization and tension generation within the actin-myosin filament
lattice that cause active extension of membrane processes across the substrate
and subsequent spreading of the cell [Nobes and Hall, 1995]. This physical
spreading of cells in itself also appears to feed back to alter intracellular
signaling, and ultimately, cell behavior. For example, adhesion-induced
activation of adenylate cyclase and subsequent increase in intracellular cAMP
is only observed in spreading cells, not in round cells [Fong and Ingber, 1996].
These shape-sensitive signals, in turn, could be critical for the regulation of
cell growth and apoptosis.
Integrin signaling and cell spreading offer diametrically different
mechanisms of biological regulation. While the former proposes that soluble
signals diffuse from integrins to the nucleus, the spreading model suggests
that mechanical structure and tension are transduced. Distinguishing
between these disparate hypotheses would be a critical initial step in directing
future research in this field.
7
Chapter 1 General Introduction
ECM regulation of growth
In vivo observations have led investigators to examine ECM as a
regulatory signal for cell growth and apoptosis.
A classic example is the life
cycle of keratinocytes, the cells that form mammalian skin. Keratinocytes
constantly multiply while attached to the basement membrane of the dermalepidermal boundary. When one is pushed off the basement membrane, it
loses integrin expression, differentiates to produce a highly crosslinked lattice
of involucrin and cytoskeletal keratins, and finally undergoes apoptosis (see
next section), leaving behind a protective sheath on the surface of skin [Fuchs,
1990; Adams and Watt, 1993]. Experimenters demonstrated in vitro that
detaching cells from their substrate would lead to terminal differentiation
and apoptosis. This response could be halted by ECM re-exposure [Adams et
al, 1993]. It was also shown that skin stem cells would only proliferate when
attached to ECM [Fuchs, 1990]. These types of experiments led to a series of
advances in understanding both growth and apoptosis regulation.
The cell cycle is a complex series of highly regulated events leading to
cell division, and is divided into Go (quiescence), Gi (preparation for DNA
synthesis), S (DNA synthesis), G2 (preparation for mitosis), and M (mitosis)
phases. Fibroblasts detached from ECM while in G1 returned to GO, while
those in S continued through cell division before arresting [Campisi et al,
1983]. Similarly, the dependence of cell cycle progression on soluble growth
factors is limited to GI [Campisi et al, 1983]. After passing a restriction point
"R" in late G1, cells will complete the cell cycle independent of mitogens [Yen
and Pardee, 1978]. These findings appear to be linked, as ECM modulates cell
sensitivity to growth factors [McNamee et al, 1993; Plopper et al, 1995; Ingber
et al., 1990]. Recent work has further demonstrated that certain cyclins and
cyclin-dependent kinases, important regulators of Gi progression, depend on
8
---I
Chapter 1 General Introduction
adhesion for expression and activation; constitutive expression of these
factors transforms cells, enabling them to grow in suspension [Guadagno et al,
1993; Zhu et al, 1996]. Our laboratory has shown that either spreading cells on
fibronectin-coated plates or binding round, suspended cells to many
fibronectin-coated beads leads to similar activation of the immediate-early
genes, including c-fos, c-myc and c-jun (the earliest signals of Gi progression).
However, only spread cells fully progress through Gi to enter S-phase [Dike
and Ingber, 1996]. This result suggests that ECM exerts its influence on cell
growth at two different levels: (1) Exposure to insoluble ECM leads to full
activation of early cell cycle events, perhaps through direct integrin signaling;
and (2) the presence of ECM on a rigid substrate promotes later cell cycle
progression, perhaps through cell spreading. Several studies have attempted
to examine the importance of spreading in growth regulation.
Folkman and Moscona (1978) first demonstrated that proliferation of
cells correlates with the degree of cell flattening against a solid substrate. Our
laboratory has since found that this adhesion is mediated through integrins
bound to insoluble ECM. When endothelial cells are cultured on increasing
densities of fibronectin, both the extent of cell spreading and the rate of
proliferation are increased [Ingber, 1990]. Interestingly, when cells are
cultured on intermediate densities of fibronectin, they differentiate and form
capillary tubes [Ingber et al, 1989]. This ECM regulated switch between
differentiation and growth has been shown for hepatocytes and adipocytes as
well [Mooney et al, 1992; Spiegelman et al, 1983]. However, since cell
spreading is achieved by increasing ECM density, this substrate system could
not distinguish whether the observed effects were due to the cell spreading
per se or to the ECM density directly. Using a microfabrication approach to
restrict cell spreading while maintaining a constant density of ECM, our
9
Chapter 1 General Introduction
laboratory demonstrated that cell growth arrests when single primary rat
hepatocytes were cultured on progressively smaller micrometer-scale
rectangular islands of laminin, another ECM glycoprotein [Singhvi et al,
1994]. Although ECM density was eliminated as a potential confounding
factor in these experiments, the total amount of ECM presented per cell
covaries with cell spreading. In the present investigation we therefore set out
to separate these two factors using an alternative micropattern design.
ECM regulation of apoptosis
Apoptosis is a highly regulated series of events that ultimately results
in cell death. Unlike necrosis, which results from tissue injury, apoptosis
does not elicit an inflammatory response and culminates in the rapid and
efficient removal of the dead cell by its neighbors. Apoptosis is a critical part
of normal development. For example, the Caenorhabditis elegans
hermaphrodite consistently produces 1030 cells during its development, and
131 of the cells die by apoptosis [Vaux et al, 1992]. In the mammalian
immune system, T cells are selected to only recognize non-self peptides; those
that bind self-peptides during their maturation in the thymus are induced to
undergo apoptosis [von Boehmer, 1992].
ECM appears to play an important role in regulating the apoptotic
program. Detachment of many cell types from ECM induces apoptosis,
including endothelium, keratinocytes, mammary epithelium,
gastrointestinal epithelium, MDCK cells, and fibroblasts [Re et al, 1994; Pullan
et al, 1996; Bates et al, 1994; Frisch et al, 1994; Meredith et al, 1993]. Increased
local degradation of ECM in vivo, by pharmacologic or genetic means,
induces apoptosis of adjacent cells and involution of surrounding tissues
[Ingber et al., 1986; Ingber and Folkman, 1989; Talhouk et al, 1992]. Using
10
Chapter 1 General Introduction
soluble antagonists to disrupt cell surface integrins from binding ECM
induces apoptosis in vitro and involution of tissues in vivo [Brooks et al,
1995]. Cells survive when attached to substrates coated with anti-integrin
antibody. These findings suggest that binding and activation of integrins
appears to be the key regulator for survival. However, in vivo and in vitro
studies suggest that the cell spreading that results from cell attachment to
ECM may be in fact the survival signal: Analysis of regressing capillaries in
vivo indicates that cells are still surrounded by the fragmented ECM when
dying, and instead cell contraction and rounding appears to act as the signal
for apoptosis [Ingber et al., 1986]. Furthermore, endothelial cells flattened
over large (100 um diameter) microcarrier beads grow, while round,
suspended cells attached to as many as 10 small (4.5 um) beads die [Ingber and
Folkman, 1989; Re et al, 1993]. However, none of these studies focused
specifically on the issue of whether cell shape per se regulates apoptosis.
Unlike for cell proliferation, little is known about the signal
transduction pathways for apoptosis. It appears that many different stimuli
induce apoptosis through distinct but convergent signaling pathways.
Although the exact level of convergence is not known, many forms of
apoptosis involve the activation of the ICE-family proteases, which carry out
the destruction of many intracellular targets. Detachment from ECM has
been shown to increase the expression of ICE [Boudreau et al., 1995]. Among
the upstream factors, the balance of intracellular bcl-2 and bax levels appears
to play a major role in control of the apoptosis program [Korsmeyer et al.,
1993]. Overexpression of bcl-2 can prevent apoptosis and override cellular
requirements for adhesion to ECM [Frisch et al., 1994]. Detachment from ECM
does not affect bcl-2 expression, but increases bax in dying mammary
epithelium [Pullan et al, 1996]. Even further upstream in ECM-mediated
11
__Z
Chapter 1 General Introduction
apoptosis, it has recently been reported that constitutive activation of
pp125FAK, a major component of the focal adhesion complex, results in
shape- and adhesion-independent survival, implicating the FAC as a
potential key mediator for apoptosis. One focus of this investigation,
therefore, is to determine the relative importance of focal adhesion formation
and cell shape changes in cell function.
Mechanics of cell deformation
Although cell and nuclear shape appear to be critical for the control of
cell function [Ingber et al., 1987; Yen and Pardee, 1979], little is known about
the mechanism by which forces transmit through ECM-integrin interactions
to control cell shape. Cells deform in response to applied forces as nonlinear
viscoelastic solids. That is, upon force application, there is an immediate
elastic deformation, followed on the order of minutes by a slower plastic
deformation [Wang et al., 1993; Evans and Yeung, 1989; Chien et al., 1984]. An
elastic deformation is defined such that release of the force results in
immediate recoil of the cell back to its original shape, while a plastic
deformation does not recoil. Early models developed to explain this
mechanical behavior assumed that the mechanical properties of the actin
cortical membrane dominated the system and was solely responsible for the
observed viscoelastic response [Elson, 1988; Drochon et al., 1990; Evans and
Yeung, 1989; Keller and Skalak, 1982]. These models would suggest that there
is no physical connection from the ECM into the interior of the cell.
However, studies of the active movements of cell protrusion and
lamellipodial extension suggest that the polymerization and extension of
cytoplasmic microfilaments and microtubules into the cortical membrane can
deform it, forcing it to protrude out into the environment [Kolega et al., 1991;
12
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Chapter 1 General Introduction
Bray and White, 1988; Condeelis and Taylor, 1977]. If membrane mechanics
dominated, these cytoplasmic filaments would be unable to deform it.
Furthermore, the mechanical properties of purified cytoskeletal filaments
suggest that all three filament systems - microfilaments, microtubules, and
intermediate filaments - contribute to the mechanical strength of a cell [Sato
et al., 1985; Janmey et al, 1991; Janmey, 1991]. In fact, cells with dysfunctional
intermediate filaments are more fragile, and rupture easily [Fuchs and Weber,
1994]. However, to truly answer the question of what role the cell cytoplasm
plays in cell mechanics, we must directly examine the deformation response
of cells to applied forces.
Engineering ECM environments
One approach to studying the role of shape in cell function is to control
cell shape through its adhesive interactions with the ECM environment, and
observe the resultant changes in cell behavior. For example, increasing the
density of ECM adsorbed to a surface increases cell adhesion and spreading
against the substrate, and alters the subsequent proliferative response [Ingber,
1990]. An alternative method has been to try to fabricate "islands" of ECM
surrounded by nonadhesive regions, such that single cells would attach and
spread only to the size of the island. Thus, by engineering the size and shape
of these islands, cell shape could be exquisitely controlled.
Historically, the investigation of cellular responses to various adhesive
environments were limited by a lack of control over the material properties,
surface chemistry, and surface topology of available substrates. It was
particularly difficult to generate substrates patterned with adjacent adhesive
and nonadhesive regions. In the past decade, the technology to engineer
patterned biological substrates has rapidly advanced, partly as a result of
13
- -2
Chapter 1 General Introduction
modification of microfabrication techniques used in the electronics industry.
This powerful class of techniques allows investigators to pattern defined
topographies and surface chemistries onto substrates with varying degrees of
precision, depending on the methods used. There are few variations on the
choice of methods to engineer surface topography, typically achieved by
anisotropic chemical or plasma etching of silicon or glass. The resulting etch
topologies are also limited to square, V-shaped, and semi-circular crosssections.
Generation of patterned surface chemistry can be achieved using
several techniques: vapor deposition, photolithography, and microcontact
printing. Vapor deposition of metals through a patterned grid onto polyhydroxyethyl methacrylate (pHEMA) results in a substrate containing
complementary patterns of metal and pHEMA. Cells can be selectively
attached to the metallic regions because they adhere to the metal, but not the
pHEMA [LeTourneau, 1975; O'Neill et al, 1986]; however, this method
produces low resolution (5 gm) patterns and the surface interactions with
proteins and cells is not well-defined. Furthermore, this technique does not
allow the immobilization of specific ECM molecules to the pattern.
Photolithography has been used to routinely produce patterns of
defined surface chemistries with resolutions better than 1 gm. This approach
uses ultraviolet light to illuminate photosensitive materials through a mask
that contains the desired pattern. This technique has been used to directly
photoablate proteins preadsorbed to a silicon or glass surface [Hammarback et
al, 1985], or to covalently link preadsorbed protein onto a photosensitive
group [Matsuda, 1995]. Photolithography has also been used in a three step
process, where photoresist was selectively removed to expose the underlying
glass or silicon surface. Then silanes could be adsorbed to the bare surfaces,
14
Chapter 1 General Introduction
followed by protein adsorption. Finally, the remaining photoresist could be
removed and filled in with a hydrophobic silane [Bhatia et al., 1994]. A major
problem with these approaches is that the "nonadhesive" regions of the
pattern are usually surfaces that actually promote protein adsorption, and
require passivation (blocking of adhesive sites) with a nonadhesive protein
such as albumin. Over a period of days, however, cells are able to migrate
onto these regions, probably as a result of degradation of the albumin and
deposition of ECM by cells. Several investigators have dealt with this issue by
using photolithography to pattern siloxane monolayers presenting perfluoroand amino-terminated moieties, demonstrating preferential adhesion of cells
to the amino-terminated siloxane without passivation of the perfluoroterminated regions by albumin. The formation of siloxane monolayers,
however, proves to be technically challenging. Furthermore, the
photolithography and clean room facilities required for all these approaches
are costly to build and maintain.
Recent advances in the chemistry of self-assembled monolayers
(SAMs) of alkanethiolates on gold surfaces has provided a different approach
to the patterning of cells. These SAMs are highly ordered molecular
assemblies that chemisorb on surfaces to produce effectively two-dimensional
crystals with controllable chemical functionality [Whitesides and Gorman,
1995]. SAMs provide one of the most promising systems to accurately and
easily control the surface chemistry of a substrate. Previous work has shown
that while hydrophobic SAMs rapidly and irreversibly adsorb proteins and
promote cell adhesion, SAMs that present ethylene glycol moieties effectively
resist protein adsorption and cell adhesion [Prime and Whitesides, 1991;
Prime and Whitesides, 1993]. Thus, patterning of these two SAMs onto a
substrate can provide a substrate that patterns cells without the need for a
15
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-
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11 Trill
111
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-
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-
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11
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In
Chapter 1 General Introduction
passivating agent like albumin. In fact, we have demonstrated that cells
cultured on substrates patterned with these two SAMs do not invade ethylene
glycol regions, even on the order of days [Mrksich et al, 1997]. But the primary
advantage to the alkanethiolate SAMs involves the relatively inexpensive
microcontact printing methods used for patterning them onto substrates.
Microcontact printing (gCP) is a fabrication technique that patterns the
formation of alkanethiolate SAMs into designated regions with dimensions
of features down to 1 gm conveniently, and down to 200 nm in special cases
[Xia et al., 1995]. This technique uses an elastomeric stamp to transfer an
alkanethiol to designated regions of a surface of gold. The stamps are usually
fabricated by pouring a prepolymer of polydimethylsiloxane (PDMS) onto a
master relief pattern; this master is often formed by photolithographic
methods, but other sources are available. Because gCP relies on self-assembly,
it does not require a dust-controlled laboratory environment, and can
produce patterned substrates at low cost relative to methods that use
photolithography. I will primarily use this method to generate substrates
containing islands of ECM to control cell shape.
Experimental Design
The literature provides compelling evidence that cell shape per se
could provide the central signal that mediates the regulation of cell
proliferation and apoptosis by adhesion to ECM. Understanding this
regulation in capillary endothelial cells would provide important insights
useful for clinical modulation of angiogenesis, particularly in cancer therapy.
Previous studies have shown that spreading of suspended capillary cells onto
large ECM-coated microcarrier beads prevents suspension-induced apoptosis,
but cells attached to small ECM-coated beads that do not support spreading die
16
=MR
Chapter 1 General Introduction
[Ingber and Folkman, 1989; Re et al., 1993]. While these findings suggest that
cell spreading may provide the survival signal, several factors including bead
curvature, surface area, and internalization confound the results. Cell
spreading also has been implicated in the regulation of proliferation: The
density of ECM adsorbed to a substrate can regulate capillary cell spreading
and growth [Ingber, 1990]. However, it remains unclear whether ECM density
or cell spreading per se is providing the signals for growth. In the present
work, I directly address the hypothesis that ECM regulates growth and
apoptosis through cell shape. The specific goals of this thesis are:
1. To explore methods to control cell shape by engineering surfaces
patterned with islands of ECM. In order to address the question of how cell
shape regulates both apoptosis and growth, I investigated the use of SAMs to
produce islands of ECM that could in turn control endothelial cell shape. By
presenting cells with constant-density, fibronectin-coated islands of varying
size, I could control cell shape independent of the density, surface curvature,
and internalization of the ECM - variables that were previously
uncontrolled.
2. To determine the role of cell shape in ECM-mediated growth and
apoptosis. Using patterned substrates to control cell shape, I determined how
the degree of cell spreading modulated cell growth and apoptosis. We
recognized that by using this approach the cell-ECM contact area (amount of
ECM visible to a cell) increased with cell spreading. To address this issue, cells
were spread across multiple small, focal adhesion-sized islands of ECM such
that, by controlling the size and spacing of the islands, cell spreading could be
varied while maintaining a constant, low area of ECM-cell contact. Thus, the
17
Chapter 1 General Introduction
role of cell shape per se in cell function could be determined. Several
measures for shape were examined in these studies, including projected cell
area, perimeter, and length.
Potential mechanisms for how cell shape might be transduced by the
cell were also explored. Since evidence suggests that FAC may play a central
role in adhesion, integrin signaling, and mechanochemical transduction
[Plopper et al., 1995], the relationship between the amount of FAC formation
and signaling in each cell, spreading, and growth was analyzed. Since
findings also indicate that the shape of the nucleus may directly regulate
growth [Yen and Pardee, 1979; Ingber and Folkman, 1989], I examined changes
in nuclear shape during cell spreading.
3. To define the mechanical basis of force transfer from ECM into the
cell and nucleus. Although ECM regulation of cell growth and apoptosis
appears to be mediated through changes in cell shape, the mechanical basis
for how ECM induces changes in cell shape is unknown. To specifically
examine whether stresses transmit across integrins to the cytoskeletal lattice
or to the cortical membrane, micromanipulation techniques were used to
examine the mechanical connectivity between cell surface integrins, the
cytoskeleton, and the nucleus.
Each of the specific goals is presented in the dissertation as a separate
chapter. The thesis concludes with a general discussion of the findings,
implications, and future applications of this work.
18
Chapter 2 Patterned Surface
CHAPTER II. DESIGN AND FABRICATION OF SUBSTRATES
Preface
While several methods have been used to control cell spreading on
substrates, we chose to explore and develop the use of self-assembled
monolayers (SAMs) for this purpose. In this chapter I describe three distinct
applications of SAMs to engineer ECM environments for studying the role of
adhesion on cell function.
The first approach uses microcontact printing to fabricate islands of
ECM surrounded by nonadhesive regions, such that single cells would attach
and spread on single islands. Microcontact printing, as previously described
[Kumar et al., 1994; Mrksich and Whitesides, 1995], uses elastomeric stamps to
"print" alkanethiolate "inks" onto flat gold substrates in defined regions, such
that the thiol forms a SAM on the gold wherever the patterned stamp
contacts it. The differential adhesivity of the different thiols is then used to
pattern islands of ECM. In this section, I describe how this approach has been
optimized for cell culture, and demonstrate that cell shape can be controlled
with this method.
The second approach elaborates on this stamping method to create
masks of SAMs that act to protect regions of a silicon substrate from chemical
etches that carve contours into the originally flat surfaces, followed by
stamping of SAMs to pattern cells on these substrates; and the third
application describes a new approach to attach cells directly to SAMs of hybrid
thiols that contain a covalently attached Arg-Gly-Asp- (RGD-) containing, cellbinding fragment of fibronectin. Although neither of these approaches were
used to study cell growth and apoptosis in this work, they provide techniques
which will be useful in future studies.
19
Chapter 2 Patterned Surface
PATTERNED SURFACES: Microcontact printing of SAMs
to pattern ECM and cells on flat surfaces
Introduction
Self-assembled monolayers of alkanethiolates on gold have been used
extensively as a model surface in the study of surface interactions of proteins
[Prime and Whitesides, 1993; Prime and Whitesides, 1991; Mrksich et al.,
1995]. The surface properties of these SAMs have been extensively described
[Dubois and Nuzzo, 1992; Whitesides and Gorman, 1995]. Alkanethiolates in
solution freely self-assemble a crystalline-like monolayer onto gold surfaces,
with the sulfur-terminated ends complexed in the nadir between 3 atoms of
gold (111) and the close-packed alkane chains extended into the solvent at a
30' angle. As a result, the attachment of a functional group to the end of the
alkane chain dominates the surface properties of the SAM. Using such
surfaces in adsorption studies have demonstrated that oligo(ethylene glycol)
terminated SAMs completely resist adsorption of proteins [Prime and
Whitesides, 1991], while methyl terminated SAMs promote their
hydrophobic adsorption [Prime and Whitesides, 1993]. Similarly cells cannot
attach to SAMs presenting ethylene glycol moieties, and can only attach to
hydrophobic SAMs if the adsorbed protein is an ECM molecule [Singhvi et al.,
1994; Mrksich et al., 1997].
Fabrication of surfaces containing patterns of different SAMs has been
accomplished using microcontact printing [Kumar et al., 1994; Mrksich and
Whitesides, 1995]. Using this method, one can "stamp" alkanethiolates with
an elastomeric stamp onto particular regions of the gold surface, followed by
immersion of the surface into a second alkanethiolate to form SAMs in the
remaining regions of bare gold. Thus, a substrate patterned with regions of
20
Chapter 2 Patterned Surface
adsorbed ECM molecules surrounded by nonadhesive regions could be
generated by stamping of the hydrophobic hexadecanethiol (HDT),
HS(CH 2 )15 CH 3 , immersing in a tri(ethylene glycol)undecanethiol (EG),
HS(CH2) 11 (OCH 2CH 2 )30H, and immersing in a solution of ECM. Recent
studies have successfully demonstrated the ability of this approach to pattern
cells [Singhvi et al., 1994; Mrksich et al., 1997]. This section describes studies
designed to further characterize and optimize the fabrication of patterned
substrates for use in cell biology studies.
Method Optimization
The conceptual steps involved in the fabrication process of patterned
SAMs have essentially remained unchanged since their proof of concept
(Figure 2.1) [Singhvi et al., 1994]. The execution of the steps following stamp
production* have since been formalized [Mrksich et al., 1997] to improve the
quality of substrates such that the standard protocol produced substrates that
effectively patterned cells from originally less than 50% to now more than
90%. This significant improvement primarily results from using higher
purity EG, immersion of patterns SAM substrates into PBS before adding ECM
protein, and rinsing thoroughly with PBS during the removal of substrates
from the protein solution [Mrksich, personal communication].
Patterned molds for making stamps were photolithographically produced using standard
techniques. Briefly, in a clean room (100), silicon <111> wafers were cleaned, spin coated with
2gm layer of poly methylmethacrylate photoresist, and baked. The wafers were exposed to
high energy UV light through a photolithographic mask containing the desired pattern. The
wafers were developed and washed, leaving 2pm thick photoresist where the UV was masked,
and naked silicon elsewhere. We prepared a poly(dimethylsiloxane) (PDMS) stamp from this
silicon master by polymerizing prepolymer on top of the master. Substrates for cells were then
prepared by evaporation of thin films of titanium (1.5nm) and gold (12nm) on glass cover slips
(0.20 mm, No.2, Corning).
*
21
A
Chapter 2 Patterned Surface
While the quality of substrates were more than adequate for use in our
experiments, several issues emerged as a result of scaling the production up
to more than a few substrates per batch. As the batch size increased, the
quality of substrates decreased, the time to produce them became
burdensome, and the amount of material used in production became costly.
As a result, each step in the production protocol was examined for
improvement. Based on these changes, a new protocol has been established:
Step C (Figure 2.1c). Drying time of HDT in nitrogen stream could be
reduced without any change in pattern quality, from 30 seconds down to the
time it takes for the ethanol to evaporate by visual inspection, or
approximately 5 seconds.
Step D (Figure 2.1d, e). Stamping of HDT could be reduced from 20
seconds to 5 seconds with no apparent difference in FN adsorption as
monitored by immunofluorescent staining. The stamp could be re-used
several times without being re-inked with fresh HDT solution. I stamped
twice before rinsing and re-inking the stamp, because by the 4th stamping,
dust would usually collect on stamp due to static electricity.
Step F (Figure 2.1f). Soaking of EG could be reduced from 12 hours to
30 minutes with no change in ability of the SAM to resist cell adhesion.
However, placing more than 15 substrates (approximately 15 square inches)
into a 15 ml bath of EG led to reduction in the quality of the SAM as
measured by the newly found ability of cells to adhere on the surface. This
quality impingement progressively worsened with each additional substrate
added to the soaking bath. Because the molar amount of EG-thiol in solution
far exceeded the amount of exposed gold surface on the substrates, these
results suggested that the substrates were bringing an adhesive thiol into the
EG-thiol solution, and this mixed-thiol solution in turn would create a
22
Chapter 2 Patterned Surface
partially adhesive SAM in the regions that were meant to be nonadhesive.
To address this issue, and to reduce batch-to-batch variability, every substrate
was place in a separate, fresh solution of EG-thiol. To reduce the volume of
EG-thiol used, substrates were place on a flat surface, and the EG-thiol
solution was dripped directly onto the substrate until it formed a concave
meniscus that covered the entire substrate. Approximately 0.25 ml is used per
square inch of substrate.
Step G (Figure 2 .1g). The use of a 20 ml PBS bath to coat protein at 50
ug/ml onto substrates proved to be the most costly fabrication step
(approximately $50 per bath, for 4 square inches of substrate). It had
previously been feared that placing substrates into a protein solution would
drag the denatured layer of protein at the air-water interface onto the
substrate, overcoating it. I found that inverting substrates and floating them
onto drops of protein solution proved to be equally effective at coating. This
method uses only 0.25 ml of protein solution per square inch of substrate,
lowering the cost by 95%.
Step H. Rinsing of substrates in a PBS stream proved to be important
in maintaining substrate quality, suggesting that the layer of denatured
protein at the air-water interface would be dragged onto the surface of a
substrate being removed from the solution. However, maintaining a stream
of PBS to flow onto an inverted substrate while removing it from the protein
dropped proved technically difficult. I noted that substrates in PBS dewetted
readily, and this dewetting probably played a major role in exposing the
surface to the air-solution interface, thereby overcoating the substrates.
Lowering the surface tension of the coating solution by directly adding 0.5 ml
of 1% bovine serum albumin (BSA) during the "washing" step made it
23
Chapter 2 Patterned Surface
possible to pull the substrate out of the solution without it dewetting. The
substrate could then be placed upright into a cell culture dish for use.
In summary, these changes reduced my stamping time from
approximately 10 substrates per hour to over 30 substrates per hour.
Importantly, the changes also reduced the use of HDT by 50%, EG by 75%, and
protein by 95%.
Different patterns
Cells were patterned onto (1) large regions where many cells attached to
each region, (2) regions on the order of single cells such that single cells
attached to single islands, and (3) regions much smaller than single cells such
that single cells could spread across multiple islands. Masks for patterns were
made either through electron beam photolithography, reduce and step
programs to shrink larger features into small ones, or use of high resolution
laserprinting to create rapid prototypes of slightly lower resolution (Figure
2.2).
Cell culture
Using the newly optimized protocols, we plated bovine and human
capillary cell onto substrates containing patterns of ECM. Phase contrast and
fluorescent microscopy studies demonstrate that cells attach specifically and
precisely to the patterns generated (Figure 2.3, 2.4). When cells were spread
on single islands of ECM, it was found that cell spreading and shape could be
precisely controlled (Figure 2.5, 2.6). Although cells displayed active ruffling
edges at the border of the ECM region, they could not spread into the
surrounding EG-SAM regions (Figure 2.7). Even when cells were spread
across multiple islands of ECM, they only formed adhesions to the ECM
24
Chapter 2 Patterned Surface
regions (Figure 2.8). Finally, the adhesion effects were not unique to FNcoated substrates (see Chapter 3).
Several additional factors in cell culture also appeared to influence the
quality of substrate patterning. In particular, cell plating numbers, serum
levels, and duration of experiment needed to be examined.
It was noted that occasionally cells were able to form "bridges" across
nonadhesive regions. The density of bridges increased with increased plating
of cells, as well as with the addition of serum. Once plated, few de novo
bridges formed during the growth phase of the cultures, suggesting that some
process during the plating caused bridge formation. Time lapse video of cells
during the first 2 hours of plating revealed two independent sources for this
phenomenon. When cell clumps (usually of 4 or more cells) landed across
two adhesive regions, spanning a nonadhesive region, cells would attach and
spread in the two adhesive regions, stretching the cells that were in the
nonadhesive region to form a bridge. When two cells were attached to
adjacent regions, and a fortuitous cell landed between them, it could
sometime form cell-cell junctions with both cells to create a bridge. Cell
clumping could be decreased if cells were used (1) at an earlier passage; (2) less
than 1 week after the previous passage; (3) cell were dissociated in trypsin
longer. Fortuitous cell-cell bridging was reduced by lowering plating density.
The presence of serum did not influence the patterning of capillary
endothelial cells (Figure 2.9). In contrast, 3T3-L1 fibroblasts in the presence of
serum, while preferentially adhering to the ECM-coated regions, easily
moved across the EG regions (Figure 2.10). Even after the initial spreading
phase, the addition of serum (as little as 1%) caused cells to migrate freely into
the EG regions. To test whether this effect was due to direct interactions of
the serum with cells or with the substrate (since serum contains many
25
Chapter 2 Patterned Surface
adhesive factors), substrates were pre-incubated with serum, rinsed, and
presented to 3T3 cells in a defined media. In this experiment, cell patterning
was preserved, indicating that serum was acting directly on cells to potentiate
their ability to adhere to EG regions. Because my thesis work focuses on the
biology of endothelial cells, which (1) can be patterned in the presence of
serum and (2) were studied in reduced or no serum, a mechanistic
explanation of this serum effect was not pursued any further.
Cells were cultured in for up to 10 days with no deleterious effects on
the patterns. This is the upper limit of time cells normally spend in a dish
during a single passage. All experiments conducted in this study were
expected to last less than 48 hours.
In summary, these results demonstrate that SAMs presenting patterns
of ECM can be used to systematically restrict, and hence control, the size
(projected area) and shape of capillary endothelial cells.
26
--
wo
Chapter 2 Contoured Surfaces
CONTOURED SURFACES: Controlling Cell Attachment on
Contoured Surfaces With Self-Assembled Monolayers of
Alkanethiolates on Gold*
Abstract
This section describes a method based on experimentally simple techniques-microcontact printing (pCP) and micromolding in capillaries (MIMIC)--to
prepare tissue culture substrates in which both the topology and molecular
structure of the interface can be controlled. The method combines optically
transparent, contoured surfaces with self-assembled monolayers (SAMs) of
alkanethiolates on gold to control interfacial characteristics; these tailored
interfaces, in turn, control the adsorption of proteins and the attachment of
cells. The technique uses replica molding in poly(dimethylsiloxane) (PDMS)
molds having micron-scale relief patterns on their surfaces to form a
contoured film of polyurethane supported on a glass slide. Evaporation of a
thin (<12 nm) film of gold on this surface-contoured polyurethane provides
an optically transparent substrate, on which SAMs of terminallyfunctionalized alkanethiolates can be formed. In one procedure, a flat PDMS
stamp was used to form a SAM of hexadecanethiolate on the raised plateaus
of the contoured surface by contact printing hexadecanethiol (HS(CH 2 )15 CH3 );
a SAM terminated in tri(ethylene glycol) groups was subsequently formed on
the bare gold remaining in the grooves by immersing the substrate in a
solution of a second alkanethiol (HS(CH 2 )1 1 (OCH 2 CH 2 )3 0H). When this
patterned substrate was immersed in a solution of fibronectin, the protein
* Contributing authors for publication in Proceedings of the National Academy of Sciences,
USA: Milan Mrksich, Christopher S. Chen, Younan Xia, Laura E. Dike, Donald E. Ingber, and
George M. Whitesides. CC carried out all the experiments in this study, under the supervision
of MM and with consultations from YX and LD.
27
Chapter 2 Contoured Surfaces
adsorbed only on the methyl-terminated, plateau regions of the substrate (the
tri(ethylene glycol)-terminated regions resisted the adsorption of protein);
bovine capillary endothelial cells attached only on the regions that adsorbed
fibronectin. A complementary procedure confined protein adsorption and
cell attachment to the grooves in this substrate.
28
Chapter 2 Contoured Surfaces
This report describes a simple and general method to fabricate optically
transparent surfaces contoured into grooves of defined size and shape, and to
use self-assembled monolayers (SAMs) of alkanethiolates on gold to control
cell attachment to these substrates. We have used SAMs extensively to
control the adsorption of proteins and the attachment of mammalian cells to
planar surfaces [Prime and Whitesides, 1991; Prime and Whitesides, 1993;
Mrksich et al., 1995; DiMilla et al., 1994; Lopez et al., 1993; for pioneering work
by other groups, see references Kleinfeld et al., 1988; O'Neill et al., 1990;
Britland et al., 1992; Stenger et al., 1992; Spargo et al., 1994]. By patterning the
formation of SAMs using microcontact printing (gCP) [Kumar et al., 1994;
Mrksich and Whitesides, 1995]--an experimentally simple and nonphotolithographic technique--into regions that promote or resist the
adsorption of protein, the attachment of cells to surfaces could be confined to
rows 10-100 pm in width [Mrksich et al., 1997], or to islands, for the
attachment of single cells [Singhvi et al., 1994]. The present work extends this
methodology to include control over the topography of surfaces used for cell
culture; the method employs an elastomeric stamp having micron-scale
patterns of relief to mold a thin film of polyurethane, and SAMs to control
the properties of these contoured surfaces.
A number of groups have used contoured surfaces to study the effects
of topography on cell alignment, migration, and metabolism [Chou et al.,
1995; Clark et al., 1991; Meyle et al., 1994; Hoch et al., 1987; Green et al., 1994;
Schmidt and con Recum et al., 1992]; this work has demonstrated the
importance of substrate topography in controlling the behavior of cells. The
procedures used to fabricate the substrates used in these studies have three
limitations: (i) The molecular properties of the surfaces are not well-
29
Chapter 2 Contoured Surfaces
controlled (nor can these properties be tailored easily); (ii) The substrates
(silicon) are optically opaque, and attached cells cannot be visualized using
conventional light microscopy; (iii) The preparation of the substrates require
photolithographic techniques that are not routinely available in biological
laboratories. The methodology described in this report uses more flexible and
convenient techniques for microfabrication--microcontact printing (pCP)
[Kumar et al., 1994; Mrksich and Whitesides, 1995] and micromolding in
capillary channels (MIMIC) [Kim et al., 1995]--to create substrates contoured
into grooves and plateaus. The methodology is general in that it allows
surfaces having a variety of topologies to be fabricated easily, and it permits
control at the molecular scale over the interfacial properties of the substrates.
Using SAMs to Control the Properties of a Surface. SAMs of
alkanethiolates on gold are prepared by immersing a substrate coated with a
thin film of gold in an ethanolic solution of a long-chain alkanethiol
(HS(CH2 )nX, 10 < n < 25). The sulfur atoms coordinate to the gold, and the
trans-extended alkyl chains pack tightly: the terminal group, X, is confined to
the interface between the SAM and the aqueous phase; the properties of the
interface are dominated by the identity of this group [Whitesides and
Gorman, 1995; Dubois and Nuzzo, 1992; Mrksich and Whitesides, 1997].
For studies involving the attachment of cells, we have used glass slides
coated with thin, optically-transparent films of gold (10-12 nm) (4). SAMs
terminated in methyl groups are hydrophobic and adsorb protein quickly and
irreversibly from solution. SAMs terminated in short oligomers of the
ethylene glycol group (-S(CH 2 )11(OCH2CH2)nOH, n=2-7) resist essentially
completely the non-specific adsorption of proteins; in situ, these SAMs resist
even the adsorption of "sticky" proteins such as fibrinogen (3). For the same
reason, SAMs terminated in oligo(ethylene glycol) groups resist the
30
Chapter 2 Contoured Surfaces
attachment of cells--and the spreading of attached cells--over periods of
several days in culture [Mrksich et al, 1997; Singhvi et al., 1994].
Materials and Methods
Materials Used in Fabrication. Poly(dimethylsiloxane) (PDMS) was
purchased from Dow Corning (Sylgard 184). PDMS stamps were prepared
from photolithographically produced masters as described previously [Kumar
et al., 1994]; flat stamps were prepared by casting the prepolymer against a
clean silicon wafer [Jeon et al., 1995]. Silicon wafers were purchased from
Silicon Sense (3", <111> orientation). Prepolyurethane (Norland Optical
Adhesive 68) was purchased from Norland Products Inc (New Brunswick,
N.J.). Hexadecanethiol was purchased from Aldrich and purified by silica gel
chromatography using 19:1 hexanes:ethyl acetate as the eluent. The
tri(ethylene glycol)-terminated alkanethiol was synthesized as described
previously [Pale-Grosdemange et al., 1991]. All other chemicals and solvents
were purchased from Aldrich and used as received.
Attachment of Cells to Substrates. The contoured substrates were
placed in Petri dishes containing phosphate-buffered saline (PBS; 20 mL; 10
mM phosphate, 100 mM sodium chloride, pH=7.4). A solution of fibronectin
(Organon Teknika-Cappel, Melvern PA) in PBS (400 pL; 2.5 mg/mL) was
added. After 2 hr, the solution was diluted by the addition of PBS (-200 mL)
and the substrates were removed from solution under a stream of buffer and
transferred immediately to Petri dishes containing defined media (low
%
glucose Dulbecco's modified eagle medium (DMEM), 10 mM Hepes, 1
bovine serum albumin (BSA), 10 gg/mL high-density lipoprotein (HDL), 10
gg/mL transferrin, 5 gg/mL fibroblast growth factor (FGF)). Bovine capillary
endothelial (BCE) cells were plated on these substrates and maintained in
31
Chapter 2 Contoured Surfaces
culture for several days (37 'C, 10 % C0 2 ) [Ingber and Folkman, 1989]; the
medium was initially exchanged 2 hr after inoculation with cells, and daily
thereafter. After three days, the cells were fixed with paraformaldehyde and
either stained for F-actin using rhodaminated-phalloidin (Sigma) or sputtered
with gold and observed by scanning electron microscopy (SEM).
Results and Discussion
Fabrication of Substrates. Our method for fabrication of contoured
substrates involved four steps (Figure 2.11): (i) Preparation of a master
pattern in silicon by micromolding in capillaries (MIMIC) using an
elastomeric stamp, followed by anisotropic chemical etching of the silicon
(other procedures would also work): (ii) Transfer of the topographical
pattern into a film of polyurethane on a glass coverslip: (iii) Evaporation of a
thin, optically transparent film of gold on the polyurethane: (iv) Formation
of patterns of SAMs of alkanethiolates on the gold.
To accomplish the first step, we prepared a poly(dimethylsiloxane)
(PDMS) stamp using the procedure described for gCP [Kumar et al., 1994]. The
stamp was placed on a silicon <100> wafer having a layer of silicon dioxide;
the recessed features of the stamp formed a network of channels (Figure 2.11).
When a drop of prepolyurethane was placed on the wafer and in contact with
the stamp, capillary action caused the liquid to fill the channels completely
(a). The prepolymer was cured with UV light and the stamp was removed
from the surface to leave a pattern of the polymer at the surface (b). This
polymer protected the underlying Si0 2 from dissolution in an aqueous
solution of HF (1%); the exposed regions of silicon were then etched
anisotropically in an aqueous solution of KOH (4 M, 15 % isopropanol, 60 C)
to give V-shaped grooves (c) [Kim et al., 1995]. A PDMS stamp was cast from
32
Chapter 2 Contoured Surfaces
this substrate (d), peeled away (e), and gently pressed onto a drop of liquid
prepolyurethane on a glass coverslip (f). The structure was cured under UV
light with the stamp in place, and the stamp was then peeled away to give the
contoured substrate (g). Figure 2.12 shows a scanning electron micrograph of
this fabricated substrate. This same PDMS stamp could be used to fabricate
multiple substrates.
Evaporation of a thin layer of titanium (1.5 nm; to promote adhesion
of the gold to the polyurethane) and a thin layer of gold (12 nm) provided a
contoured substrate (h) on which SAMs could be assembled. In one example,
the plateaus of the substrate were derivatized selectively with a SAM of
hexadecanethiolate by contact printing with a flat stamp [Jeon et al., 1995] (i);
this procedure left the gold surface of the grooves unmodified (j). A SAM
terminated in tri(ethylene glycol) groups was formed in the grooves by
immersing the substrate in a solution of the second alkanethiol
(HS(CH 2 )11 (OCH2 CH 2 )3 0H) (k). Substrates having a reversed pattern of SAM
were prepared by first printing the tri(ethylene glycol)-terminated alkanethiol
onto the plateaus, and then immersing in a solution of hexadecanethiol*.
Directed Attachment of Cells. We examined the attachment of bovine
capillary endothelial (BCE) cells on two fibronectin-coated contoured surfaces;
one having ridges 25 pm in width and separated by V-shaped trenches of
equal width, and the second having ridges and grooves 50 pm in width. For
all substrates, the attachment of cells depended strictly on the properties of the
SAM and not on the topology of the substrate; SAMs presenting tri(ethylene
glycol) groups resisted the adsorption of fibronectin and the subsequent
* Ellipsometric measurements showed that microcontact printing of the tri(ethylene glycol)terminated alkanethiol resulted in < 50% formation of SAM. We determined empirically that
it was necessary to repeat the microcontact printing three times before immersing the substrate
in a solution of hexadecanethiol to passivate the ridges of the contoured substrates.
33
Chapter 2 Contoured Surfaces
attachment of BCE cells; fibronectin adsorbed to methyl-terminated SAMs,
and allowed efficient attachment of cells in these areas. Substrates modified
uniformly with a SAM of hexadecanethiolate presented fibronectin at all
regions and allowed efficient attachment of the BCE cells on both the plateaus
and grooves, with little preference for either region (Figure 2.13a). For
substrates presenting fibronectin only on their plateaus, cells attached
exclusively to the plateaus; no cells attached to the grooves presenting a SAM
of tri(ethylene glycol) groups (Figure 2.13b). For substrates whose grooves
were coated with fibronectin, cells attached only to the sides of the grooves:
many cells stretched across both sides of the grooves without contacting the
bottom edge (Figure 2.13c).
These contoured substrates have many characteristics that make them
useful for experimental manipulation of cultured cells. Because the
substrates are optically transparent, attached cells can be observed in culture
using standard light microscopy. Figure 2.14 shows optical micrographs of
cells that were stained with Comassie Blue; cells were also visible by phase
contrast without staining.
The gold-coated substrates are compatible with fluorescence
microscopy. Figure 2.14c shows a fluorescent micrograph of the F-actin
network of cells confined to ridges after staining with rhodaminatedphalloidin. These substrates also have the stability required for use in cell
culture. After a period of five days, the BCE cells remained attached to the
contoured substrates and continued to divide; the cells also did not invade
regions that were modified with a SAM terminated in tri(ethylene glycol)
groups.
SAMs of alkanethiolates in this methodology provide many
opportunities for tailoring the molecular structures of the surfaces to control
34
Chapter 2 Contoured Surfaces
their interfacial characteristics. For example, the properties of SAMs that
present electroactive groups can be switched by applying a potential to the
gold substrate [Abbott and Whitesides, 1994; Wong et al., 1994]; the thin,
optically transparent films of gold used here still have the electrical
conductivity of bulk gold [Gorman et al., 1995]. SAMs that present ligands of
low molecular weight have been prepared for fundamental studies of biospecific adsorption of proteins at interfaces [Mrksich et al., 1995]. SAMs
presenting chelates of Ni(II) are useful for immobilizing his-tagged proteins
from cell extracts [Sigal et al., 1996]. A variety of analytical techniques--surface
plasmon resonance (SPR) spectroscopy [Mrksich et al., 1995; Mrksich et al.,
1995; Sigal et al., 1996], ellipsometry [Prime and Whitesides, 1993; Prime and
Whitesides, 1991], scanning electron microscopy [Lopez et al., 1993], and quartz
crystal microbalance [Ward and Buttry, 1990]--can be used to study the
interactions of proteins with SAMs on gold. SPR is especially useful because
it is a non-invasive technique that can detect ~2% of a monolayer of protein,
and it provides both kinetic and thermodynamic parameters.
In summary, this report describes a flexible methodology to prepare
optically transparent, contoured surfaces appropriate for fundamental studies
of the relationships between the molecular structure and topology of a surface
and the behavior of cells attached to the surface. This experimental system
may also find use in applied cell culture, including the development of
supports for the immobilization of cells in bioreactors, and substrates for
tissue engineering. The range of geometries of features that can be formed is
limited only by the availability of appropriate master templates; these
templates are often created using techniques common in microfabrication,
but are also available from other sources (e.g. diffraction gratings). This
methodology can be used to prepare contoured substrates having features
35
Chapter 2 Contoured Surfaces
with dimensions down to the sub-micron range without requiring access to
the special facilities and instrumentation used currently in microfabrication
[Wilbur et al., 1995].
Acknowledgments.
This work was supported by the National Institutes of
Health (GM 30367 to G.M.W. and CA 55833 to D.E.I.), the Office of Naval
Resarch, and the Advanced Research Projects Agency. D.E.I. is a recipient of a
Faculty Research Award from the American Cancer Society. M. M. is grateful
to the American Cancer Society for a postdoctoral fellowship.
36
Chapter 2 Biospecific Surfaces
BIOSPECIFIC SURFACES: Direct attachment and spreading of
cells to mixed self-assembled monolayers presenting GRGD and
(EG) 3 0H groups*
ABSTRACT
This paper describes the interactions of cells and proteins with self-assembled
monolayers (SAMs) of alkanethiolates on gold that present mixtures of
glycine-arginine-glycine-aspartate (GRGD), a tetrapeptide that promotes cell
adhesion by binding to cell surface integrin receptors, and
oligo(ethyleneglycol) moieties, groups that resist non-biospecific adsorption of
proteins. Surface plasmon resonance (SPR) spectroscopy was used to measure
the adsorption of carbonic anhydrase and fibrinogen to mixed SAMs
comprising trityl groups (EG 60GRGD) and oligo(ethylene glycol) groups
(EG 30H); SAMs having values of the mole fraction of GRGD (XGRGD)
0.05 do
not adsorb carbonic anhydrase or fibrinogen. Bovine capillary endothelial
cells attached and spread on SAMs at XGRGD
0.00001, with spreading of cells
reaching a maximum at XGRGD > 0.001. These mixed SAMs prevented the
deposition of proteins by attached cells relative to both fibronectin-coated
SAMs of hexadecanethiolate and RGD peptide-coated glass. After allowing
cells to attach for 2 or 4 h onto surfaces presenting RGD, addition of soluble
GRGDSP to the adherent cells rapidly released them from the surfaces. In
contrast, if cells were allowed to attach onto surfaces for 24 h, only cells
attached to the mixed SAM surface could be released using the soluble
GRGDSP. These results demonstrate that the integrin-RGD interaction alone
is sufficient for adhesion and survival of cells over 24 h.
* Contributing authors for publication in Journal of the American Chemical Society: Carmichael
Roberts, Christopher S. Chen, Milan Mrksich, Valerie Martichonok, Donald E. Ingber and
George M. Whitesides. CC performed all cell culture experiments in this section. CR, MM, and
VM synthesized the new thiols and CR aided in all experiments.
37
Chapter 2 Biospecific Surfaces
INTRODUCTION
Adhesion of cells to the extracellular matrix (ECM) influences the
shape, growth, viability, differentiation, migration, and metabolism of these
cells [Ingber, 1990; Boudreau et al., 1995; Watt et al., 1988; Flaumenhaft and
Rifkin, 1991; Basson et al., 1992; Salomon et al., 1981]. However, it has been
difficult to characterize the biological activities of specific constituents of the
ECM (e.g., fibronectin, laminin, vitronectin, collagens, and proteoglycans),
primarily because within hours after plating cells onto substrates presenting
specific ECM proteins, cells can degrade and redeposit a new ECM. We
believe that the combination of mixed self-assembled monolayers (SAMs)
that present specific ECM moieties with an "inert", non-adsorbing interface
could provide a surface technology that would promote attachment through
specific cell adhesion receptors while preventing the remodeling of the
substrate. Here, we used mixed SAMs and SPR to study attachment and
spreading of bovine capillary endothelial cells on GRGD presented in a
background of (EG) 30H groups that resists the deposition of extracellular
matrix by the cell (Figure 2.15).
To promote cell adhesion mediated by specific interactions with cellsurface adhesion receptors, ECM proteins or peptide fragments are often
immobilized onto the surfaces of biomedical materials either by nonspecific
adsorption onto hydrophobic surfaces or by nonspecific covalent attachment
onto chemically reactive substrates [Ingber, 1990; Massia and Hubbell, 1991;
Stenn et al., 1983; Ohji et al., 1993; Aznavoorian et al., 1990; Rannels et al.,
1992]. Surfaces prepared using these procedures are heterogenous and poorly
characterized; it is impractical, in these systems, to count or control the
number of ligands that are biologically functional. Surfaces fabricated to date
also have not been shown to resist the deposition of additional adhesive
38
Chapter 2 Biospecific Surfaces
ligands expressed by the attached cells; within hours, the molecular
composition of the surface, and therefore the spectrum of cell surface
receptors being engaged, change uncontrollably. Although the use of protein
synthesis inhibitors to block synthesis and deposition of new ECM proteins
may be useful for a few hours [Aznavoorian et al., 1990], this approach
severely impairs many processes within cells and is lethal to them within 8 to
24 hours [Lewis et al., 1995; Lor et al., 19941. As a result it is difficult to assess
the effects of specific ligand-cell interactions on many biological processes,
including adhesion itself.
Cell attachment to the RGD peptide, found in many proteins of the
ECM, through specific cell-surface integrin receptors has been described well
by Ruoslahti and Pierschbacher [1987]. Using SAMs of alkanethiolates on
gold, we have previously demonstrated that surfaces presenting
oligo(ethylene glycol) moieties prevent the adsorption of protein [PaleGrosdemange et al., 1991; Prime and Whitesides, 1991; Prime and Whitesides,
1993]. In this study we develop surfaces that promote cell attachment by the
specific interaction of RGD with cell surface integrin receptors, and resist
significant deposition of cell-derived matrix components. Using this model
system, we demonstrate that GRGD alone is sufficient to maintain long term
biospecific attachment and survival of cells.
EXPERIMENTAL PROCEDURE
Synthesis of Alkanethiol 1 (see Figure 2.16).
Materials and Methods: Reactions were monitored by thin layer
chromatography (TLC) using 0.25-mm silica gel plates (E. Merck).
39
Chapter 2 Biospecific Surfaces
Column chromatography was performed using silica gel-60 (particle
size 0.040-0.063) (E. Merck). All reactions in non-aqueous solvents were
executed under nitrogen.
Z-NHGR(PMC)-OH (2b). To a solution Z-NH-G-ONHS (3.81 g, 12.4
mmol) in DMF (40 mL) cooled to 0 'C was added H 2 NR(PMC)-OH 2a (5.0 g,
11.3 mmol) followed by dropwise addition of diisopropylethylamine (DIPEA)
(7.48 mL, 43 mmol). The reaction mixture was stirred at 0 'C for 1 h, allowed
to warm to rt and stirred at rt for an additional 3 h. Product 2b precipitated
from cold H 20 after acidification with 1N aqueous HCl (26 mL). The obtained
precipitate was filtered, washed with cold H 2 0 and dried in vacuo to afford 2b
(6.7 g, 94%).
1H NMR (500 MHz, CDCl 3 ): 8 1.27 (s, 6 H), 1.55 (bs, 2 H), 1.75 (m,
3 H),1.85 (bs, 1 H), 2.06 (s, 3 H), 2.49 ( s, 3 H), 2.50 (s, 3 H), 2.56 (bs, 2 H), 3.14 (bs, 2
H), 3.89 (bs, 2 H), 4.48 (bs, 1 H), 5.00 (s, 2 H), 6.15-6.50 (m, 4 H), 7.24 (m, 5 H), 7.61
(bs, 1 H).
Z-NHGD(OtBu)-OtBu (3b). To a solution of Z-NH-G-ONHS (6.87 g, 22.4
mmol) in DMF (40 mL) cooled to 0 'C was added H 2ND(OtBu)-OtBu 3a (5.0 g,
20.4 mmol) followed by dropwise addition of DIPEA (7.48 mL, 43 mmol). The
reaction mixture was stirred at 0 'C for 1 h, allowed to warm to rt and stirred
at rt for an additional 3 h. The reaction mixture was added dropwise to cold
H 20, and left on ice for 2 h. The obtained precipitate was filtered, washed
with cold H 2 0 and dried in vacuo to afford 3b (7.3 g, 82%).
H NMR (400
MHz, CDCl 3 ): 8 1.41 (s, 9 H), 1.43 (s, 9 H), 2.71 (dd, J= 3.48, 17.11 Hz, 1H), 2.88
(dd, J = 4.17, 17.11 Hz, 1H), 3.92 (m, 2 H), 4.68 (dt, J= 4.29, 8.36 Hz, 1H), 5.12 (s,
2 H), 5.38 (bs, 1 H), 6.85 (d, J = 7.90 Hz, 1H), 7.29-7.36 (m, 5 H).
40
Chapter 2 Biospecific Surfaces
Z-NHGR(PMC)GD(OtBu)-OtBu (4). Compound 3b (1.46 g, 3.35 mmol)
was hydrogenated in EtOH (30 mL) over 10% Pd/C (0.3 g) until the TLC (33%
EtOAc in hexanes) indicated that 3b had been consumed. The reaction
mixture was filtered through Celite and concentrated in vacuo to give the
crude amine that was used in the next step without further purification. The
flask containing crude amine from 3b was purged with N 2 , acid 2b (2.3 g, 3.65
mmol) and dry DMF (20 mL) were added and the stirred solution was cooled
to 0 0C. Diphenylphosphoryl azide (DPPA) (0.94 mL, 4.38 mmol) was added,
followed by a solution of DIPEA (0.76 mL, 4.38 mmol) in DMF (5 mL) and the
stirring was continued at 0 0C for 10 h. The mixture was diluted with EtOAc
(100 mL) and washed successively with H 2 0 (3 x 20 mL), 5% aqueous NaHCO 3
(20 mL) and brine (2 x 20 mL). The organic phase was dried (MgSO 4) and the
solvent was removed in vacuo to give the residue which was
chromatographed (eluting with 10% EtOH in EtOAc) to give product 4 (2.23 g,
71%).
H NMR (500 MHz, CDCl 3 ): 6 1.26 (s, 6 H), 1.36 (s, 9 H), 1.38 (s, 9 H), 1.54
(m, 2 H), 1.63 (m, 2 H), 1.75 (t, J = 6.78 Hz, 2 H), 1.85 (m, 2 H), 2.05 (s, 3 H), 2.50
(s, 3 H), 2.51 (s, 3 H), 2.57 (t, J = 6.72 Hz 2 H), 2.65 (dd, J = 3.36, 17.00 Hz, 1 H),
2.78 (dd, J = 4.20, 17.00 Hz, 1 H), 3.18 (bs, 2 H), 3.81-4.01 (m, 4 H), 4.50 (m, 1 H),
4.63 (m, 1 H), 5.03 (s, 2 H), 6.10-6.40 (m, 4 H), 7.25 (m, 5 H), 7.43 (m, 1 H), 7.77
(m, 1 H). HRMS (FAB) calcd for C 44 H 65 N 7 0 12 SNa (M + Na) 938.4310, found
938.4310.
CH2=CH-(CH 2)9-(OCH 2CH 2)6-OCH 2COO-tBu (6). To a solution of
alcohol 51718 (1.04 g, 2.4 mmol) in dry DMF (5 mL) cooled to 0 *C was added
NaH (144 mg of 60% suspension in oil, 3.6 mmol). The mixture was stirred at
0 'C for 10 min, t-butylbromoacetate (532 gL, 3.6 mmol) was added in one
portion, and the mixture was allowed to warm up to rt. After stirring for 6 h
41
Chapter 2 Biospecific Surfaces
additional t-butylbromoacetate (532 gL, 3.6 mmol) was added and the stirring
was continued at 40 'C for 10 h. After cooling to rt EtOAc (50 mL) was added,
organic phase was washed with H 20, brine, dried (MgSO 4 ) and concentrated in
vacuo. Column chromatography (3% MeOH in CH 2Cl 2 ) afforded product 6
(0.81 g, 62%). 1H NMR (400 MHz, CDCl 3 ):
8
1.25 (bs, 10 H), 1.29-1.41 (m, 2 H),
1.45 (s, 9 H), 1.55 (m, 2 H), 2.01 (m, 2 H), 3.42 (t, J = 6.80 Hz, 2H), 3.56 (m, 2 H),
3.60-3.73 (m, 22 H), 4.00 (s, 2 H), 4.89 (m, 2 H), 5.73-5.84 (m, 1 H). MS (FAB)
calcd for C2 9 H 5 6 O9 Na (M + Na) 571, found 571.
CH2=CH-(CH 2 )9-(OCH 2CH 2)-OCH 2C(O)NHGR(PMC)GD(OtBu)-OtBu (8).
To a solution of 6 (376 mg, 0.69 mmol) in CH 2 Cl 2 (5mL) was added TFA (5 mL)
and the mixture was stirred for 3 h. After concentration in vacuo and
-
column chromatography (10% MeOH in CH 2Cl 2 ) CH 2=CH-(CH 2)9-(OCH 2CH 2)6
OCH 2COOH (290 mg, 0.59 mmol, 86%) was obtained.
Z-protected tetrapeptide
4 (507 mg, 0.65 mmol) was hydrogenated in EtOH (10 mL) over 10% Pd/C (0.1
g). After filtration and concentration in vacuo the crude amine was obtained.
The amine and CH 2=CH-(CH 2)9-(OCH 2CH 2)6-OCH2COOH 7 were combined,
the flask was purged with N2 , dry DMF (3 mL) was added and the stirred
solution was cooled to 0 0C. DPPA (150 gL, 4.38 mmol) was added, followed
by a solution of DIPEA (120 kL, 4.38 mmol) in DMF (1 mL) and the stirring
was continued at 0 'C for 10 h. The mixture was diluted with EtOAc (10 mL)
and washed successively with H 2 0, 5% aqueous NaHCO 3 and brine. The
organic phase was dried (MgSO 4) and the solvent was removed in vacuo to
give a residue which was chromatographed (5% MeOH in CH 2Cl 2 -+ 10%
MeOH in CH 2 Cl 2 ) to give product 8 (355 mg, 48%).
42
H NMR (500 MHz,
Chapter 2 Biospecific Surfaces
CDC13): 8 1.25 (s, 10 H), 1.28 (s, 6 H), 1.30-1.36 (m, 2 H), 1.39 (s, 9 H), 1.40 (s, 9 H),
1.48-1.63 (m, 4 H), 1.68-1.75 (m, 1 H), 1.78 (t, j = 6.80 Hz, 2 H), 1.89-1.95 (m, 1 H),
2.01 (q, J = 6.85 Hz, 2 H), 2.07 (s, 3 H), 2.53 (s, 3 H), 2.55 (s, 3 H), 2.60 (t, J = 6.70
Hz, 2 H), 2.67 (dd, j = 4.75, 16.94 Hz, 1 H), 2.79 (dd, J = 5.01, 16.99 Hz, 1 H), 3.22
(m, 2 H), 3.40 (t, J = 6.85 Hz, 2 H), 3.52-3.55 (m, 2 H), 3.58-3.64 (m, 22 H), 3.663.71 (m, 2 H), 3.88 (dd, J = 5.68, 16.62 Hz, 1 H), 3.95 (dd, J = 5.64, 14.47 Hz, 1 H),
3.98-4.05 (m, 2 H), 4.01 (s, 2 H), 4.46 (q, J = 7.61 Hz, 1 H), 4.63 (dt, J= 4.86, 8.04 Hz,
1 H), 4.88-5.00 (m, 2 H), 5.73-5.83 (m, 1 H), 6.36 (bs, 2 H), 7.11 (d, J= 8.06 Hz, 1
H), 7.43 (bs, 1 H), 7.63 (bt, J = 5.29 Hz, 1 H), 7.90 (t, J = 5.76 Hz, 1 H). MS (FAB)
calcd for C61 H 105N 7 0 18SNa (M + Na) 1278, found 1278.
AcS-(CH 2)11-(OCH 2CH2)6-OCH 2C(O)NHGR(PMC)GD(OtBu)-OtBu (9). A
mixture of 8 (650 mg, 0.52 mmol), thioacetic acid (742 pL, 10.4 mmol), and
AIBN (20 mg) in THF (10 mL) was irradiated by UV. After 2 h additional
AIBN (20 mg) was added and irradiation was continued for 3 h. The mixture
was concentrated in vacuo and chromatographed (5% MeOH in CH 2Cl 2 ->
10% MeOH in CH2Cl 2) to give thioacetate 9 (641 mg, 93%). 1H NMR (500
MHz, CDCl 3 ): 6 1.26 (bs, 12 H), 1.28 (s, 6 H), 1.28-1.33 (m, 2 H), 1.39 (s, 9 H), 1.40
(s, 9 H), 1.47-1.64 (m, 6 H), 1.68-1.75 (m, 1 H), 1.78 (t, j= 6.80 Hz, 2 H), 1.88-1.96
(m, 1 H), 2.07 (s, 3 H), 2.28 (s, 3 H), 2.53 (s, 3 H), 2.54 (s, 3 H), 2.60 (t, J = 6.68 Hz, 2
H), 2.68 (dd, J = 4.80, 16.94 Hz, 1 H), 2.78 (dd, j= 5.10, 16.96 Hz, 1 H), 2.82 (t, J=
7.36, 2 H), 3.23 (m, 2 H), 3.40 (t, j= 6.85 Hz, 2 H), 3.52-3.55 (m, 2 H), 3.58-3.64 (m,
22 H), 3.66-3.71 (m, 2 H), 3.87 (dd, J = 5.70, 16.64 Hz, 1 H), 3.95 (dd, j= 5.68, 14.48
Hz, 1 H), 3.98-4.05 (m, 2 H), 4.01 (s, 2 H), 4.45 (bq, J= 7.52 Hz, 1 H), 4.63 (dt, j=
4.90, 8.18 Hz, 1 H), 6.36 (bs, 2 H), 7.14 (d, J = 8.07 Hz, 1 H), 7.52 (bs, 1 H), 7.71 (bs,
43
Chapter 2 Biospecific Surfaces
1 H), 7.94 (t, J = 5.78Hz, 1 H). MS (FAB) calcd for C63 H10 9 N 7 0 19 S2Na (M + Na)
.
1354, found 1354
HS-(CH 2)11-(OCH 2CH 2)6 -OCH 2C(O)NHGRGD-OH (1). A mixture of
CF 3COOH/PhSMe/HS(CH2) 2SH/H20 37:1:1:1 (v/v) was used for cleavage of
the protective groups in 9. A solution of 9 (320 mg, 0.25 mmol) in CH 2Cl 2 (2
mL) was cooled 0 'C and the cleavage mixture (10 mL) was added. The
obtained solution was stirred at 0 'C for 1 h, allowed to warm to rt and stirred
for 2 h at rt. The mixture was concentrated in vacuo, CH 2Cl 2 (10 mL) was
added and the mixture was concentrated in vacuo again. The residue was
dissolved in a minimum amount of CH 2Cl 2, cold Et2O ( 50 mL) was added
dropwise into the flask with vigorous stirring, the obtained precipitate was
-
filtered and dried in vacuo to afford AcS-(CH 2)11-(OCH 2 CH 2)6
OCH 2C(O)NHGRGD-OH (195 mg, 0.20 mmol, 80%) as a white solid. MS (FAB)
calcd for C 41H 75N 70 16 SNa (M + Na) 976, found 976. The obtained thioacetate
was dissolved in absolute MeOH (10 mL) and the solution was cooled 0 'C.
NaOMe (1.54 mL of 0.39 M in MeOH, 0.60 mmol) was added and the mixture
was stirred at 0 'C for 3 h. The mixture was neutralized with Dowex exchange
resin (H-form), filtered and concentrated in vacuo. Repeated precipitation (3
times) from CH 2Cl 2 with cold Et2 O afforded pure 1 (63 mg, 34%).
1H
NMR
(500 MHz, CD 3 0D): 8 1.27-1.46 (m, 12 H), 1.52-1.61 (m, 4 H), 1.63-1.71 (m, 2 H),
1.72-1.81 (m, 2 H), 1.92-2.00 (m, 2 H), 2.48 (t, J = 7.15 Hz, 2 H), 2.82 (t, J = 5.41 Hz,
2 H), 3.20 (t, J = 6.76 Hz, 2 H), 3.46 (t, J = 6.62 Hz, 2 H), 3.57 (m, 2 H), 3.61-3.66 (m,
18 H), 3.68 (m, 2 H), 3.71 (m, 2 H), 3.84-4.02 (m, 4 H), 4.06 (s, 2 H), 4.40 (t, J = 7.32
Hz, 1 H), 4.63 (t, J = 5.50 Hz, 1 H);
13C
NMR (125 MHz, CDCl 3): 8 24.97, 25.94,
27.20, 29.41, 30.22, 30.57,30.65, 30.72, 35.22, 37.62, 42.02, 43.13, 43.39, 50.77, 54.33,
71.14, 71.19, 71.33, 71.43, 71.47, 71.52, 72.08, 72.37, 158.63, 171.06, 171.58, 173.74,
44
Chapter 2 Biospecific Surfaces
174.17, 174.92, 175.02. HRMS (FAB) calcd for C 39 H 73N 7 01 5 SNa (M + Na)
934.4783, found 934.4805.
Preparation of SAMs: Substrates were prepared as previously
described[Prime and Whitesides, 1991; Prime and Whitesides, 1993],
begining with evaporation of titanium (1 nm) and gold onto glass
slides (38 nm of Au for SPR experiments; and 12 nm of Au for cell
culture)*. The slides were immersed in ethanolic solutions containing
mixtures of HSC 1 1EG 6 0GRGD and HSC 11EG 3 0H (2 mM total thiol) in
ethanol for 4h. Ellipsometric measurements of mixed SAMs of
HSC 11 EG 6 0GRGD and HSC11EG 3 0H resulted in thicknesses ranging
from 22.8 to 23.7 A; these values are in good agreement with those
expected for a well-packed SAM containing trans-extended
alkanethiolates [Prime and Whitesides, 1993]. The relative amount of
GRGD, XGRGD, is based on the mole fractions of EG6 0GRGD and EG 30H
in solution (Equation 1); therefore surface values may differ
systematically**.
The SPR spectroscopy requires a reflective gold surface (38 nm of Au). In contrast, a
transparent surface (12 nm of Au) is required for the study of cells.
** Prime and Whitesides [1991; 1993] found that the thickness of mixed SAMs of EGn
alkanethiolates increases almost linearly with composition.
45
Chapter 2 Biospecific Surfaces
%GRGD
=
[EG6GRGD](1)
[EG6OGRGD]+[EG30H]
Fibronectin (FN) coated substrates, used as a positive control for
adhesion assays, were prepared by immersing metallized substrates in
hexadecanethiol (2 mM) in ethanol for 4h; then, coated substrates were
placed in a solution of fibronectin in PBS (Collaborative Biomedical, 50
gg/ml) for 1 hour.
Surface Plasmon Resonance Spectroscopy. We used the
Biacore
1000 (Pharmacia) for all studies described here. We modified
the manufacturer's chips to accept our substrates, as described
previously [Prime and Whitesides, 1993]. Phosphate-buffered saline
(P3813), fibrinogen (F4883; 94% clottability) and carbonic anhydrase
(C3934) were purchased from Sigma and used as received. Solutions of
proteins were filtered through 0.22 pm filters immediately before use.
Cell Culture. Bovine capillary endothelial (BCE) cells were
isolated from adrenal cortex and cultured as described previously
[Ingber and Folkman, 1989] . Cells were dissociated with trypsin-EDTA,
washed in Dulbecco's Modified Eagle's Medium (DMEM) containing
1% bovine serum albumin (1%BSA/DMEM), and plated onto
46
Chapter 2 Biospecific Surfaces
substrates in chemically defined media (10gg/mL high density
lipoprotein, 5gg/mL transferrin, 5ng/mL basic fibroblast growth factor
in 1%BSA/DMEM) [Ingber, 1990]. Cells were incubated in 10% CO2 at
370 C.
Assessment of efficiency of cell attachment. A fixed number of
cells were plated onto substrates (15,000 cells/cm 2) containing varying
amounts of GRGD peptide. After 4 hours, substrates were gently
washed in PBS and fixed with 4% paraformaldehyde in PBS for 30
minutes. The number of cells attached per field was determined from
photographs taken of samples on a Nikon Axiophot microscope at
200X magnification. At least 6 fields were counted per experiment, and
the experiment was repeated on 3 different occasions.
Measurement of de novo matrix deposition. Cells were
preincubated in 95% cysteine-free, methionine-free medium
(DMEM/cm-) containing 1% dialysed fetal calf serum for 24 h. After
dissociation with trypsin-EDTA, cells were washed in DMEM/cmcontaining 1% BSA and plated for 4 or 24 h onto substrates in the
presence of
3 5S-methionine
and 35S -cysteine (50 pCi/mL, Amersham:
Promix). Substrates were washed 3 times with PBS, and cells were
47
Chapter 2 Biospecific Surfaces
gently extracted with 0.1% ammonium hydroxide, leaving the
deposited matrix on the substrate as previously described [Ingber et al.,
1986]. Deposited matrix was then removed from substrates with
radioimmunoprecipitate assay (RIPA) buffer (1% Triton X-100, 1%
deoxycholate, 0.1% sodium dodecylsulfate, 150mM NaCl, 50 mM TrisHCl, pH 7.2) [Plopper and Ingber, 1993]. Sodium hydroxide and
hydrogen peroxide were added (to 1 M and 2% w/w respectively), and
incubated at 37 C for 10 minutes. Samples were precipitated in ice cold
25% trichloroacetic acid containing 2% casein proteolytic fragments
(Amersham: casamino acids), and filtered through glass fiber filters
(Whatman). After rinsing the precipitate on the filters 3 times in 5%
trichloroacetic acid and once in acetone, samples were dried and
counted in a scintillation counter (Wallac).
Detachment of cells using soluble GRGDSP peptide. Cells were
plated onto substrates for 2, 4 or 24 hours, after which the samples were
transferred to an Omega RTD 0.1 stage heating ring mounted on a
Nikon Diaphot inverted microscope. Cells were then immersed in
defined media containing soluble GRGDSP [Dejana et al., 1987; Sims et
al., 1992] (1 mg/mL, Peninsula Laboratories). Photographs were taken
before, and at 1 to 10 min intervals after, the addition of the soluble
48
-1
Chapter 2 Biospecific Surfaces
peptide. Projected cell areas were determined using these photographs
with image analysis software (Oncor:BDS). Between 25 and 50 cells
were analyzed for each condition and time.
RESULTS
Mixed SAMs of EG 3 0H and EG 60GRGD that contain
1%
EG 6 0GRGD resist the. adsorption of proteins. Using SPR, we measured
the amount of carbonic anhydrase and fibrinogen that adsorbed to
mixed SAMs containing different mole fractions of EG6 0GRGD and
EG 30H (Equation 1). Previous studies using SPR have determined that
a monolayer of carbonic anhydrase and fibrinogen formed on the
surface of the SAMs of hexadecane alkanethiolates gives a AO of 0.150
and 0.45', respectively [Mrksich et al., 1995]. Our results show that
SAMs having values of the mole fraction of GRGD (XGRGD)
0.05 do
not adsorb carbonic anhydrase or fibrinogen, while mixed SAMs
having values of
XGRGD >
0.05 adsorbed less than 5% of a monolayer of
these proteins (Figure 2.17).
Cells adhere and spread to GRGD SAMs in a concentration
dependent manner. Figure 2.18 shows the density of cells attaching to
SAMs presenting different mixtures of EG6 0GRGD and EG3 0H groups.
49
Chapter 2 Biospecific Surfaces
SAMs having
XGRGD
1
X
10-6 completely resisted attachment of
endothelial cells. Cells readily attached to SAMs having XGRGD
but maximal cell spreading was only observed when XGRGD
X
10
1 X 10-3
.
5,
1
Engineered surfaces reduce the deposition of extracellular matrix
by cells. Although SAMs composed of GRGD between XGRGD
and
XGRGD =
=
0-001
0.01 promote complete cell adhesion and spreading, SPR
established that they resist nonspecific protein adsorption (Figure 2.17).
These results suggest that such surfaces allow cells to attach, but resist
the deposition of new extracellular matrix by cells. To examine directly
whether cells can deposit ECM on mixed SAM surfaces, attached cells
were cultured in the presence of radioactive amino acids, followed by
direct measurement of de novo deposition of radioactive proteins onto
the surfaces. Results show that mixed SAMs of GRGD significantly
reduced deposition of new proteins as compared to surfaces presenting
fibronectin nonspecifically adsorbed onto hexadecanethiolate SAMs or
GRGD-containing peptide nonspecifically adsorbed onto glass (Figure
2.19)* . While SAM substrates do not completely resist matrix
*
SAMs of pure EG 3 OH alkanethiolate also resulted in similarly low (<500
cpm) nonspecific
adsorption of radioactive amino acids in the absence of cells. Since cells presented with an
EG3 0H SAM do not attach, they rapidly die. Therefore, this surface was not presented as a
control substrate in the experiment. Attached cells were lysed and radiolabel-counted after
culture to estimate number of cells attached to each substrate. No significant difference in cell
counts were found.
50
Tal W 1===
-
--
*Mer-mm.allimanism
MII,
,1811
.IIIb
iI
.
.
-
Chapter 2 Biospecific Surfaces
deposition, the reduction of protein accumulation on SAM substrates
may increase the window of time during which the effects of specific
ligand-receptor interactions on cellular processes can be studied.
Cells adhere biospecifically to surfaces presenting GRGD ligand.
The previous experiment demonstrates that a newly deposited layer of
ECM forms on surfaces within several hours after attachment of cells,
possibly allowing cells to attach via mechanisms that do not involve
binding to RGD. To test whether cells over time continued to attach to
surfaces via GRGD moieties, or if multiple interactions developed, cells
were allowed to spread on substrates for up to 24 hours, and challenged
with soluble GRGDSP peptide. Cells were first allowed to attach to the
GRGD-presenting substrates for 2 h. In that interval, they spread
equally on the 4 different substrates - mixed SAMs containing either
0.1% or 1% GRGD, fibronectin nonspecifically adsorbed to SAMs of
hexadecanethiolate, and GRGD-containing peptide nonspecifically
adsorbed to glass. Addition of the soluble peptide caused the rapid
release (2-30 min) of cells from all substrates (1%, 0.1% GRGD, FN, and
RGD coated directly on glass). These results support previous findings
that cell attachment via GRGD is reversible, and demonstrates that it is
51
A
Chapter 2 Biospecific Surfaces
only necessary to antagonize interactions with binding sites for GRGD
to cause release of an attached cell from a surface presenting this ligand.
When cells were attached for 4 h before challenging with soluble
GRGDSP, cells partially resisted detachment from FN-coated SAMs and
RGD-coated glass. In contrast, cells continued to rapidly detach from
mixed SAMs upon addition of soluble GRGDSP (Figure 2.20). Even
after culturing cells on substrates for 24 hours, GRGD mixed SAMs
continued to detach readily from cells in response to soluble peptide,
whereas cells could not be detached from FN-coated SAMs and RGDcoated glass substrates.
The significance of these results is to suggest that only the
specific interactions of GRGD with cellular receptors are involved in
initial adhesion (i.e., less than 2 h) of cells to RGD-containing
substrates. Events are more complex, however, with cells allowed to
attach for 24 h to these systems. Within 4 hours cells were able to
deposit a functional non-RGD matrix on FN-coated SAMs and RGDcoated glass substrates such that cell adhesion was partially mediated
through binding of other cell-surface adhesive receptors to this matrix.
By 24 hours these undefined cell-matrix interactions could support the
adhesion and spreading of cells entirely, as evidenced by the inability of
soluble GRGDSP to detach cells from these substrates. In contrast,
52
Chapter 2 Biospecific Surfaces
adhesion of cells to the mixed SAMs presenting EG3 0H and
EG 6 0GRGD do not lose specificity over time; only interactions with
GRGD remain important.
The rapidity with which cells detached from the SAM is
remarkable, and has not previously been reported. Past studies describe
the average time of detachment of these cells to be between 20 and 60
minutes [Sims et al., 1992], whereas most cells were detached from
SAM substrates by 2 minutes. These data suggest that substrates that
are traditionally used to present RGD also contain additional unknown
adhesive factors in the background of the substrate. They also suggest
that substrates such as these mixed SAMs might be useful in
experiments that require rapid release of attached cells without the
damage to cell surfaces caused by treatment with proteases or by other,
non-discriminate methods of releasing cells.
These results define sharply the molecular interaction involved
in adhesion between these cells and mixed SAMs presenting GRGD
and EG 3 OH moieties, and eliminates the ambiguity in interpretation
that has obscured results obtained with surfaces that are less carefully
tailored than these to limit adhesion to only one interaction.
CONCLUSIONS
53
Chapter 2 Biospecific Surfaces
Fundamental studies of mechanisms of cell adhesion have been
limited in the past by the inability to design and generate surfaces with
defined molecular structure. Using SAMs to address these technical
limitations, we have demonstrated a general approach to engineer
surfaces with precise molecular and chemical compositions to study
specific cell-substrate interactions.
Previous studies have shown that the binding of specific cellsurface integrins to the RGD peptide can mediate adhesion of cells to
substrates [Massia and Hubbell, 1991; Ruoslahti and Pierschbacher,
1987]. Interpretation of these studies, however, was limited because it
was not possible to determine the initial density of GRGD moieties
required for efficient cell attachment and spreading, as well as the
changes of the surface over time which result from the active
degradation and redeposition of extracellular matrix onto the substrate
by the cells.
By engineering a surface containing GRGD peptide directly
linked to the protein-resistant ethylene glycol SAM interface, we have
created a substrate that allows biospecific adsorption of cells directly to
the SAM. The use of an ethylene glycol in the SAM reduces the
deposition of new matrix onto the surface. As a result, the specificity of
the cell-substrate interaction was maintained for at least 24 hours. By
54
Chapter 2 Biospecific Surfaces
using substrates that present GRGD in an otherwise inert background,
we demonstrate that integrin-RGD interactions alone are sufficient for
long term attachment and survival of cells.
Acknowledgments. This work was supported by the National Institutes of
Health (GM 30367 to G.M.W. and CA 55833 to D.E.I.), the Office of Naval
Research, the Advanced Research Projects Agency and the National Science
Foundation (DMR-94-00369). C.S.R and M.M. are grateful to the National
Science Foundation and the American Cancer Society, respectively, for
postdoctoral fellowships. C.S.C. was partially supported by the Harvard-M.I.T.
Division of Health Sciences and Technology Medical Engineering and
Medical Physics Program.
55
Chapter 2 Figures
Figure 2.1. The microcontact printing (gCP) process. A
poly(dimethylsiloxane) (PDMS) stamp is fabricated by casting the prepolymer
against a master pattern (a) to give a stamp having a complementary pattern
of relief (b). The stamp is "inked" with an alkanethiol (c) and brought into
conformal contact with a surface of gold (d). A SAM of alkanethiolates is
formed only at those regions where the stamp contacts the surface (e); the
bare regions of gold remaining after the printing process can be modified with
a different SAM by immersing the substrate in a solution of a second
alkanethiol (f). If the stamped alkanethiol promotes protein adsorption (e.g.,
hexadecanethiol) and the immersion thiol resists protein adsorption (e.g.,
oligo(ethylene glycol) undecanethiol), then when the substrate is immersed
in a solution of fibronectin, or other protein, the protein selectively adsorbs to
the stamped regions (g).
56
~-1 gm
Master
PDMS
&PDMS
c
--
K-
p
CH 3 (CH2 1 5 SH
d
PDMS,/
e
0.2 -100g m
111 i
11
11111 1i1111
f
r ria
SAM (2-3 nm)
Au (10-200 nm)
Ti (1-10 nm)
Si (0.5-2 mm)
EG
FIBRONECTIN
57
~2I
Chapter 2 Figures
Figure 2.2. Photoresist patterns on silicon masters generated from different
types of masks. Photoresist spin coated onto silicon was etched away
photolithographically through either a high resolution laser printed mask (A,
B), chromium on glass mask produced by step reduction methods (C, D), or
chromium on glass mask produced by electron-beam etching (E, F).
Micrographs show low (A, C, E) and high (B, D, F) magnification of features.
(A, B) 30 um squares; (C) 3 to 50 um squares; (D) 40 um square; (E, F) 5 um
circles.
58
B
A
II
D
F
100gm
59
20gm
Chapter 2 Figures
Figure 2.3. Patterning of cells onto a SAM substrate. A SAM substrate is
produced from a stamp having the pattern diagrammed in (A), where shaded
regions indicate where the stamp will print hydrophobic, adhesive SAM onto
a gold coated substrate. (B) When this substrate is immersed in a solution of
fibronectin (50 ug/ml in PBS) for 1 hr, the fibronectin is selectively adsorbed
onto these regions, as shown by immunofluorescent staining of the substrate.
(C) Cell plated onto such a substrate only adhere to the fibronectin-coated
regions, as shown by phase-contrast microscopy.
60
E
20
5
75 330
.5
5
s
47 .
S
-
F 43.
1
30
2
0
E
4E
55
3
7
5
20
E
3
10
0m
7
2C
0
B
C
m
-
C1
61
73r.74
51
7p
Chapter 2 Figures
Figure 2.4. Phase-contrast micrographs of bovine capillary endothelial cells
cultured on fibronectin-coated lines of 90 and 10 um widths (A and B,
respectively).
62
90
AM
106M
63
Chapter 2 Figures
Figure 2.5. (A) Phase-contrast micrograph of bovine capillary endothelial cells
cultured on a substrate containing fibronectin-coated circles 20 um in
diameter (top left region), lines 20 um in width (top right region), and a large
unpatterned region (bottom). (B) High magnification micrograph of cells on
these circular islands.
64
B
gm
40 gm
65
Chapter 2 Figures
Figure 2.6. Differential interference contrast micrograph of bovine capillary
endothelial cells cultured on fibronectin-coated squares of various sizes (5, 10,
20, and 40 um side length).
66
10pgm
40 gm
204m
5
E
67
m
Chapter 2 Figures
Figure 2.7. Differential interference contrast micrograph of a single bovine
capillary endothelial cell cultured on a fibronectin-coated square (50 um side
length). To the left is a large region of fibronectin where cell spreading is
unrestricted.
68
NONADHESIVE
50 gm
69
Chapter 2 Figures
Figure 2.8. Cells spread across multiple, circular islands coated with
fibronectin. Fluorescence micrographs of substrates simultaneously stained
for fibronectin (top) and vinculin (bottom). The cell borders have been
outlined (top) to illustrate their location.
70
Fibronectin
Vinculin
71
Chapter 2 Figures
Figure 2.9. Effect of serum on patterning of BCE cells. Cells were plated in the
absence (A) or presence (B) of serum for 24 hours.
72
Bovine Capillary Endothelial Cells
IA
A. Serum Free
B. 10% Calf Serum
73
Chapter 2 Figures
Figure 2.10. Effect of serum on patterning of 3T3-L1 preadipocytes. (A) Cells
were plated in the absence of serum for 24 hours. (B) After 24 hours, cells
were exposed to serum for 1 hr.
74
Mouse 3T3-L1 Pre-Adipocytes
A. Serum Free
B. 10% Calf Serum
75
-nub"!
Chapter 2 Figures
Figure 2.11. Procedure for preparing contoured substrates. The method is
explained in the text.
76
SiO 2
Si
Glass Slide
[~$><$P~..tK
g
5a
Glass Slide
b
cs
rs
ss
h
ESSE
Au (12 rm).
Ti (1.5i
c
Glass Slide
d
PDMS/%e
PDMS/ *,.%.-,
Glass Slide
*e
sPDMS.
Glass Slide
.polyurethane
Glass Slide
Glass Slide
77
-
ME -
-
--
Chapter 2 Figures
Figure 2.12. Scanning electron micrograph of a contoured film of
polyurethane supported on a glass slide (as in Figure 2.6, part g). The
contoured substrate was frozen in nitrogen, fractured along a plane
perpendicular to the array of lines, and sputtered with gold (20 nm) prior to
the electron microscopy.
78
50 tm
79
Chapter 2 Figures
Figure 2.13. SEM images of endothelial cells cultured on contoured substrates
having ridges and grooves 25 gm in width. The substrates were tailored with
SAMs presenting either methyl or tri(ethylene glycol) groups; fibronectin
(FN) adsorbed only on the methyl-terminated regions; cells attached only to
these regions presenting fibronectin. (A) The entire substrate presents
fibronectin (FN). (B) Only the ridges present fibronectin. (C) Only the
grooves present fibronectin. (D) None of the areas present fibronectin. After
2 days in culture, cells were fixed in Karnovsky's fixative, critical point dried,
and sputtered with 20 nm of gold. The scale bar applies to all images.
80
-CH 3 /FN
-CH 3 /FN
-CH3/FN
-EG30H
-EG 3 OH
-CH 3 /FN
-EG 3 OH
J -EG 30H
100 gm
81
Chapter 2 Figures
Figure 2.14. Optical micrographs of endothelial cells cultured on
contoured substrates having ridges and grooves 50 gm in width. (A)
The entire substrate was tailored with a SAM presenting methyl groups
and fibronectin (FN). (B) Only the plateau regions were tailored with a
SAM presenting methyl groups and fibronectin. Both photographs are
at the same magnification and include an unpatterned planar region to
the left. Cells were fixed in 3.7% paraformaldehyde and stained with
1:1 Giemsa:Comassie. (C) Optical micrograph of endothelial cells
attached on a contoured surface after fluorescence staining of the Factin microfilaments with rhodaminated-phalloidin.
The upper
region shows cells on a planar non-patterned region and the lower
region shows cells confined to a plateau 50 gm in width. Cells were
fixed in 3.7 % paraformaldehyde prior to staining with the phalloidin.
82
CH3 /FN
CH3 /FN
Z: EC3 OH
1OO sm
-CH 3 /FN
-EG0H
-CH/FN
30 Rm.
83
I
Chapter 2 Figures
Figure 2.15. Diagram of self-assembled monolayer of alkanethiolates
on gold presenting EG6 0GRGD and EG 3 0H groups.
84
NHGRGDCOH
oq (NHGRGDc O2H
L\O-\,o
0)
T
GRGD
Of0
OH
0
HO
HO
(EG)nO i
(%H?
0
00
0
0H
0(
0
0CH0)
0
0 0
S
S
///////Au
s
s
/ //
s
/I
85
s
/
/
0I
2
11
Thiol
Substrate
Chapter 2 Figures
Figure 2.16. Synthesis of GRGD hexaethyleneglycol alkanethiol (1). (i)
ZNHCH 2 CONHS, DIPEA; (ii) ZNHCH 2 CONHS, DIPEA, DMF; (iii)
DPPA, DIPEA, DMF; (iv) NaH, DMF than BrCH 2COOt-Bu; (v) TFA,
CH2C 2; (vi) 10% Pd/C, EtOH on 4 than DPPA, DIPEA, DMF; (vii)
CH 3COSH, AIBN, THF with UV irradiation; (viii) TFA, PhSMe,
HS(CH 2 ) 2 SH, CH2 C
2
86
PMCNH
=NH
iv
62%
RlN
OH
+
RN N
H
H
2
94
0
3
R = COCH 2NHZ
iii
82%
R -OCH2NH2
71%
v 86%
)NH
HN
00
2
o
+
O
ZHN,JK
N0
HH
4
0
vi
48 %
0
PMcNH
0-40
0
0
vii
93
HN )=NH
0
~PMCNHN
HN
00
viii
3 %
0
>=NH
2
0
HS0
0
87
0
A-
Chapter 2 Figures
Figure 2.17. Percentage of a monolayer of protein irreversibly adsorbed
to mixed SAMs presenting GRGD groups and tri(ethylene glycol)
groups. SPR response curves were obtained for the adsorption of
carbonic anhydrase and fibrinogen to SAMs containing different mole
fractions of GRGD groups (top, middle panels, respectively). From
these curves, steady state adsorption of proteins were determined as a
function of XGRGD. The steady state change in resonance angles (AO)
were normalized to previously measured values for a full monolayer
of these proteins (AO of 0.15' and 0.45', respectively) [Mrksich et al.,
1995]; values of XGRGD are indicated on the plot.
88
Buffer SDS
Buffer
Protein
Buffer
II
0.16
Carbonic Anhydrase
0.140)
0
~0
I
0.12-
xRGD
0.10
~----
-0
XRGD
-
.005
0.080.06
0.04
- -----------------
0.020
---
0.12-
300
400
500
~---
600
700
Fibrinogen
0.10
XRGD
=
0.08
- -------- XRGD
=
1.0
0.005
0.06
0.04
----------------
0.02
---------------------
0
200
)
100
300
400
500
600
700
800
Time (s)
10
-
0)
0
"0
200
100
0.14-
o
*
Carbonic Anhydrase
Fibrinogen
8-
6-
4-
I
00I
*
U
2U
o
'
0
0
0
0.6
0.8
I
0.2
0'
0.05
0.4
XGRGD
89
1.0
Chapter 2 Figures
Figure 2.18. Adhesion of endothelial cells to mixed SAMs presenting
GRGD groups and tri(ethylene glycol) groups; nominal values of
XRGD
are indicated on the plot. The number of cells attached per field are
indicated on the vertical axis. The photographs above the bar graph
illustrate the amount of cell attachment and cell spreading observed for
a given SAM: (A) cells on fibronectin-coated on (CH2) 15 CH3 SAM; (B)
cells are attached and spread; (C) cells are attached and spread; (D) cells
are attached but do not spread; (E) cells are unable to attach.
90
DPTW'w
-r-7
...... ......................... ...........
.....
.......
.......
200
...........
...........
-
......
...........
150
L
.............
...
.......
...........
-
loo
............. .........................
50
0
FN
1
10-1
10-2
10-3
10-4
10-5
X(EG)60GRGD
91
10-6
10-7
10-8
0
Chapter 2 Figures
Figure 2.19. Direct measurement of de novo deposition of radioactive
proteins onto mixed SAMs presenting GRGD groups and tri(ethylene
glycol) groups. Cells were cultured on mixed SAMs presenting GRGD
groups and tri(ethylene glycol) groups, fibronectin-coated SAMs of
hexadecanethiolate (C16-SAM) or RGD-coated glass for 4 or 24 hours in
35
the presence of radioactive ( S) amino acids. (A) Measurement of
radioactivity (cpm) of substrate after removal of cells at 4 or 24 hours,
indicating amount of de novo deposition of proteins. (B)
Measurement of radioactivity (cpm) of control substrates immersed in
radioactive medium in the absence of cells for 4 or 24 hours, indicating
amount of passive adsorption of amino acids.
92
-1
20000
15000-
0
10000
/
LO
/
E
cCL
/
10
5000-
/
/
-
-
-
//
z
U,
0
N
0.1% GRGD-SAM
1% GRGD-SAM
0
Z
Fibronectin on C16-SAM
c15000-
RGD on glass
0
C
0
10000
-
C.)
C)
c,
5000'4-
0
z
00
0
Time of Incubation
93
--I
Chapter 2 Figures
Figure 2.20. Retraction of cells caused by soluble GRGDSP peptide.
Bovine capillary endothelial cells were allowed to attach to these
substrates: mixed SAMs presenting GRGD and tri(ethylene glycol)
groups, fibronectin-coated SAMs of hexadecanethiolate, and RGDcoated glass. After allowing attachment to surfaces for 2, 4 or 24 h,
soluble GRGDSP peptide to a concentration of 0.5 mg/ml. (A, B) Phase
contrast micrographs of cells (attached 4 or 24 hours, respectively)
immediately before and 10 minutes after addition of peptide. (C) Plots
of projected cell area over time after the addition of peptide.
94
A
B
Plated for
24 Hours
Plated for GRGDSP for
10 Minutes
4 Hours
C
GRGDSP for
10 Minutes
1001
for2 hours
8060-
.
....
Fibronectin/ C16
400
4)
N
0
....................................
RGD-coated Glass
0100
Cells plated
for 4 hours
80
pq
i
60
-U---
FN /C16
-----
0.1%GRGD
1%GRGD
40
RGD on Glass
20
0
a)
0.1% GRGD
0"
Cells plated
1004
for 24 hours
8060-
........... .....
.
.
-
40
20
1% GRGD
-
(IN
.....
.....
20-
............
-
o~
o
-
I
0
Time after adding soluble GRGDS P (minutes)
Chapter 3 ECM , growth and apoptosis
CHAPTER III. GEOMETRIC CONTROL OF CELL LIFE AND
DEATH*
Preface
This chapter describes the study of the effect of cell shape on cell
proliferation and apoptosis. As a submission to SCIENCE, this study omitted
many of the background details and supporting evidence collected. For
completeness, this chapter includes more complete scientific methods and
additional results in Appendix A, leaving the text virtually unchanged.
Capillary endothelial cells were switched from growth to apoptosis using
micropatterned substrates containing extracellular matrix-coated adhesive
islands of decreasing size to progressively restrict cell extension. Cell
spreading also was varied while maintaining total cell-matrix contact area
constant by changing the spacing between multiple focal adhesion-sized
islands. Cell shape was found to govern whether individual cells will grow
or die, regardless of the type of matrix protein or anti-integrin antibody used
to mediate adhesion. Local geometric control of cell growth and viability may
therefore represent a fundamental mechanism for developmental regulation
within the tissue microenvironment.
* Contributing authors: Christopher S. Chen, Milan Mrksich, Sui Huang, George M. Whitesides
and Donald E. Ingber. CC performed all experiments in this chapter. MM aided in the
production of substrates, and SH aided in studies with human cells.
96
Chapter 3 ECM , growth and apoptosis
The local differentials in cell growth and viability that drive
morphogenesis in complex tissues, such as branching capillary networks
[Clark and Clark, 1938; Ingber et al., 1986], are controlled through modulation
of cell binding to extracellular matrix (ECM) [Ingber and Folkman, 1989;
Roskelley et al., 1995; Ingber and Folkman, 1989a; Wicha et al., 1980; Drake et
al., 1995; Sympson et al., 1994]. Local disruption of ECM by pharmacological or
genetic means results in programmed cell death (apoptosis) selectively within
adjacent cells [Ingber et al., 1986; Sympson et al., 1994; Boudreau et al., 1995].
Soluble integrin caVP3 antagonists also induce apoptosis in cultured
endothelial cells and promote capillary involution in vivo [Brooks et al.,
1994; Brooks et al., 1994; Brooks et al., 1995; Stromblad et al., 1996].
Furthermore, death can be prevented by allowing suspended cells to attach to
immobilized anti-integrin antibodies or by inhibiting tyrosine phosphatases
[Boudreau et al., 1995; Meredith et al., 1993; Zhang et al., 1995]. For these
reasons, adhesion-dependent control of apoptosis has been assumed to be
mediated by changes in integrin signaling. Analysis of capillary regression in
vivo has revealed, however, that dying capillary cells remain in contact with
ECM fragments and instead suggest that the cell foreshortening caused by
ECM dissolution may be the signal that initiates the death program [Ingber et
al., 1986]. This possibility is supported by the finding that endothelial cells
spread and grow on large (> 100 um diameter) microcarrier beads [Ingber and
Folkman, 1989] whereas they rapidly die when bound to small (4.5 um) ECMcoated beads [Re et al., 1994] that cluster integrins and activate signaling, but
do not support cell extension [McNamee et al., 1993; Schwartz et al., 1991; Dike
and Ingber, 1996].
Understanding how this apoptotic switch is controlled in capillary cells
has enormous clinical implications since angiogenesis is a prerequisite for
97
Chapter 3 ECM , growth and apoptosis
tumor growth [Folkman 1971; Hanahan and Folkman, 1996; Folkman et al.,
1989, Ingber et al., 1990]. Thus, we set out to determine whether cell shape or
integrin binding per se governs life versus death in these cells. We first
measured apoptosis rates in suspended cells attached to a range of different
sized beads coated with fibronectin (FN).
Nearly all cells survived when
spread on FN-coated planar dishes in medium containing saturating
amounts of growth factors whereas approximately 60% of non-adherent cells
entered the death program within 24 hr (Fig. 3.1A). In contrast, less than 10%
of cells adherent to large (> 25 um) FN-coated beads underwent programmed
cell death (Fig. 3.1B). Unlike suspended cells which remained small and
spherical, these cells and their nuclei appeared to flatten as they extended
around the beads (Fig. 3.1A). Importantly, as the bead diameter was decreased
to 10 um, cells became more round, and the apoptotic index increased to
match the levels in non-adherent cells (Fig. 3.1B).
The size of these spherical beads not only affects the degree of cell and
nuclear spreading, but also ECM curvature and bead internalization. The 10
um beads appeared fully engulfed by cells within 4hr whereas 25 um beads
were never fully internalized. To eliminate these complicating factors, we
fabricated planar adhesive islands of defined size and shape, separated by nonadhesive regions, using a microscale patterning technique [Singhvi et al.,
1994; Prime and Whitesides, 1991; Kumar et al., 1994; Mrksich and
Whitesides, 1995; Mrksich et al., 1997]. When plated on circular FN-coated
islands 10 or 20 um in diameter, cells spread until they took on the size and
shape of the underlying adhesive island (Fig. 3.1C). Significantly more cells
entered apoptosis when held in a round form on 20 um circles than when
spread on identically-fabricated unpatterned substrates (Fig. 3.1D).
Furthermore, the subtle decrease in cell and nuclear spreading observed in
98
ni-a
--
-
-.
-vaans.hewima
.
-man
s.
midlilMI AINI
i iE.. ni
,.
'-
"''-
Chapter 3 ECM , growth and apoptosis
cells on 10 versus 20 um islands (Fig. 3.1C) was also accompanied by a
statistically significant increase in apoptosis (Fig. 3.1D).
Capillary cell spreading on ECM also has been shown to modulate cell
cycle progression [Ingber and Folkman, 1989; McNamee et al., 1993; Schwartz
et al., 1991; Dike and Ingber, 1996; Folkman and Moscona, 1978; Ingber, 1990].
To determine the precise spreading requirements for survival versus growth,
GO-synchronized cells were cultured on different sized FN-coated adhesive
islands. The different sized islands were contained within a single substrate
(Fig. 3.2A) to rule out the possibility that changes in cell behavior could be
due to release of paracrine growth-modulators. When cells were plated on
square shaped islands coated with FN, square shaped cells were produced that
closely matched the size and shape of the adhesive island (Fig. 3.2A).
Apoptosis progressively declined when the island size was increased from 75
to 3000 um2 whereas DNA synthesis was concomitantly switched on as cell
and nuclear spreading were promoted (Fig. 3.2A,B).
These results demonstrate that increasing cell spreading on a
homogeneous, high-density coating of FN leads to cell survival and growth.
However, the total area of cell-ECM contact also increases under these
conditions and thus, integrin binding, focal adhesion formation and
accessibility to matrix-bound growth factors [Folkman et al., 1988; SpivakKroizman et al., 1994; Falcone et al., 1993] may all vary in parallel. To explore
this mechanism more fully, apoptosis and growth were evaluated in single
cells spread across multiple, closely-spaced adhesive islands either 3 or 5 urn
in diameter, to approximate the size of individual focal adhesions (Fig.
3.3A,B). Cell bodies spread across the intervening non-adhesive areas of the
substrate, stretching processes from one small adhesive island to another.
Immunofluorescence staining confirmed that adherent cells only attached
99
.___ -4
0_0
Chapter 3 ECM , growth and apoptosis
and formed vinculin-containing focal adhesions on the engineered islands
(Fig. 3.3B,C). By changing the spacing between adhesive islands, cell
spreading could be increased more than 10 fold without significantly altering
the total cell-ECM contact area (Fig. 3.3D). On these substrates, DNA synthesis
scaled directly with projected cell area and not with cell-ECM contact area (Fig.
3.3D). Apoptosis was similarly switched off by cell spreading, even though
the cell-ECM contact area remained constant under these conditions (Fig.
3.3D). Thus, cell shape per se appears to be the critical determinant that
switches cells between life and death and between proliferation and
quiescence.
In vivo studies demonstrate that programmed cell death and capillary
regression can be induced by inhibiting integrin oVP3 binding whereas
apoptosis can be prevented in vitro by cell attachment to immobilized antiintegrin
P1
antibodies [Brooks et al., 1994; Brooks et al., 1994; Brooks et al.,
1995; Stromblad et al., 1996; Meredith et alk., 1993; Zhang et al., 1995].
Because cell binding to FN is mediated by both
P1 and P3 integrins,
.
---
we chose
to explore their role in shape-dependent control of apoptosis. Unpatterned
substrates and 20 um circular islands were coated with antibodies specific for
1 or ixVf3 integrins, FN, or physiological ECM ligands that preferentially
utilize integrin f1 (type I collagen) or aVP3 (vitronectin). Apoptosis was
greatly inhibited relative to the 60% level observed in suspended cells when
cells spread on unpatterned substrates, regardless of the integrin ligand
utilized (Fig. 3.4). However, survival was consistently greater in cells
adherent to intact ECM proteins. Interestingly, when spreading was restricted
by use of 20 um circles, cells adherent to integrin
P1 ligands
(FN, type I
collagen, anti-1 antibody, or anti-1 combined with anti-aVP3) exhibited
much greater increases in apoptosis compared with those on either intact
100
Chapter 3 ECM , growth and apoptosis
vitronectin or anti-xVP3 antibody alone (Fig. 3.4). This was due to a change
in sensitivity, rather than a lack of response, since similar high levels of
apoptosis were induced when cell spreading was further restricted by plating
on 10 um circular islands. Thus, while geometric switching between growth
and apoptosis is a general phenomenon, different adhesion receptors appear
to be able to convey distinct death signals and thereby tune the cellular
response to shape distortion.
The mechanism by which cells transduce changes in cell geometry into
different biochemical responses remains unclear. The specialized cytoskeletal
structure, or focal adhesion complex, that forms intracellularly at the site of
integrin binding is a molecular bridge that mechanically couples integrins,
and hence ECM, to the actin cytoskeleton [Burridge et al., 1988; Craig and
Johnson, 1996; Wang et al., 1993; Wang and Ingber 1994; Wang and Ingber,
1995; Maniotis et al., 1997]. Because focal adhesions also orient much of the
signal transduction machinery of the cell [Clarke and Brugge, 1995; Schwartz
et al., 1995;Ingber, 1993; Plopper et al., 1995; Miyamoto et al., 1995], they may
integrate mechanical signals associated with changes in cell shape with
chemical signals elicited directly by integrin binding and thereby, modulate
downstream signaling [Ingber, 1997]. In fact, constitutive activation of FAK
kinase in the focal adhesion complex can lead to shape- and adhesionindependent cell survival and growth [Owens et al., 1995; Frisch et al., 1996].
Alternatively, growth and viability may be altered directly via mechanical
stress-dependent changes in the organization or stiffness of the cytoskeleton
and nucleus [Wang et al., 1993; Wang and Ingber 1994; Wang and Ingber, 1995;
Maniotis et al., 1997; Ingber, 1997; Stamenovic et al., 1996; Ingber, 1993]. For
example, the increased flexibility of the cytoskeleton observed in rounded
101
Chapter 3 ECM , growth and apoptosis
cells [Wang et al., 1993; Wang and Ingber 1994; Wang and Ingber, 1995] may
permit intracellular structural rearrangements that are lethal, including the
characteristic structural degeneration of the cell and nucleus that are
hallmarks of apoptosis. The finding that cell survival is more tightly coupled
to cell shape in cells adherent to ligands for integrin
also consistent with the observation that
P1 provides
P1 compared
to XVp3 is
stronger ECM anchoring
to resist cytoskeletal tension [Wang et al., 1993; Wang and Ingber 1994; Wang
and Ingber, 1995]. From this perspective, adhesive substrates may prevent cell
death and promote growth by resisting contractile forces transmitted across
integrins and thereby, mechanically stabilizing the nucleocytoskeletal lattice.
During morphogenesis, growing, quiescent and dying cells often coexist
within the same microenvironment [Clark and Clark, 1938; Ingber et al., 1986].
In fact, it is the establishment of local differentials in cell growth and viability
that drives pattern formation. Our results suggest that living cells can filter
the same set of chemical inputs (activation of integrin and growth factor
receptor signaling) to produce different functional outputs (growth versus
apoptosis) as a result of local mechanical deformation of the cell or nucleus.
By sensing their degree of extension or compression, cells therefore may be
able to monitor local changes in cell crowding or ECM compliance (e.g., due to
enhanced ECM remodeling or local application of cell tension) and thereby,
couple changes in ECM extension to cell mass expansion within the local
tissue microenvironment.
Tissue involution may be promoted in other
microenvironments by inducing rapid breakdown of ECM and associated cell
retraction. During malignant transformation, progressive loss of shapedependent regulation also may lead to cell survival in the absence of ECM
extension, unrestricted mass expansion, and hence, neoplastic
disorganization of tissue architecture [Ingber et al., 1981; Ingber and Jamieson,
102
Chapter 3 ECM , growth and apoptosis
1982; Ingber and Jamieson, 1985; Ingber et al., 1985; MacPherson and
Montagnier, 1964; Stoker et al., 1968; Wittelsberger et al., 1981; Tucker et al.,
.
1981; Folkman and Greenspan, 1975]*
This work was supported by grants from NIH (HL57669, CA55833, & GM30367), DARPA, and
ONR; postdoctoral fellowships from American Cancer Society (M.M) and the Swiss National
Science Foundation (S.H.); and partial salary support (C.C.) from the Harvard-MIT Health
Sciences Technology Program.
*
103
Chapter 3 Appendix A
APPENDIX A. SUPPORTING METHODS AND RESULTS
METHODS
Cell Culture
Prior to experiments, cells were cultured in standard growth media
(bovine capillary endothelial (BCE) cells in 10% C02 on gelatin-coated plastic
in DMEM; 10% calf serum; 2mM glutamine; 100u per ml streptomycin; 100u
per ml penicillin; ing per ml bFGF, and human endothelial cells (HMVEC) in
EBM; 10% fetal calf serum; 1 ug per ml hydrocortisone; 10 ng per ml EGF; 10
ug per ml bovine brain extract; 50ug per ml gentamycin; 50 ug per ml
amphotericin-B). In experimental medium, serum was reduced (to 2% for
HMVECs) or removed (for BCEs), and saturating amount of basic fibroblast
growth factor (5 ng/ml) , human high density lipoprotein (10 gg/ml), and
transferrin (10 gg/ml) were added. Although experiments were conducted
with reduced or no serum, several preliminary studies had indicated that the
presence of serum did not significantly alter the shape-dependent induction
of apoptosis (Figure 3.5).
For attaching cells to beads, endothelial cells were suspended with
trypsin and washed with experimental media. Beads coated overnight with
50pg/ml of fibronectin in 0.1M carbonate buffer, pH=9.4, were incubated with
the cells at a 1:1 ratio (106 beads/ml) for 1 hour to allow cells to attach to the
beads. The mixture was then diluted 1:10 into 2% methylcellulose in
experimental media to maintain them in suspension. Cells were recovered
for analysis by fixing with 4% formaldehyde for 30 minutes, diluting the
suspension with PBS, and spinning the samples in a centrifuge. Cells were
resuspended in 1 ml of PBS and dried onto gelatin-coated slides at 400 C.
Samples were stained directly on the slides.
104
Chapter 3 Appendix A
Microfabrication of Patterned Substrates
We used microcontact printing (gCP) techniques to fabricate substrates
patterned with regions that adsorb ECM and regions that resist such
adsorption, as previously described (Singhvi). Briefly, patterned substrates
were prepared as follows (Figure 2.1). Patterned molds for making stamps
were photolithographically produced using standard techniques. Briefly, in a
clean room (100), silicon <111> wafers were cleaned, spin coated with 2gm
layer of poly methylmethacrylate photoresist, and baked. The wafers were
exposed to high energy UV light through a photolithographic mask
containing the desired pattern. The wafers were developed and washed,
leaving 2gm thick photoresist where the UV was masked, and naked silicon
elsewhere. We prepared a poly(dimethylsiloxane) (PDMS) stamp from this
silicon master by polymerizing prepolymer on top of the master. Substrates
for cells were then prepared by evaporation of thin films of titanium (1.5 nm)
and gold (12 nm) on glass cover slips (0.20 mm, No.2, Corning). A cotton
swab was wetted with a solution of hexadecanethiol (HS(CH2)15CH3, 2mM in
ethanol) and dragged once across the face of a PDMS stamp molded from a
silicon master as described above; the stamp was dried with a stream of
nitrogen for 10 s, and placed gently on a metallized glass slide with sufficient
pressure to promote conformal contact between the stamp and the substrate.
After 5 s, the stamp was removed from the substrate, taking care not to
"double-stamp" the substrate. The slide was immersed immediately in a
solution of the tri(ethylene glycol)-terminated alkanethiol in ethanol
(HS(CH2)11(OCH2CH2)30H, 2mM) for 30 minutes; the slide was removed,
rinsed with ethanol, and dried with a stream of nitrogen. Hexadecanethiol
was purchased from Aldrich and purified by silica gel column
105
Chapter 3 Appendix A
chromatography; the tri(ethylene glycol)-terminated alkanethiol was
synthesized as described previously. The adsorption of protein on
hydrophobic SAMs of hexadecanethiolate is usually rapid and irreversible.
SAMs presenting oligomers of the ethylene glycol group are very effective at
resisting the adsorption of protein. Therefore, when these substrate were
immersed in 50 ug/ml of FN in PBS, FN rapidly adsorbed to the stamped
regions. After rinsing with PBS, the substrates were treated using standard
cell culture techniques.
Detection of S-phase entry
After cells were cultured on different patterns for 24 hours, the
percentage of cells that entered S phase was determined by staining for 3
different antigens:
(1) BrdU incorporation (2) Ki-67, a protein expressed in S
phase, and (3) Proliferating Cell Nuclear Antigen, PCNA, also a cell cycle
protein expressed in S phase. Cells to be stained for BrdU were fixed in 90%
ethanol/5% H20/5% acetic acid for 30 minutes, washed in PBS, and incubated
with primary mouse anti-BrdU antibody (Amersham) containing nuclease.
Cells stained for Ki-67 were fixed in 4% paraformaldehyde for 20 minutes,
washed in PBS, and incubated with primary mouse anti-Ki-67 antibody in
immunofluorescence buffer (IFB) containing 0.1% Triton X100, 0.1% BSA in
PBS. Cells stained for PCNA were fixed in methanol on ice for 10 minutes,
washed in PBS, and incubated with primary mouse anti-PCNA antibody in
immunofluorescence buffer (IFB) containing 0.1% Triton X100, 0.1% BSA in
PBS. In all cases, mouse antibody was detected using goat-anti-mouse
antibody conjugated to fluorescein (Amersham) for 1 hour.
106
Chapter 3 Appendix A
Immunofluorescence Staining of Cytoskeletal and Matrix Proteins
Cytoskeletal proteins and fibronectin were stained as follows. Samples
were first permeabilized in a cytoskeletal stabilizing buffer (300mM sucrose,
100mM NaCl, 3mM MgCl2, 0.5%Triton X100, 10mM pipes, pH 6.8), then fixed
in 4% formaldehyde for 30 minutes and washed in immunofluorescence
buffer (IFB), containing 0.1% Triton X100, 0.1% BSA in PBS. The sample was
incubated with primary antibody in IFB for 1 hour, washed, and incubated in
fluorescent secondary antibody in IFB for 1 hour, and washed. Actin was
stained with TRITC conjugated phalloidin in IFB for 1 hour, and washed.
Determination of cell , ECM-cell contact, and nuclear areas
Image processing software was used to calculate projected cell and
nuclear areas from images grabbed from the microscope through a CCD
camera.
Projected cell area, perimeter, and diameter were determined from
interactive tracing of cell edges of phase images. Nuclei were visualized
fluorescently by 4,6-diamidino-2-phenylindole (DAPI) staining. ECM area was
calculated based on intersection of images of direct fluorescence staining of
ECM islands with projected cell areas.
Fluorescence quantitation of FAC quantity and signaling
FAC quantity and signaling were assessed by measuring vinculin and
phosphorylated tyrosine accumulation, respectively, with quantitative
immunofluorescent detection with confocal microscopy. Fluorescence
quantitation was calculated from the volume*intensity maps imaged by laser
confocal microscopy of samples labelled by immunofluorescent antibodies to
vinculin.
107
_
~
-~
Chapter 3 Appendix A
Statistical analysis of correlating factors with cell growth
For each series of experiments, a correlation table was generated
between growth and the factors considered. After identifying factors that
showed significant correlation coefficients, stepwise regression was then
performed to determine if any factor provided additional predictive value as
a secondary factor. Scatter plots were then generated, and best-to-fit mean
regression curves were estimated for significant correlations.
RESULTS
Regulation of apoptosis by size of FN island
Nearly all the results obtained in this study were duplicated in two
different cell lines: capillary endothelial cells isolated from bovine adrenal
cortex (BCE) and microvascular endothelial cells isolated from human
pulmonary tissue (HMVEC). Both cell lines attached to FN-coated beads or
microfabricated islands demonstrated a shape-dependent induction of
apoptosis (Figure 3.6). Although the apoptosis rates in bovine cells were
consistently lower than those of human cells, the trends remained apparent.
At longer times, the apoptotic index of bovine cells increased to match those
of human cells. (Figure 3.7).
Regulation of growth by size of FN island
Closer examination of cells attached to square islands reveals that they
not only spread to the size and shape of the engineered islands (Figure 2.3, 2.5,
2.6, 3.2A), but also continue to form active ruffling edges at the FNnonadhesive boundary, indicating their inability to spread beyond this
boundary (Figure 3.8). A switch from apoptosis to growth was observed in
bovine ECs as cells were spread on progressively larger islands (Figure 3.2B).
108
Chapter 3 Appendix A
In this study, DNA synthesis was measured by incorporation of a thymidine
analog (BrdU). Although positive staining for BrdU incorporation
sufficiently indicates S-phase entry, negative staining could be a result of
changes in nucleotide transport that prevent BrdU from entering the nucleus;
thus, the increase in BrdU incorporation with island size could be interpreted
to arise from a shape-sensitive transport mechanism that masks cell cycle
progression. Therefore, EC entry into S-phase on different sized islands was
additionally measured by presence of proteins expressed during S-phase (Ki67 or Proliferating Cell Nuclear Antigen, or PCNA). Results clearly indicated
that cell cycle progression is regulated by the size of ECM island (Figure 3.9).
Human ECs also exhibited a similar shape-dependent switch between growth
and survival (Fig. 3.10). The human experiment shows a 4 hour pulse
incubation of BrdU between 20 and 24 hours after plating, rather than a
cumulative 24 incubation with BrdU. A full time course of BrdU
incorporation in 4 hour pulses reveals that the shape-dependent block of Sphase entry changes the percentage of cells entering S-phase, and not the rate
of progression through the cell cycle (Figure 3.11).
Response of cells to focal adhesion-sized micropatterns
Attachment of cells to FN-coated lines separated by different spacings
demonstrated that cells could not cross 20 um spaces, but could cross 10 um
spaces though spreading is significantly reduced. Across 5 um spaces, cells
could spread almost as much as on unpatterned substrates (Figure 3.12). At
no time were cell extensions observed to terminate in a nonadhesive region.
Cells spread across small, focal adhesion-sized circles (3 or 5 um
diameter) of FN separated by 5 or 10 um spaces were stained for focal
adhesion and CSK proteins to determine how cells were attaching to these
109
Chapter 3 Appendix A
substrates. Simultaneous staining for FN and vinculin indicated that focal
adhesions were forming only above the FN-coated islands and not in the
intervening, nonadhesive spaces (Figure 3.4C). Staining for other structural
proteins of the FAC (talin and paxillin), signaling proteins (FAK,
phosphorylated tyrosines), and integrin receptors (ix5p1 and (xVP3) supported
the finding that focal adhesions were only forming on the grid of adhesive
islands (Figure 3.13). The termination of stress fibers at these focal adhesions
demonstrates that these FACs are functional anchors for the microfilament
lattice (Figure 3.14). Similar results were obtained on other ECM proteins,
such as vitronectin and collagen I (Figure 3.15).
Shape regulation of cell function
The ability to separate cell spreading from total ECM-cell contact using
these focal adhesion-sized substrates indicated that cell growth and apoptosis
were regulated by cell shape (Figure 3.3D). Allowing cells to spread across
multiple lines further supported that growth depends on shape per se, not the
amount of ECM-cell contact (Figure 3.16).
To examine more closely the nature of shape-dependent growth, by
using several additional types of patterns, DNA synthesis was measured in
cells adherent to all the different types of substrates (including squares, with 1
cell per square, focal adhesion-sized circles, lines, and unpatterned substrates),
and plotted as a function of several measured shape parameters, including
projected cell area, cell-ECM contact area, cell length (long axis), and cell
perimeter (Figure 3.17). These results indicate that even over this wide range
of environments, projected cell area consistently predicted growth regardless
of the series of patterns used, such that the data from all substrates appeared
to lie on one curve. Statistical analysis showed that the correlation was
110
Chapter 3 Appendix A
highly significant (r2 = 0.97). In contrast, cell-ECM contact area showed no
consistent predictive value for growth. Other measures for the shape of a cell
also did not show any relationship with cell growth.
Mechanism of regulation of cell function by shape
Two different hypotheses can be proposed that may provide a
mechanistic explanation for how cell spreading regulates growth, both of
which can be indirectly tested in this system. The first proposal, as suggested
in the discussion of this chapter, is that focal adhesions act as a
mechanochemical transducer for changes in cell shape, tension, and integrin
binding and activation. If focal adhesions acted as the central signal for
growth and survival, then one would expect that as cell spreading increased,
the total amount of focal adhesion formation would increase. To test this
hypothesis, the total amount of focal adhesion was quantitated by measuring
total vinculin at the cell-substrate interface by using confocal microscopy.
These studies revealed that there exists a trend of increased focal adhesion
amount with cell spreading independent of the amount of ECM-cell contact
(Figure 3.18A, B), suggesting that cell spreading acts as a permissive signal for
FAC formation. Similarly, FAC signaling increased with increased cell
spreading. When FAC amount and signaling of per cell was low, it correlated
tightly with growth. However, this relationship became less significant on
patterns that induced higher FAC formation (Figure 3.18C).
The second proposal, as suggested in the discussion of this chapter, is
that changes in cell shape act to coordinate changes in the CSK and nucleus.
If this mechanical interconnection existed between cell membrane and
nucleus, we would expect increases in cell spreading to result in spreading of
the nucleus. The resulting changes in nuclear shape could either
111
Chapter 3 Appendix A
mechanically open nuclear pores to change cytoplasmic-nuclear transport, or
physically distort the nuclear matrix and DNA to alter transcription site
accessibility. We measured the projected area of the nucleus in cells attached
to all the different substrates available, and found without exception that cell
and nuclear shape increased in parallel (Figure 3.19).
112
Chapter 3 Figures
Figure 3.1. Effect of cell spreading on apoptosis. (A) Combined phase
contrast-fluorescence micrographs of human capillary endothelial cells
cultured in suspension in the absence or presence of different sized
microbeads or on a planar culture dish coated with FN for 24 hours (29). In
the highly spread cell on the 25 um bead, only the flattened DAPI-stained
nuclei is clearly visible. (B) Apoptosis in cells attached to different size beads.
The apoptotic index was quantitated by measuring the percentage of cells
exhibiting positive TUNEL staining (Boehringer-Mannheim) which detects
DNA fragmentation; similar results were obtained by analyzing changes in
nuclear condensation and fragmentation in cells stained with DAPI at 24
hours. Apoptotic indices were only determined within single cells bound to
single beads. Error bars indicate standard error of the mean. (C) Differential
interference contrast micrographs of cells plated on substrates micropatterned
with 10 or 20 um diameter circles coated with FN, using a microcontact
printing method* , or on a similarly coated unpatterned substrate (right). (D)
Apoptotic index of cells attached to different sized adhesive islands coated
with a constant density of FN for 24 hours; similar results were obtained with
bovine capillary endothelial cells**.
* Beads were coated with FN (Collaborative Biomedical, 50 ug/ml) using carbonate buffer
[Wang et al., 1993; Plopper and Ingber, 1993]. Patterned substrates containing islands coated
with FN were fabricated using a microcontact printing method [Singhvi et al., 1994; Prime and
Whitesides, 1991; Kumar et al., 1994; Mrksich and Whitesides, 1995; Mrksich et al., 1997].
Briefly, hexadecanethiol (HS(CH2)15CH3) was printed onto gold-coated substrates with a
flexible stamp containing a relief of the desired pattern. The substrate was immersed
immediately in a 2 mM solution of the tri(ethylene glycol)-terminated alkanethiol
(HS(CH2)11(OCH2CH2)30H in ethanol), which coated the remaining bare regions of gold.
When these substrates were immersed in a solution of FN, vitronectin, or type I collagen (50
ug/ml in PBS), the protein rapidly adsorbed only to the stamped regions. Antibody-coated
substrates were prepared by first immersing surfaces in a solution of goat anti-mouse IgG Fc
antibody (50ug/ml) and washed with 1%BSA/DMEM prior to immobilizing the mouse antiintegrin antibodies to integrin aVP3 (1 ug/ml; LM609, Chemicon), P1 (1 ug/ml; BD15, Biosource),
or combination of both (0.5 ug/ml each). Cells cultured on substrates with no mouse antibody or
antibodies to intracellular proteins did not adhere under these conditions.
** Human pulmonary microvascular endothelial cells (Clonetics) were cultured in EGM medium
(Clonetics) supplemented with 2% fetal calf serum, EGF (10 ng/ml) and FGF (5 ng/ml). Bovine
adrenal capillary endothelial cells were cultured in serum-free, chemically-defined medium
supplemented with FGF (5 ng/ml) [Ingber, 1990; Ingber and Folkman, 1989].
113
AB"
ETUNEL
ANuclear Fragmentation
-
60
CI)
T
0
0
0
--
10 m
10gm Bead
20Jm
25gm Bead
c
C
C,
o
Ca
Bead diameter (ptm)
Attached
D
U0 TUNEL
E Nuclear Fragmentation
60-
C
0
a
400
-
No Bead
20
0-
E
E
0
0
a
Chapter 3 Figures
Figure 3.2. Effect of spreading on cell growth and apoptosis. (A) Schematic
diagram showing the initial pattern design containing different sized square
adhesive islands and Nomarski views of the final shapes of bovine adrenal
capillary endothelial cells adherent to the fabricated substrate. Distances
indicate lengths of the square's sides. (B) Apoptotic index (% cells exhibiting
positive TUNEL staining) and DNA synthesis index (% nuclei labelled with 5
BrDU) plotted as a function of the projected cell area. Data were obtained only
from islands that contained single adherent cells; similar results were
obtained with circular or square islands and with human or bovine
endothelial cells.
115
A
5gm
20gm
304m
3g2
5040gm
-50
(D 20
c
-a
-40
15
-
-
"-
_
-30
-20
U)
.F5
0.
-0
00
4-0
0
C.
-10<
0
0
0
Adhesive Island Area (ptm)
116
a)
Chapter 3 Figures
Figure 3.3. Cell-ECM contact area versus cell spreading as a regulator of cell
fate. (A) Diagram of substrates used to vary cell shape independently of the
cell-ECM contact area. Substrates were patterned with small, closely-spaced
circular islands (center), such that cell spreading could be promoted as in cells
on larger single round islands, yet the ECM contact area would be low as in
cells on the small islands. (B) Phase contrast micrograph of cells spread on
single 20 or 50 um diameter circles or multiple 5um circles patterned as
shown in A. (C) Immunofluorescence micrographs of cells on a
micropatterned substrate stained for FN (top) and vinculin (bottom). White
outline indicates cell borders; note the circular rings of vinculin staining
which coincides precisely with edges of the FN-coated adhesive islands. (D)
Plots of projected cell area and total ECM contact area per cell (top), growth
index (middle), and apoptotic index (bottom), when cells were cultured on
single 20um circles or on multiple circles 5 or 3 um in diameter separated by
40, 10, and 6 um, respectively.
117
C
Apoptosis
2000
a
Growth
E 1500
S 1000
-
????
mCell Area
- 0 ECM Area
-
AQ
CL
20gm
500
50m
10
, , ,
25-
0
200
15-
00b
010-
0- 6C.
<
O420
:
20:40
5:10
Pattern
118
3:6
Chapter 3 Figures
Figure 3.4. Role of different integrin ligands in cell shape-regulated apoptosis.
Apoptotic indices (% positive TUNEL staining) for cells cultured for 24 hr on
unpatterned or 20 um circles coated with either FN, type I collagen (Col I),
vitronectin (VN), anti-integrin
P1
antibody, anti-integrin (xVP3 or both
combined (28).
119
VN-
Col-
FN-
1 (cx*
ocV P3 ..........
CD
--
-
,-.-
-3
0
CalC
Apoptotic Index (%)
-1-l
Chapter 3 Figures
Figure 3.5. Effect of serum in adhesion-modulated apoptosis. (A) Suspended
bovine endothelial cells were rescued from apoptosis by allowing them to
attach to single 25 um diameter beads coated with fibronectin. (B) Similarly,
human endothelial cells were rescued from apoptosis by spreading them on
higher density fibronectin. In either case the absence or presence of serum
resulted in an adhesion-dependent induction of apoptosis.
121
A
0
BCE cells
80
T
60-
X
40
-
(1)
70
0~
00~
U1
Defined Media (0% CS)
El
Full Media (10% CS)
U
Experimental Media (2%FCS)
Sl
Full Media (10% FCS)
20-
0
E
CL
Cm
LO
C'j
C/)
0
0
0
HMVEC cells
2
T
1.5-
1
-
B
-- j
0.5
-
I
0
-
0
0-
zU-
zU-
0
-j
122
-1
Chapter 3 Figures
Figure 3.6. Effect of cell spreading on apoptosis. Apoptosis of bovine (A, B) or
human (C, D) endothelial cells attached for 24 hours to FN-coated beads (A, C)
or patterned substrates (B, D) of different size. The apoptotic index represents
the percentage of cells positively identified for DNA fragmentation by either
TUNEL (Boehringer-Mannheim) or DAPI staining. Only single cells attached
to single beads or islands were scored. Error bars indicate standard error of the
mean.
123
Unrestricted
20um circles
10um circles
Suspended
Attached
45um Bead
25um Bead
10um Bead
Suspended
-
--
W-
a
7
3
-- I- - -----
0
- -- ---
MN
IF*,*, !"o
Unrestricted
.
-
--
se
Il
-4
01
n
~N0N)I
(D
0
0
Apoptotic Index (%)
. ......
m
1
20um circles
10um circles
Suspended
-......
4b.
S<
m
(0
CD
0.C zm
z
OE 0
c
Attaiched
25um Bead
20um Bead
15um Bed
1 oum Bead
Suspe nded
0
Apoptotic Index (%)
N)
ED
00
-
0
CI
o
aa
0
(D
,D
z
F m
z
E N
Ca,
Apoptotic Index (%)
0
Apoptotic Index (%)
C,
ro
o
0
CD
3
(C
C
z
F-
m
z
C
m
-z
r'n
-c
Elu
w
Cl)
z
m
H
T
w
C/)
m
--
--
.
. ----
Chapter 3 Figures
Figure 3.7. Apoptosis of bovine endothelial cells over time. Apoptosis of BCE
cells attached for either 24 or 48 hours to substrates patterned with different
sized islands coated with FN.
125
BCE Cells
50
T
400
30-
T
0
*0-~
0~
20-
0
10-r
0a,
70
V
a,
a
C,)
ci')
0
a)
E
E
=3
0
c\J
126
DP
U
24hours after plating
E
48hours after plating
Chapter 3 Figures
Figure 3.8. Differential interference contrast micrograph of a single bovine
capillary endothelial cell attached to a square island (50 um side length) coated
with FN. The central body contains the nucleus, and the raised cell boundary
suggests a ruffling edge.
127
00
Chapter 3 Figures
Figure 3.9. Effect of spreading on cell growth. Go synchronized bovine
endothelial cells were cultured onto square islands of different size for 24
hours in the presence of saturation basic fibroblast growth factor (5 ng/ml),
and assayed for entry into S-phase. The proliferation index represent the
percentage of cells that are synthesizing new DNA, as measured by either
BrdU incorporation or expression of Ki-67 or PCNA. Cells were synchronized
into Go by growing to confluence and starving in 1% serum for 24 hours prior
to the experiment. Error bars indicate standard error of the mean.
129
60
50
U BrdU
L
Ki-67
~40
4I PCNA
.2
30-
-20
zn
q
cz~~.
..
....
0
0
LO
10
co
O
0
0
N'.
0
CO
Square size (side length, gin)
130
_0
0
C
D
Chapter 3 Figures
Figure 3.10. Effect of spreading on cell growth and apoptosis in human
endothelial cells. Apoptotic index and DNA synthesis index were plotted as a
function of projected cell area. Apoptotic index represents the percentage of
cells that died within 24 hours after plating. DNA synthesis index represents
the percentage of cells that were incorporating BrdU between 20 and 24 hours
after plating (i.e., during S-phase).
131
00
3
CD
Q.
CD
0
%-.
-u
0
5000
0
4000-
3000-
2000 -
1000-
0
0
0
DNA Synthesis (%) -0-
1
0
Apoptotic Index (%) -U-
0
0
CD
C
)
m
Chapter 3 Figures
Figure 3.11. Effect of cell spreading on the kinetics of cell cycle progression.
The percentage of cells incorporating BrdU, indicating S-phase progression,
was assayed at 4 hour intervals. The plotted times indicate the end of the 4
hour period in which BrdU was pulse incubated with cells (e.g., a 20 hour
time point represents percentage of cells incorporating BrdU from 16 to 20
hours).
133
40
(D,
U)
CL)
0~
30-
-0
0-
M-20-
C')
C')
C/)
10-
lTFmW~Yr~~ 1
z
n
0 S i
0
I 1
It
af
1
1 -
Pt Cn ( ou N(00
Time after Plating (hours)
134
1
Unpatterned
-0--.....
50um circles
0 -- --
20um circles
Chapter 3 Figures
Figure 3.12. Effect of width of nonadhesive gaps on cell spreading. Cells were
attached to FN-coated lines separated by gaps of 20, 10, or 5 um. Phasecontrast micrographs were taken 24 hours after plating.
135
>-20 urn space
-10
urn line
0 urn space
- 5' urn line
-5
urn space
-3 urn line
U np attemed
136
Chapter 3 Figures
Figure 3.13. Localization of various FAC proteins in cells spread across small,
FN-coated islands. Immunofluorescent staining of vinculin, paxillin, talin,
phosphotyrosine, FAK, and integrins a5s1 and cxVp3. Cells were plated for 6
to 24 hours, permeabilized with a buffer containing 0.5% Triton X100 [CSK+,
Plopper and Ingber, 1993], and fixed for 20 minutes in 4% paraformaldehyde
in PBS.
137
VINCULIN
TALIN
INTEGRIN a51
INTEGRIN aV03
PAXILLIN
PHOSPHOTYROSINE
PHOSPHOTYROSINE
FAK
CO
Chapter 3 Figures
Figure 3.14. Localization of actin and vinculin in cells spread across small,
FN-coated islands. Immunofluorescent staining of vinculin (A, C) and
microfilaments (B, D) in cells spread on a grid of circles either 3 um in
diameter separated by 5 um spaces (A, B) or 5 um in diameter separated by 10
um (C, D).
139
VINCULIN
ACTIN
140
Chapter 3 Figures
Figure 3.15. Localization of vinculin in cells on islands coated with different
ECMs. Immunofluorescent staining of vinculin after plating cells on
substrates coated with fibronectin, collagen I, or vitronectin.
141
VINCULIN
VITRONECTIN
142
Chapter 3 Figures
Figure 3.16. Cell-ECM contact area versus cell spreading as a regulator of cell
growth. Plots of projected cell area and total ECM contact area per cell (top),
and corresponding growth of cells attached to indicated patterns of ECM.
143
3000-
U
T
e
3000
I
Y
Projected Cell Area
l
2000-
ECM Area
2000
T
I.
ci
T
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Chapter 3 Figures
Figure 3.17. Correlation of growth with geometric parameters of cell and
substrate. Projected cell area correlates tightly with percentage of cells
synthesizing DNA (r2=0.951), while total ECM contact area per cell, projected
cell perimeter, and projected cell length do not. Different plot symbols
indicate geometry of patterned ECM on the substrate.
145
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Chapter 3 Figures
Figure 3.18. Relationship between total amount of FAC per cell with cell
spreading, ECM area, and growth. (A) Plots of projected cell area and total
ECM contact area per cell (top), and corresponding total amount of FAC per
cell (bottom) attached to indicated patterned of ECM. (B) Plot of percentage of
cells synthesizing DNA as a function of total amount of FAC per cell. FAC
was quantitated by confocal microscopy measurements of fluorescence
intensity in cells immunofluorescently stained for vinculin.
147
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148
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Chapter 3 Figures
Figure 3.19. Relationship between nuclear spreading and growth. Plot of
percentage of cells synthesizing DNA as a function of projected nuclear area,
showing a high correlation (r2=0.937).
149
60r2 =0.937
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150
LO
-1
Chapter 4 Deformation of Cell and Nucleus
CHAPTER IV. MECHANICAL BASIS OF CELL
AND NUCLEAR DEFORMATION
Preface
The previous chapter contains compelling evidence that ECM regulates
cell growth and apoptosis by modulating cell shape, not by the amount of
ECM in contact with cells and not by the amount of FAC formed
intracellularly. Results also suggest that ECM molds cell shape through
physical, integrin-mediated connections to the cytoskeleton and that, as with
previous studies, cell and nuclear area appear to be coupled. We hypothesize
that the cell and nucleus responds as an integrated mechanical structure to
physical forces exerted through the ECM. Current thinking has focused on
membrane proteins and focal adhesion complexes as the major candidates for
mechanosensory apparatus, while a mechanically integrated cell raises the
possibility that any site within the cell could harbor mechanochemical
transduction machinery. In this chapter, we use micromanipulation
techniques to examine whether integrins are mechanically coupled to the
nucleus through discrete load-bearing elements within the cytoskeletal lattice.
151
Chapter 4 Deformation of Cell and Nucleus
Demonstration of mechanical connections between integrins,
cytoskeletal filaments and nucleoplasm that stabilize nuclear
structure*
ABSTRACT
We report here that living cells and nuclei are hard-wired such that a
mechanical tug on cell surface receptors can immediately change the
organization of molecular assemblies in the cytoplasm and nucleus. When
integrins were pulled by micromanipulating bound microbeads or
micropipettes, cytoskeletal filaments reoriented, nuclei distorted, and nucleoli
redistributed along the axis of the applied tension field. These effects were
specific for integrins, independent of cortical membrane distortion, and
mediated by direct linkages between the cytoskeleton and nucleus. Actin
microfilaments mediated force transfer to the nucleus at low strain, however,
tearing of the actin gel resulted with greater distortion. In contrast,
intermediate filaments effectively mediated force transfer to the nucleus
under both conditions. These filament systems also acted as molecular guy
wires to mechanically stiffen the nucleus and anchor it in place whereas
microtubules acted to hold open the intermediate filament lattice and to
stabilize the nucleus against lateral compression. Molecular connections
between integrins, cytoskeletal filaments, and nuclear scaffolds may therefore
provide a discrete path for mechanical signal transfer through cells as well as
a mechanism for producing integrated changes in cell and nuclear structure
in response to changes in extracellular matrix adhesivity or mechanics.
* Contributing authors for publication in Proceedings of the National Academy of Sciences,
USA: Andrew J. Maniotis, Christopher S. Chen, and Donald E. Ingber. CC designed some of the
experiments involving cell manipulations, and peformed the analysis of deformation behavior
of the cells. AM performed all cell manipulation experiments.
152
Chapter 4 Deformation of Cell and Nucleus
INTRODUCTION
Cells generate mechanical tension in their actin cytoskeleton (CSK) and
exert tractional forces on their adhesions to extracellular matrix (ECM)
[Ingber, 1991]. Changes in the balance of forces between cells and ECM,
induced by altering matrix flexibility or adhesivity, can change cell shape and
switch cells between growth and differentiation [Ingber, 1991; Ingber and
Folkman, 1989; Singhvi et al., 1994; Maniotis, 1991; Li et al., 1987]. The precise
mechanism by which cell shape changes influence gene expression and cell
cycle progression remains unclear. However, these regulatory effects appear
to be mediated, at least in part, by associated changes in CSK and nuclear
structure [Ingber, 1991; Ingber et al., 1987; Ingber et al., 1995; Bohmer et al.,
1996; Pienta and Coffey, 1992, Yen and Pardee, 1978]. Thus, it is critical to
understand how mechanical stresses applied to the surface membrane can
promote coordinated alterations in cell, CSK, and nuclear form.
Understanding this mechanism also could provide insight into
mechanotransduction, the process by which cells sense and respond to
external mechanical stimuli.
One explanation for integrated cell shape control is that transmembrane
ECM receptors, CSK filaments, and nuclear scaffolds are "hard-wired"
together such that a mechanical pull on the surface membrane results in
coordinated realignment of structural elements throughout this
interconnected molecular network [Ingber, 1993; Wang et al., 1993]. This
model is in direct contrast to many current models of cell mechanics which
envision the viscous fluid-like cytoplasm and surrounding elastic membrane
to be the major load-bearing elements in living cells [Fung, 1988; Evans and
Yeung, 1989; Lauffenburger and Linderman, 1993].
153
On the other hand,
1,
Chapter 4 Deformation of Cell and Nucleus
microscopic studies demonstrate structural continuity between ECM
molecules, transmembrane proteins, CSK filaments and nuclear scaffolds in
detergent-extracted cells [Berezney and Coffey, 1975; Fey et al., 1984; Osborn
and Weber, 1977]. However, the mechanical relevance of these structural
interconnections remains unclear.
We reasoned that if the CSK provides a discrete path for mechanical
force transfer from the surface to the nucleus, then we should be able to
demonstrate mechanical continuity between cell surface receptors and the
nucleus in living cells. To test this hypothesis, we used micropipettes to
micromanipulate ligand-coated microbeads (4.5 gm diameter) bound to
membrane receptors on cultured endothelial cells. When cells bind to these
beads coated with ECM ligands (e.g., fibronectin, RGD peptide) for
transmembrane integrin receptors, focal adhesions rapidly form that mediate
transfer of mechanical stresses to the internal CSK [Wang et al., 1993; Plopper
and Ingber, 1993]. In contrast, binding of beads coated with acetylated low
density lipoprotein (AcLDL), a ligand for transmembrane metabolic receptors,
neither promotes focal adhesion formation nor supports efficient stress
transfer across the plasma membrane; only a weak connection to the elastic
submembranous CSK can be detected [Wang et al., 1993; Plopper and Ingber,
1993]. In the present study, both types of surface-bound beads were pulled at a
rate (approximately 5 to 10 jm/sec) that was 10 to more than 100 times faster
than the fastest assembly rates that have been reported for CSK filaments in
mammalian cells [Condeelis, 1993; Gelfand and Bershadsky, 1991; Fuchs and
Weber, 1994]. Using this approach, we now show that cell surface integrin
receptors, CSK filaments, and nuclear scaffolds are mechanically coupled in
living cells. We also show that the mechanical properties of the cytoplasm
154
Chapter 4 Deformation of Cell and Nucleus
and nucleus depend on cooperative force transfer between all three CSK
filament systems.
MATERIALS AND METHODS
Experimental System.
Bovine capillary endothelial cells were
cultured in chemically-defined medium on glass coverslips coated with a
density of fibronectin (200-400 ng/cm 2 ; Cappel, PA) that promotes moderate
cell spreading using a carbonate buffer coating method [Ingber and Folkman
1989; Ingber et al., 1995]. More highly extended cells are much stiffer [Wang
and Ingber, 1994] and thus, are less amenable to micromanipulation.
Microbeads (4.5 gm, tosyl-activated; Dynal, Norway) were coated with
fibronectin, synthetic RGD-peptide (Peptide 2000; Telios, CA), or AcLDL
(Biomedical Technologies, MA) at 50 gg/ml, as described [Wang et al., 1993;
Plopper and Ingber, 1993], and added to cells (1 to 4 beads/cell) for 10-15 min
at 37 0 C prior to transfer to an Omega RTD 0.1 stage heating ring coupled to a
Nikon Diaphot inverted microscope. An uncoated glass micropipette was
placed alongside the surface-bound beads using a Leitz micromanipulator and
then rapidly pulled away from the cell(about 5 to 10 gm/sec), parallel to dish
surface.
The micropipettes were formed with tips approximately 1-5 gm
wide along a length of 40-100 gm. In one study, cells with bound beads were
permeabilized with 0.5% Triton X-100 in 60 mM PIPES, pH 7.4/ 25 mM
Hepes/ 8 mM EGTA/ 2 mM MgCl 2 for 2 min at 37 0 C prior to force
application. In other experiments, glass micropipettes were coated directly
with integrin ligands (fibronectin or RGD peptide) using the coating
procedure described above. ECM-coated pipettes were held in close contact
with the surface of adherent cells for greater than 5 min prior to stress
application. Control experiments confirmed that cells do not bind to the
155
Chapter 4 Deformation of Cell and Nucleus
uncoated glass pipettes in the absence of serum and that surface-bound
pipettes could be detached by adding soluble RGD-peptide. For polarization
microscopy, optics were adjusted to near complete extinction using a quarter
wave plate polarizer in conjunction with Hoya analyzers.
Analysis of Stress Transfer through the CSK. To identify the molecular
basis of force transfer between the CSK and nucleus, we harpooned the
cytoplasm 10 pm from the nuclear border using an uncoated glass
micropipette and then pulled away first 10 and then 20 gm at a rate of 5-10
gm/sec. Cells were plated in the absence or presence of 10 gg/ml nocodazole
(Noc; Sigma, MO) for 5 hr; 5 mM acrylamide (Acryl; Biorad, ) for 24 hr; 0.1
gg/ml cytochalasin D (CytoD; Sigma) for 2 hr; or 10 gg/ml Noc for 4 hr
followed by 0.1 gg/ml CytoD for 1 hr. These drug doses alter CSK mechanics
in these endothelial cells without completely blocking cell spreading [Wang et
al., 1993].
Resultant changes in deformation induced by the 10 and 20 gm
pulls were simultaneously measured using real-time videomicroscopy in
conjunction with a Macintosh Quadra 800 computer and Oncor Image
Analysis software (Oncor, CA). Nuclear strains in the direction of pull at 10
and 20 gm displacements were calculated as (d'-d)/d and (d"-d)/d,
respectively (Fig. 4.3A). Nuclear movement was defined as displacement of
the rear border of the nucleus in the direction of pull (x' and x"). Negative
lateral nuclear strain (nuclear narrowing) was calculated by measuring
changes in nuclear width perpendicular to the direction of pull.
Analysis of Mechanical Stiffness and Connectivity (Poisson's Ratio) in
the Cytoplasm and Nucleus. The stiffness (E) of any material equals stress (a;
force / cross-sectional area) divided by strain (E; change in length / initial
length). Because only induced strains were measured in this study, the
stiffness of the cytoplasm and nucleus could not be determined directly.
156
Chapter 4 Deformation of Cell and Nucleus
However, we were able to estimate the ratio of stiffnesses in the cytoplasm (c)
and nucleus (n) using the following approach (see Appendix B for a more
detailed treatment). As diagrammed in Fig. 4.4A, the ratio of nuclear to
cytoplasmic stiffness (En / EC) will equal the ratio of cytoplasmic to nuclear
strain (EC / en) measured in these regions when exposed to the same stress. If
the cell can be treated isotropically and homogeneously over short
(micrometer) distances, then the stress tensor (three dimensional stress field)
produced at any point will depend primarily on its location relative to the site
of force application. Thus, the ratio of nuclear to cytoplasmic stiffness could
be calculated by determining the ratio of induced strains measured in regions
of the cytoplasm and nucleus when placed at the same distance from the
micropipette.
Strains in the direction of pull were measured within regions of the
nucleus and cytoplasm located at the same distance from a pipette that was
pulled 10 pm toward the cell periphery; this was accomplished by respectively
placing the pipette 5 or 10 gm from the nuclear border (Fig. 4.4A). When the
pipet was placed 10 gm from the nuclear border, induced strains in the
direction of pull were measured in the cytoplasm adjacent to the pipette (0-5
pm from the tip), in distal cytoplasm adjacent to the nucleus (5-10 gm away)
and in the proximal portion of the nucleus (10-15 gm). Strain was
determined using computerized image analysis by measuring changes in the
distances between different intracytoplasmic or nucleoplasmic phase-dense
particles (e.g., vesicles, nucleoli). Identical measurements were then carried
out in similarly treated cells with a pipette placed 5pm from the nuclear
border to determine strains at the same distances (0-5, 5-10 or 10-15 pm) from
the pipette tip and hence, under similar stresses (see Appendix B for a more
157
Chapter 4 Deformation of Cell and Nucleus
complete explanation). These locations now fell in the cytoplasm adjacent to
the nucleus, in the proximal nucleus, and in the distal nucleus, respectively
(Fig. 4.4A). The ratio of nuclear to cytoskeletal stiffness was calculated by
determining the ratio of strains measured in the adjacent cytoplasm and
proximal nucleus (i.e., 5-10 gm away from pipettes placed 10 and 5 gm away
from the nuclear border, respectively). We also tested our basic assumption
by comparing strains measured within adjacent areas in the nucleus (e.g.
proximal versus distal) as well as neighboring regions in the cytoplasm (0-5
versus 5-10 pm from the nuclear border), when placed at the same distance
from the pipet. Strains measured in these regions did not differ significantly
from each other (nucleus/nucleus and cytoplasm/cytoplasm strain ratios ~ 1),
confirming that the stresses were transmitted isotropically and
homogeneously, at least over the micrometer distances we analyzed. This
approach also assumes that the strength of cell-substrate adhesions and height
of the cell in the adjacent 5 pm regions being stressed (n and c) do not vary
significantly within similarly-treated cells; electron microscopic analysis
confirmed that basal adhesions remained relatively constant and that height
values only differed by approximately 15%.
Apparent Poisson's ratios were measured in the cytoplasm and
nucleoplasm by harpooning cells 10 gm from the nuclear envelope, pulling
the pipette 5 gm away from the nuclear border and calculating the ratio of the
strain in the region along the axis perpendicular to the direction of pull
divided by the strain in the direction of pull. All strains were measured in
equal areas (9 gm 2 ) equally distant (4 to 5 gm) from both the pipet and the
nuclear border, and all displacements were of equal magnitude.
The ratio we
report here must be viewed as an "apparent" rather than absolute Poisson's
ratio because we calculate the ratio based on a two dimensional projection of a
158
Chapter 4 Deformation of Cell and Nucleus
three dimensional material in cells adherent to an underlying solid substrate.
However, variables were kept constant between measurements and thus,
relative changes in Poisson's ratios may be compared under different
experimental conditions.
RESULTS AND DISCUSSION
Mechanical stresses were applied directly to cell surface integrin
receptors by allowing cells to bind RGD-coated microbeads (4.5 pm diameter)
for 10 min and then pulling these beads laterally using uncoated glass
micropipettes and a micromanipulator. When a single RGD-coated
microbead was pulled away from the cell, the nucleus deformed and
elongated in the direction of the pull even though it was separated by many
microns from the site of force application (Fig. 4.1A,B). Coordinated changes
in intranuclear structure also were produced, as indicated by increases in the
spacing between nucleoli (Fig. 4.1A,B).
In contrast, when we pulled on
AcLDL-beads that bound to transmembrane metabolic receptors that only
physically connect to the submembranous CSK (i.e., as opposed to the focal
adhesion complex [Wang et al., 1993; Plopper and Ingber, 1993]), they detached
from the cell surface and no changes in nuclear shape or nucleolar
distribution were observed (Fig. 4.1C, D). To determine whether the observed
mechanical coupling between integrins and nuclei required changes in
diffusion-based chemical signaling or protein polymerization, we pulled
integrin-bound beads on cells after membranes and cytosolic components had
been extracted using 0.5 % Triton-X-100 (Fig. 4.1 E,F). Again, coordinated
distortion of the nucleus and nucleoli was observed, despite the absence of
membranes, surface tension, osmotic forces or ATP, thus confirming that
stress can be transferred directly through the CSK lattice.
159
Chapter 4 Deformation of Cell and Nucleus
Living cells were then pulled using glass micropipettes that were coated
directly with integrin ligands to apply stress over larger areas andto rule out
potential complications associated with bead internalization. When we
pulled fibronectin-coated pipettes that were initially bound to the cell surface
many micrometers away from the nucleus, extensive changes in nuclear
structure were observed including evagination of the nuclear boundary and
elongation of nucleoli along the principal axis of the tension field (Fig.
4.1G,H). Stress-induced molecular reorganization also could be observed
within individual nucleoli, as indicated by the appearance of birefringence
(i.e., a direct measure of multimolecular realignment) when viewed under
polarization optics (Fig. 4.1 IJ). In contrast, birefringence of nucleoli was
never observed in control cells, regardless of cell or nuclear orientation
relative to the direction of the polarizing light. Furthermore, birefringent
cytoplasmic filament bundles oriented perpendicular to the pull immediately
changed their birefringent sign and thus, reoriented (i.e., turned 900) along
the major axis of the tension field in response to stress application (Fig. 4.2
A,B). These bundles stained positively for F-actin using rhodaminatedphalloidin (not shown) and similar realignment of intermediate filaments
has been demonstrated in response to prolonged pipette pulling by electron
microscopy [Kolega, 1986]. Nuclear components might be expected to
disconnect from integrins in mitotic cells which lose most of their ECM
contacts as well as their nuclear lamina. Nevertheless, when integrins were
pulled using RGD-coated micropipettes, rotation of the mitotic spindle axis
and partial separation of chromosomes were observed (Fig. 4.2 C-F).
To analyze the molecular basis of force transfer through the cytoplasm,
we used RGD-coated micropipettes to pull on cells that were treated with
cytochalasin D (CytoD) and thus, lacked intact microfilaments. The surface of
160
Chapter 4 Deformation of Cell and Nucleus
these cells distended easily when bound integrins were stressed, at times
extending more than 100 pm in length, yet this deformation produced little
change in nuclear shape or nucleolar distribution (Fig. 4.2 GH). Thus,
deformation of the cortical membrane is not sufficient to produce the nuclear
changes that we observed in intact cells.
Because CSK modifying drugs, such as CytoD, disrupt mechanical signal
transfer between integrins and the CSK [Wang et al., 1993], we used a
"harpooning" approach to determine how stress is transmitted from the CSK
to the nucleus. The tip of an uncoated micropipette was rapidly inserted into
the cytoplasm 10 prm from the outer boundary of the nucleus and pulled first
10 and then 20 pm away toward the cell periphery (Fig. 4.3A). Pulling directly
on the CSK resulted in immediate force transfer to the nucleus as indicated by
associated nuclear extension (i.e., increase in percent nuclear strain; Fig. 4.3 B)
and movement of the nucleus in the direction of the pull (Fig. 4.3 C) as well
+
as slight narrowing of the nucleus in the perpendicular direction (e.g., -3.7
0.1 % lateral strain with 10 gm pipette displacement).
To rule out the possibility that these changes in nuclear shape were
produced indirectly by narrowing of the surrounding CSK in response to
pulling (i.e., a "sausage-casing" effect), we applied tension via pipettes placed
closer to the nuclear border. If force was transferred to the nucleus indirectly,
then tension application would result in global nuclear elongation in the
direction of the applied stress, regardless of the site of force application. In
contrast, if the CSK transfers stresses to the nucleus across direct mechanical
connections, then decreasing the distance between the pipet tip and the
nucleus should result in mechanical distortion of progressively smaller
regions of the nucleus, with greatest deformation being produced directly
along the main axis of the applied tension field. In fact, pulling closer to the
161
Chapter 4 Deformation of Cell and Nucleus
nucleus (2-4 gm) caused a small region of the nuclear envelope to protrude
locally toward the pipette in the region of highest stress (Fig. 4.3 D).
Furthermore, the nuclear border and associated cytoplasm also could be made
to indent locally by harpooning the nucleoplasm and pulling inward (Fig. 4.3
E). A discrete nucleoplasmic thread could be seen stretching from the site of
nuclear envelope indentation in these experiments (Fig. 4.3 E). These results
can not be explained by a sausage-casing effect and thus, they confirm that
tensional forces are transferred directly from CSK filaments to discrete sites
on the nuclear envelope and from there to distinct filamentous networks
within the nucleoplasm.
To examine the role of the microfilaments independently of
microtubules or intermediate filaments in nuclear shape control, cells were
plated in the presence of nocodazole (Noc) which depolymerizes
microtubules and induces formation of a contracted intermediate filament
cap at one end of the cell (Fig. 4.3 F), but permits cell spreading (Fig. 4.3 G).
The retracted intermediate filament cap can be detected by phase contrast
microscopy as a perinuclear zone of cytoplasm that excludes granules and
other organelles. When the opposite side of the cell that contained only actin
filaments was harpooned, mechanical stress was initially transferred to the
nucleus as indicated by localized evagination of the nuclear boundary (Fig. 4.3
H) as well as a small increase in nuclear strain (elongation) in the direction of
the pull (Fig. 4.3 B). But the actin network consistently ruptured in response
to larger deformations (Fig. 4.3 H,I), causing the stress to dissipate, nuclear
movement to cease (Fig. 4.3C), and the extended nucleus to retract (Fig. 4.3B).
Importantly, when these cells were pulled from the pole that retained both
microfilaments and intermediate filaments, tearing was never observed and
near normal nuclear deformation resulted (Fig. 4.3B,J). However, the absence
162
~~~LVk~2L~
-
-~---
-- M
-
-
Chapter 4 Deformation of Cell and Nucleus
of microtubules resulted in release of the normal restriction to nuclear
movement (Fig. 4.3 C) as well as a decrease in the ability of the nucleus to
resist lateral compression (-13.3 + 0.7 % lateral nuclear strain; Fig. 3 J). Similar
increases in movement (Fig. 4.3 C) and lateral compression of the nucleus (12.8 + 0.4 % strain) were produced when acrylamide was used to disorganize
the intermediate filament network in otherwise intact cells, yet nuclear
deformation in the direction of the pull was not altered (Fig. 4.3 B).
In
contrast, disruption of microfilaments with CytoD completely destabilized
nuclear shape as well as position, causing the nucleus to become more
deformable in both directions (Fig. 4.3 B; -8.1 + 0.4 % lateral nuclear strain)
and to move freely inside the cell (Fig. 4.3 C). Simultaneous administration
of CytoD and Noc resulted in additive inhibitory effects on lateral nuclear
stability (-21.5 + 3.4 % lateral strain) in addition to destabilizing nuclear
position (Fig. 4.3 C). Nevertheless, pulling on these cells that lacked both
microfilaments and microtubules (i.e., only retained intact intermediate
filaments) still produced nuclear deformation and movement in the
direction of pull, even at low strains (Fig. 4.3 B,C). Thus, the intermediate
filament network alone is sufficient to transmit mechanical stress to the
nucleus. In round mitotic cells that lack intermediate filaments (Fig. 4.2),
residual actin microfilaments and nuclear matrix scaffolds appear to preserve
coupling between the CSK and individual chromosomes [Maniotis et al.,
1997; Nickerson et al., 1992].
To explore how these CSK interconnections and associated mechanical
force transfer contribute to nuclear structure, we measured changes in the
relative stiffness of the nucleus compared to the cytoplasm (Fig. 4.4 A). This
analysis revealed that the nucleus behaved as if it were approximately 9 times
stiffer than the cytoplasm in control cells and that this difference in structural
163
Chapter 4 Deformation of Cell and Nucleus
rigidity could be completely or partially negated by interfering with either
microfilaments, intermediate filaments, or microtubules using appropriate
CSK modulators (Fig. 4.4B). Disruption of these filament systems also
decreases CSK stiffness [Wang et al., 1993] and released cytoplasmic
restrictions to nuclear movement in these cells (Fig. 4.3 C). Thus, given that
the nuclear to cytoplasmic stiffness ratio also decreases, treatment with these
CSK modulators must result in an even greater loss in nuclear stiffness.
Acrylamide could alter nuclear structure directly by compromising the
integrity of the nuclear lamina or the intermediate filament cage that
surrounds the nucleus [Hay and De Boni, 1991]. CytoD also could have direct
effects on internal nuclear scaffolds since actin has been identified within
interphase nuclei Amankwah and De Boni, 1994].
To determine where acrylamide and CytoD exert their destabilizing
actions, we compared their effects on the Poisson's ratio within the cytoplasm
versus nucleoplasm (Fig. 4.4B).
Poisson's ratio is a direct mechanical
measure of microstructural organization and connectivity within any
network (13,30). We estimated the apparent (two-dimensional) Poisson's
ratios by measuring the ratio of the strain perpendicular to the direction of
pull divided by the strain in the direction of pull. Treatment of cells with
CytoD and acrylamide each independently increased the Poisson's ratio in the
cytoplasm (Fig. 4.4 C), but not in the nucleus (nuclear Poisson's ratios of 0.60.+
0.10 versus 0.56 + 0.16, respectively). These drug treatments also did not
detectably alter force transfer within the nucleus, as measured using the
method shown in Fig. 4.3 E (not shown). Acrylamide and CytoD therefore do
not appear to directly influence the structural organization of the
nucleoplasm, rather they appear to change the mechanical stability of the
164
-1
Chapter 4 Deformation of Cell and Nucleus
nucleus by altering the ability of CSK filaments to act as molecular guy wires
within the surrounding cytoplasm.
In solid mechanics, Poisson's ratio typically ranges between 0.3 and 0.5
[Crandall et al., 1978], however, it can reach much higher values in certain
solids, such as foams and particularly in fabrics [Gibson and Ashby, 1988].
Our results suggest that the nuclei of living cells also fall into this latter
category as does the cytoplasm after CSK filament disruption. The increase in
the cytoplasmic Poisson's ratio in response to microfilament disruption may
be due to transformation of the CSK from a gel, which typically exhibits a low
Poisson's ratio, to an open lattice network (due to remaining intermediate
filaments and microtubules) that exhibits greater lateral contraction when
similarly strained. Acrylamide may increase Poisson's ratio by altering the
organization of intermediate filaments or breaking their connections with
other filament systems and thereby, altering network connectivity.
Surprisingly, loss of microtubules induced by Noc resulted in both increased
lateral contraction of the cytoplasm (i.e, increased Poisson's ratio; Fig. 4.4 C)
and enhanced narrowing of the nucleus (increased negative lateral nuclear
strain; Fig. 4.3
J)
in response to tension, even though the gel properties of the
actin CSK should become more dominant. Intact microtubules therefore may
normally act to stabilize the entire nucleo-CSK lattice against lateral
compression, in addition to holding the intermediate filament lattice open in
an extended form.
We currently have little understanding of how dynamic and integrated
changes in cell form take place or how transmission of mechanical stresses
between cells and ECM alters cell and nuclear structure, even though these
events clearly play a critical role in growth and differentiation [[Ingber, 1991;
Ingber and Folkman, 1989; Singhvi et al., 1994; Maniotis, 1991; Li et al., 1987;
165
Chapter 4 Deformation of Cell and Nucleus
Ingber et al., 1987; Ingber et al., 1995; Bohmer et al., 1996; Pienta and Coffey,
1992, Yen and Pardee, 1978; Folkman and Moscona, 1978; Ingber et al., 1986].
Our results show that cell surface integrin receptors transmit tensile stresses
that mechanically distort interconnected CSK and nucleoskeletal networks
and thereby, drive changes in cell and nuclear form in time periods much
faster than those required for polymerization. While the CSK is surrounded
by membranes and penetrated by viscous cytosol, it is this discrete
filamentous network that provides the main path for mechanical signal
transfer through the cytoplasm.
The efficiency of force transfer depends
directly on the mechanical properties of the CSK and nucleus which, in turn,
are governed by higher order cooperative interactions between
microfilaments, intermediate filaments, and microtubules acting in the
cytoplasm. The CSK also provides the cytoplasm's principal mechanical
strength whereas the mechanical properties of the surface membrane play a
relatively insignificant role in the force balances that determine cell and
nuclear form. Importantly, these results not only indicate the
inappropriateness of generalizing conventional engineering models of cell
mechanics which treat the cell as a viscous fluid surrounded by an elastic
membrane or cortical CSK [Fung, 1988; Evans and Yeung, 1989; Lauffenburger
and Linderman, 1993], they also provide direct evidence in support of more
recent efforts to mathematically describe cellular mechanics [Stamenovic et
al., 1996; Forgacs, 1995].
Quantitative analysis of the mechanical properties of the cytoplasm and
nucleus confirmed that structural interplay in the CSK is complex and that
the behaviors of these different filament systems are not simply additive or
superimposable. Actin microfilaments form a volume filling gel that
efficiently bears compression, but it does not have the strength to resist
166
Chapter 4 Deformation of Cell and Nucleus
external tension and thus, it tears at high strain. The intermediate filament
network is itself poor at resisting lateral compression, yet it efficiently resists
tension and hardens at high strains.
Similar results have been obtained
studying purified filament systems in vitro [Janmey et al., 1991]. However,
when these two filament systems are combined in living cells, a composite
higher order structure is formed that provides both load-bearing functions
with greater efficiency. Full mechanical responsiveness and structural
stability, however, requires the added presence of microtubules to locally
resist the inward contraction of the surrounding tensile CSK and thereby, to
impose an internal stress or "prestress" in this interconnected molecular
network. Cytoplasmic microfilaments and intermediate filaments also appear
to act as tensile guy wires that anchor the nucleus in place, coordinate changes
in cell and nuclear form, and provide the nucleus with its own mechanical
stiffness.
This observed dependence on discrete load-bearing elements, tensional
continuity and prestress for shape stability is consistent with a model of cell
and tissue structure that is based on tensegrity architecture [Ingber, 1993].
Tensegrity can explain how local stresses produce coordinated changes in cell,
CSK, and nuclear structure in the absence of protein polymerization or
diffusion-based signaling ([Wang et al., 1993] ; Fig. 4.1) and how different types
of CSK filaments can contribute uniquely to the overall mechanical behavior
of the cell. It also provides a mathematical basis to predict the material
properties and architectural features of living cells, independently of changes
in CSK connections [Ingber, 1993; Stamenovic et al., 1996].
This is in contrast
to percolation theory which is a mathematical method for analyzing the
importance of phase-transitions and connectivity within networks [Forgacs,
1995]. While tensegrity provides a mathematical basis for shape stability
167
Chapter 4 Deformation of Cell and Nucleus
[Stamenovic et al., 1996], percolation provides a complementary approach to
describe how the mechanical behavior of tensegrity-based networks may
change in response to alterations in CSK polymerization or cross-linking.
Taken together, these results indicate that cells and nuclei are literally
built to respond directly to mechanical stresses applied to cell surface
receptors, such as integrins. Other types of adhesion receptors that couple to
the CSK (e.g., cadherins) may exhibit similar behavior. The demonstration of
direct mechanical linkages throughout living cells raises the possibility that
regulatory information, in the form of mechanical stresses or vibrations, may
be rapidly transferred from these cell surface receptors to distinct structures in
the cell and nucleus, including ion channels, nuclear pores, nucleoli,
chromosomes, and perhaps even individual genes, independently of ongoing
chemical signaling mechanisms. As an example: neurotransmitter release
from motor nerve terminals can be detected within 10 to 20 milliseconds after
cell surface integrins are mechanically stressed [Chen, 1995]. Direct
mechanical stress transfer across these CSK linkages also may explain the
coupling between cell and nuclear shape that is observed in spreading [Ingber
et al., 1987; Pienta and Coffey, 1992] and retracting cells [Sims et al., 1992]; why
nuclear pores expand and nuclear transport rates increase when cells
physically extend [Feldherr and Akin, 1990]; and how changes in the
distribution of mechanical stresses transmitted across integrins might redirect
the axis of cell division, a process that is critical for morphogenesis of plants
[Lintilhac and Vesecky, 1984] as well as animals. This type of "mechanical
signaling" (i.e., structural coupling) could serve to coordinate, complement,
and constrain slower diffusion-based chemical signaling pathways [Ingber,
1991; Ingber, 1993] and thus, explain in part how mechanical distortion of
ECM caused by gravity, hemodynamic forces, or cell tension can change cell
168
Chapter 4 Deformation of Cell and Nucleus
shape, alter nuclear functions, and switch cells between different genetic
programs.
ACKNOWLEDGEMENTS
We thank T.N. Chen, R. Ezzell, T. Pedersen, N. Wang, K. Mi-Lee,
Fong, and M. Chicurel for helpful suggestions and
J.
J.
Folkman and R. Cotran
for their continued support. This work was supported by grants from NIH
(CA-45548) and NASA (NAG-9-430).
169
Chapter 4 Appendix B
APPENDIX B. ANALYSIS OF MATERIAL PROPERTIES
The experimental approach taken to examine mechanical properties of
the cytoplasm is treated in greater depth here. Additional definitions of terms
are introduced in this appendix that are not used in the body of the chapter.
Definitions
Let the cell be modeled as an infinite two-dimensional cytoplasm with
a circular nucleus embedded inside. The transformation of a threedimensional system into a two-dimensional one assumes that the variations
of mechanical properties in the vertical dimension (such as cell height,
adhesion to the substrate, and subcellular inhomogeneities) are incorporated
into the material properties of the model. These will be discussed in more
detail below.
Placing a pipet into a cell and pulling laterally along the dish surface
can be treated as a point deformation within the model. Let us define a
cartesian coordinate system such that its origin is the point of pipet insertion
and its X axis bissects the nucleus. Let x be the distance between the origin
and the nucleus, and r the radius of the nucleus (Figure 4.5).
Nuclear-cytoplasmic boundary conditions
To examine whether cytoplasm and nucleus are physically connected,
we observed deformations of the boundary when stressed locally. To create
local stresses in the nuclear-cytoplasmic boundary, the pipet was placed
nearby (less than r/2); in any viscoelastic material, a point deformation only
creates high stress locally, dissipating rapidly with distance from the origin of
stress. When the pipet was moved parallel to the nuclear boundary, the
170
Chapter 4 Appendix B
nucleus rotated with the pipet with no time-dependent lag, and adjacent
points within the cytoplasm and nucleus remained near each other.
Therefore, the nuclear-cytoplasmic boundary supported a no-slip condition.
Pulling the pipet away from the nucleus caused it to locally protrude in
the direction of pull (Figure 4.3D). There are several explanations for why the
nucleus and cytoplasm did not separate: Either they are physically connected
by a boundary that can bear tension, or the nucleus can be treated as a fluid
that flows into voids left by a deformed cytoplasm. The latter case would
imply that stress ceases at the cytoplasmic boundary, and the nucleus can not
bear anisotropic tensile stresses of the order experienced by the cytoplasm (i.e.,
acts like a fluid) at this time scale (seconds). To test this hypothesis, the pipet
was placed inside the nucleus, near the boundary, and pulled away from the
boundary (i.e., into the nucleus). This experiment resulted again in local
deformation of the boundary toward the pipet, with no separation of nucleus
from cytoplasm (Figure 4.3E). This finding demonstrates that nuclear
material can carry nonisotropic stresses great enough to deform the cytoplasm
into the nucleus, and that it cannot be treated as a fluid. Therefore, the
nuclear-cytoplasmic boundary is treated as a no slip boundary that physically
connects the two regions together.
In physical terms, the interactions between the nuclear envelope and
the cytoskeletal elements prevent the separation of these two structures at the
time scales observed. Whether this implies a true cross-linked interaction or
an interpenetration of polymer filaments can not be distinguished from these
experiments. If there exists a stress relaxation component resulting from the
sliding of filaments past one another, it is possible that at long time scale, the
nuclear envelope could theoretically separate from cytoskeletal polymers.
171
Chapter 4 Appendix B
Nuclear to Cytoplasmic Stiffness Ratio
Since our system could not measure applied forces or internal stresses
in this experiment, absolute values for Young's modulus (true material
stiffness) of nucleus and cytoplasm could not be calculated. However, by
examining the relative deformation of cytoplasm and nucleus to controlled
lateral stretching of the cytoplasm using a micropipet, we estimated the
relative apparent stiffness of these two compartments with respect to each
other.
To appreciate the complexity of this analysis, we first compared three
thought experiments in our two-dimensional model described above.
Consider the case of a nucleus with the identical material properties as the
surrounding cytoplasm. The deformation of this cell in response to a stretch
by a pipet would be identical to that of a model cell with no nucleus (i.e., the
space where the nucleus would be is occupied by cytoplasmic material). This
deformation would be high in magnitude near the pipet, and dissipate to a
constant value far away as the stress field distributes through the cytoplasm
(Figure 4.6A, B). Along the axis of the pipet, the principal stress and strain
would also decrease with distance from the pipet (Figure 4.6C).
Let us compare this situation with the case where the nucleus was
significantly stiffer than the cytoplasm. Based on first principles, the stiff
nucleus would deform less given the same experimental conditions (Figure
4.7A). Because the stiff nucleus deforms less than its surrounding cytoplasm,
it will experience some stress concentration (Figure 4.7B). Along the axis of
pull, because the stiff nucleus deforms less and is concentrating stress, there
may be a slight compensatory increase in cytoplasmic stress and strain near
the nucleus relative to the case with no nucleus. Within the stiff nucleus
172
Chapter 4 Appendix B
itself, stress would be increased but strain would be decreased relative to a
cytoplasm-filled nucleus, since we have imposed the same point deformation
in a stiffer system. Similar arguments can be made for a nucleus less stiff
than its cytoplasm.
We can take the results of the above thought experiment to estimate
the relative stiffness of nucleus to cytoplasm. From continuum mechanics,
the stiffness of a material (E) can be related to plane strains (e) and stresses (T)
as,
[ -v] [ x(xY)
E.[Ex
Y)
~
EYX)
= - -U(')
V
where the subscripts denote the direction of stress or strain, and v refers to
the Poisson's ratio associated with the material. Taking the first of the two
embedded equations, stiffness then could be experimentally determined as,
E= Ux(x,Y) -V -Uy(xY)
Ex(X,y)
In the case of a point deformation in an infinite plane, solutions for the stress
distribution have been derived [Timoshenko and Goodier, 1970]. These
solutions show that for Poisson's ratio ranging from 0.3 to 1, that
cX(x'O)1< 0.24; and
0xJ(x,)
v
(Tx0) <
0.07
GTx(X,O)
for stresses along the axis of pull, such that neglecting the cy term in
calculating E introduces only a maximum 7% error. In our experimental
model system, the stiffness of the stiff nucleus (En) can therefore be calculated
as the ratio of stress (axn(x,O)) to strain (Exn(x,O)) in the x-direction, measured
173
1
Chapter 4 Appendix B
inside the nucleus. The same holds true for the stiffness of the "cytoplasmic"
nucleus (subscript c).
= rxn (x,0)
E
o xc(X,0)
Exn(x,0)
EXC(X,0)
The ratio of nuclear to cytoplasmic stiffness then is a ratio of the stresses and
strains in the two types of cells, for a given x distance from the pipet.
En_
Exc(x,O)
Oxn(x,O).
Ec
Exn (x,0)
Oxc(x,0)
Because a stiff nucleus concentrates stress,
xn (x,0)
> 1
,
therefore
Uxc(X,0)
En
Ec
>
Exc(X,0)
Exn (x-,0)
Thus, if we simply measure strains along the axis of pull in the two
experiments for a given distance from the pipet, we have a lower bound, first
order estimate of how much stiffer the nucleus is relative to the cytoplasm.
In reality, we do not have a cell containing a nucleus with cytoplasmic
material properties with which to compare the deformation of real cells.
Instead, we proposed to use a cytoplasmic region of a cell as our control,
"cytoplasmic" nucleus. Suppose a pipet was place further from a nucleus, and
engineering strain (change in length normalized to initial length)
measurements were taken in the cytoplasmic region proximal to the nucleus
(Figure 4.8B). The deformation of this region should be similar to that of our
previous thought experiment with the "cytoplasmic" nucleus, since the
distance from pipet to the region of interest is held constant between the
experiments. Cytoplasmic strain as a function of distance from a pipet should
look similar whether a pipet was placed closer to the nucleus or farther,
provided that the material properties of the cytoplasm to not change
174
Chapter 4 Appendix B
dramatically as a function of distance from the nucleus. To check this,
engineering strains were measure in cells 0-5 um from pipets placed either 5
um or 10 um from the nuclear border. These strains were not statistically
different. Thus, proceeding with our analysis, we found that strains
measured in the nucleus 5-10 um from pipets (Figure 4.8A) were 9 times
lower than strains measured in cytoplasm at similar distances from a pipet
(Figure 4.8B). Taking these engineering strains as estimates of the true strain
in these experiments, equation (1) predicts that the nucleus is behaving at
least 9 times stiffer than the surrounding cytoplasm.
The apparent material stiffness of the cytoplasm and nucleus that we
estimated includes not only the actual material stiffness of these materials,
but also the 3 dimensional geometric conditions of the cell's shape in this
particular experiment. So for example, the cytoplasm becomes thinner
towards the periphery of the cell, and therefore its apparent stiffness would
decrease. In addition, the basal adhesions of a cell act as attachment points
that can dissipate stresses without deforming the cell. Therefore, in a purely
theoretical sense, one could interpret the apparently less deformable nucleus
to be due not to an actual increased stiffness in the material within the
nucleus, but rather to a 9 fold increase in cell height above the nuclear region,
or a large increase in cell-substrate adhesion at the nuclear-cytoplasmic
border. Based on electron micrographs of vertical sections of our cells, there
appears be no more than a 10% increase in a cell height from cytoplasm to
nucleus, and there appears to be no increase in immunofluorescent staining
of focal adhesions near the nucleus.
Therefore, we treat the cell cytoplasm as relatively homogenous, and
that the large difference in nuclear and cytoplasmic deformation to be
primarily a reflection of differences in material properties of nucleus and
cytoplasm.
175
-4
Chapter 4 Figures
Figure 4.1. Phase contrast (A-H) and polarization optics (I,J) views of
endothelial cells before (A,C,E,G,I) and after (B,D,F,H,J) mechanical stresses
were applied to cell surface receptors. A,B) Pulling on a single RGD-coated
microbead (4.5 gm diameter) 15 min after binding to integrins using an
uncoated glass micropipette; only 2 sec passed between (A) and (B). C,D)
Similar displacement of a surface-bound AcLDL-coated microbead. E,F)
Mechanical displacement of RGD-coated beads bound to the surface of a cell
permeabilized with 0.5% Triton X-100 prior to force application. G,H) A
spread cell before (G) and after (H) a fibronectin-coated micropipette was
bound to cell surface integrins for 5 min and pulled laterally (downward in
this view).
IJ) The same cell shown in (G,H) viewed under polarization
optics; arrowheads indicate white bifringent spots in the region of nucleoli.
The movement of the pipette is oriented downward and vertical black arrows
indicate the extent of pipette displacement in all views. (x 900)
176
N
Chapter 4 Figures
Figure 4.2. Polarization optics (A,B) and phase contrast (C-H) views of
interphase (A,B,G,H) and mitotic (C-F) cells whose integrin receptors were
mechanically stressed using surface-bound glass micropipettes coated with
fibronectin.
A) Cells exhibiting positively (white) and negatively (black)
birefringent CSK bundles aligned horizontally and vertically, respectively. B)
White arrow indicates birefringent CSK bundles which originally appeared
white in (A) that immediately changed black as they turned 900 and realigned
vertically along the axis of the applied tension field when integrins were
pulled. C-F) A series of micrographs showing a living mitotic cell. Pulling on
a fibronectin-coated micropipette bound to the cell surface resulted in
counterclockwise rotation of the spindle axis.
Partial separation of
chromosomes also can be seen in (D). Arrowheads point to the main axis of
the spindle in C & F; curved arrow indicates the direction of spindle rotation.
G) An interphase cell treated with 0.1 gg/ml CytoD for 1 hr. H) The same cell
as in (G) after tension was applied to integrins by pulling on a surface-bound,
matrix-coated micropipette (uncoated 4.5 gm diameter beads were included
only for size reference). (A,B x 700; C-G x 1,500; G,H x 900).
178
N
Chapter 4 Figures
Figure 4.3. Analysis of the Molecular Basis of Stress Transfer between the CSK
and Nucleus. A) Diagram of the method used to determine changes in
nuclear strain and movement (See Methods for details). The effects of CSKmodifying drugs on nuclear strain and movement in the direction of pull are
shown in (B) and (C), respectively; standard error was consistently less than
10% of the mean. Closed Circle/Control, absence of drugs; Closed
Square/Noc(MF), cells plated in 10 gg/ml nocodazole for 5 hr and harpooned
in the pole of the cell containing only microfilaments; Open Square/Noc(IF),
the same Noc-treated cells that were harpooned in the opposite pole
containing intermediate filaments; Open Diamond/Acryl, cells treated with 5
mM acrylamide for 24 hr; Closed Triangle/CytoD, cells treated with 0.1 gg/ml
CytoD for 2 hr; Open Triangle/CytoD+Noc, cells in Noc for 4 hr and then in
CytoD for 1 hr. D) A control cell harpooned in the cytoplasm 2-4 gm from the
nuclear border; arrow indicates a local tongue-like protrusion of the nuclear
envelope.
E) Invagination of the nuclear envelope (large arrow) in response
to harpooning the nucleoplasm. Four small arrows indicate the stressed
nucleoplasmic thread stretching to the pipet tip. Parallel
immunofluorescence (F,I, insets in G,H) and phase contrast (G,H,J) views of a
cell that was plated in the presence of Noc which induced formation of a
vimentin-positive intermediate filament cap at one pole of the cell (F)
although it did not prevent cell or nuclear spreading (G).
H) Harpooning and
pulling the intermediate filament-free pole of the cell caused nuclear
elongation in the direction of the pull, however, cytoplasmic tearing also
resulted. I) Rhodamine-phalloidin staining of cell depicted in (H) showing
tearing of the F actin-rich pole of the cell that lacked intermediate filaments.
J) Cell
in (H) after pipet was removed and used to harpoon the cytoplasm on
the opposite side of the same cell. Note extensive deformation of the nucleus
and narrowing in the perpendicular direction. Insets show nuclei stained for
DNA with DAPI.
180
H
C.)
(WI
f
a-w
OOAO
II ("4*#"k
Io*
PAW**
UK am"
UO "m8O
~z~m jIAJOW
~~EhI
imusWOA
at
181
- - I-- -
,- -
-
-.
1., 1,1W1,111il
I'm
Chapter 4 Figures
Figure 4.4. Analysis of Mechanical Stiffness and Connectivity (Poisson's
Ratio) in the Cytoplasm and Nucleus. A) Diagram of the method used to
estimate the ratio of nuclear to cytoplasmic stiffness. B) Ratio of nuclear to
cytoplasmic stiffness in cells cultured in the absence (Control) or presence of
CytoD, Acryl, or Noc (IF) using the conditions described in Fig. 4.3C.
C)
Poisson's ratio measured in the cytoplasm of control cells (Control) and in
cells treated with CytoD, Acryl, or Noc (IF). (see Methods for details)
182
0
Noc (IF)
Acryl
Cyto D
Control
Noc (IF)
Acryl
Cyto D
Control
0
0
w
-
-- q
C)
Cytoplasmic Poisson's Ratio
-I
01
Ratio of Nuclear to
Cytoplasmic Stiffness
JT1I
En
Chapter 4 Figures
Figure 4.5. Model of cell deformed with a micropipette. Two-dimensional
model (A) of cell being deformed by a micropipette inserted into the
cytoplasm and pulled away from the nucleus (B).
184
I
.. ...
I
I.........................................
..........
""O
I
cytopla
............
sm
0r
................................................
Ifl
pipefte
cytoplasm
nucleus
185
Chapter 4 Figures
Figure 4.6. Stress-strain behavior of a model cell where nucleus and
cytoplasm have equal stiffness. (A) Diagram of cell before the pipette deforms
the cell. (B) Stress (z-axis) calculated numerically as a function of x-y position,
with a point load in the x-direction imposed at the origin (analytic solutions
were taken from Timoshenko and Goodier, 1970). (C) Diagram of cell after
the pipette deforms the cell (top), and stress and strain along the axis of
pulling (bottom). Shading depicts qualitative form of stress isotherms.
Actual isotherms were calculated numerically for different ranges of stress,
Young's modulus, and Poisson's ratio which indicated little change in the
qualitative form of the isotherms. Stress and strain are shown in these
diagrams to illustrate the general trends observed in any viscoelastic material,
and do not represent a quantitative analysis for this particular experiment.
Theoretical approaches demonstrate a hyperbolic solution of stress and strain
as a function of distance from a point deformation in an infinite plane.
186
A
Y
nucleus
)(
B
ly
x
C
U,
cin
ci,
+
Distance from pipet
C
+
Distance from pipet
187
Chapter 4 Figures
Figure 4.7 Behavior of a model cell with a stiff nucleus in response to
deformation. (A) Diagram of general shape of a nucleus before and after the
pipette deforms the cell. (B) Qualitative diagram of the expected changes in
stress and strain along the axis of pulling (X-axis).
188
A
Y
nucleus
G
Mx
r n ot s t iff
stiff
B
neus
(I)
(1)
+
Distance from pipet
189
Chapter 4 Figures
Figure 4.8. Diagram of experimental approach taken to compare nuclear and
cytoplasmic responses to deformation. By initially placing the pipette at
different locations within a cell, strains could be measured in the nucleus (A)
and in the cytoplasm (B) at the same distance from a pipette.
190
A
Y
C,
x
5um
B
x
10 um
.......................
191
-A
Chapter 5 Conclusions
CHAPTER V. CONCLUSIONS
The underlying hypothesis that guided this study is that cell shape per
se regulates cell function. Based on previous work, I chose to examine the
technical potential of using SAMs of alkanethiolates on gold surfaces to
engineer micrometer-scale islands of ECM, pattern cell attachment, and
control cell shape. Using this approach, I showed that cell spreading per se
regulates an integrin- and ECM-mediated switch between growth and
apoptosis programs. In examining potential mechanisms by which ECM
regulates cell shape and growth, I noted that the cell and nucleus always
spread in a coordinated manner. We found that this coordinated spreading
results from a direct physical link between cell surface integrins and the
nucleus through the cytoskeletal lattice.
Potential mechanisms for ECM-regulation of growth and apoptosis
In this thesis, ECM-integrin interactions were found to regulate growth
and apoptosis through cell spreading. However, the molecular mechanisms
of how spreading is translated into an intracellular signal remains unclear.
The focal adhesion complex that forms intracellularly at the site of integrin
binding orients much of the signal transduction machinery of the cell
[Burridge et al., 1988; Craig and Johnson, 1996; Clarke and Brugge, 1995;
Schwartz et al., 1995;Ingber, 1993; Plopper et al., 1995; Miyamoto et al., 1995],
and appears to integrate adhesion signals with those from soluble factors.
Previous work has shown that PDGF stimulation of phosphoinositol lipid
signaling, known to be involved in cell survival and migration, requires
integrin signaling to generate the lipid substrate to the growth factor receptor
192
Chapter 5 Conclusions
complex [McNamee et al., 1993]. Maximal activation by EGF of ERK, a signal
transduction molecule involved in proliferation, also requires prior
recruitment of the EGF receptor to focal adhesion complexes by induced
integrin engagement [Miyamoto et al., 1996]. Despite this link between
integrins and soluble factors within the FAC, our data show that their
synergistic action is still insufficient to lead to the downstream signaling
events required for growth and survival.
It is possible that the FAC may also integrate mechanical signals
associated with changes in cell shape with chemical signals elicited directly by
integrin binding [Ingber, 1997]. FAC is a molecular bridge that mechanically
couples integrins and ECM to the actin cytoskeleton, and thus, provides a site
for sensing mechanical forces [Wang et al., 1993; Wang and Ingber 1994; Wang
and Ingber, 1995; Maniotis et al., 1997]. Integrin binding activates the rhoGTPase family of rho kinases, known to induce filopodial and lamellipodial
extension (i.e., spreading), stress fiber formation (i.e., cytoskeletal contractile
tension), and subsequent FAC formation [Nobes and Hall, 1995; Burridge and
Chrzanowska-Wodnicka, 1996]. Thus, integrin binding causes cell spreading
and FAC formation. In our studies, we examined the quantity and signaling
of FAC in cells spread to different degrees, and found at low spreading that
FAC formation and activity depends on the degree of spreading. Thus, FAC
formation also requires a threshhold amount of cell spreading. Taken
together, these findings suggest that integrin binding, FAC formation, and cell
spreading interplay in a positive feedback cycle that drives cells to spread to
their maximum ability.
However, it remains unclear if the observed increase in survival and
growth with cell spreading is mediated by focal adhesions themselves. The
potential importance of focal adhesion signaling for the integration of ECM193
-1
Chapter 5 Conclusions
binding and cell function comes from the recent finding that constitutive
activation of FAK kinase, a protein tyrosine kinase and a major structural
(cytoskeletal) component of the focal adhesion complex [Shaller et al., 1992;
Hanks et al., 1992], can lead to shape- and adhesion-independent cell survival
and growth [Owens et al., 1995; Frisch et al., 1996]. Perhaps the physiologic
survival and growth signal provided by cell spreading can be circumvented by
the constitutive activation of FAK kinase. In our studies, at high cell
spreading, the correlation between cell growth and FAC amount and activity
breaks down, while its correlation with spreading does not. Thus, while
integrin clustering, FAC formation, and recruitment of growth factor
receptors optimizes soluble signaling from the outside environment as seen
in early signaling events (where cell spreading appears irrelevant), cell
spreading per se appears intrinsically to provide a separate signal that allows
for later events in cell function to occur.
These changes in cell shape may be able to exert control over cell
growth and viability through many potential intrinsic mechanisms: As cells
spread on a solid substrate, the mechanical stiffness of the entire cytoskeleton
increases as a result of tension dependent structural rearrangements within
the cytoskeletal lattice that stretches from the focal adhesions to the nucleus
[Wang et al., 1993; Wang and Ingber, 1994; Wang and Ingber, 1995; Maniotis et
al., 1997; Ingber, 1997]. The resulting changes in nuclear shape could
mechanically open nuclear pores to increase cytoplasmic-nuclear transport,
allowing transcription factors or signaling molecules in and out of the
nucleus. Stretching the nucleus could also physically distort the nuclear
matrix and chromatin structure to alter transcription site accessibility
[Maniotis et al., 1997; Pienta and Coffey, 1992]. The isometric tension
generated in the cytoskeleton of spread cells also mechanically stabilizes cell
194
Chapter 5 Conclusions
and nuclear structure [Maniotis et al., 1997; Stamenovic et al., 1996]. Thus, it
is possible that the destabilization of cytoskeletal structure and increased
flexibility observed in round cells [Wang and Ingber, 1994; Wang and Ingber,
1995] may permit intracellular structural rearrangements that are lethal to the
cell. For example, loss of cell spreading may allow increased accessibility of
self-destructive enzymes to their substrates, and thus, lead to the characteristic
breakdown of cytoskeletal and nuclear architecture that is the hallmark of
apoptosis. These possibilities are consistent with our finding that cell
survival promoted by integrin $1 binding is more dependent on cell
deformation than integrin cxVP3 signaling; integrin
P1 provides
stronger
ECM anchoring to resist cytoskeletal tension [Wang and Ingber, 1995]. From
this perspective, cell adhesion to a substrate that can resist cell tension
transmitted across integrins may prevent cell death by mechanically stiffening
and stabilizing the entire nucleocytoskeletal lattice and thereby suppressing
the apoptosis structural degeneration program.
Implications of shape-regulated cell function
The existence of a geometric or mechanical control mechanism for
switching between several cell fates points to an integration of growth and
apoptosis regulation which is critical for both tissue mass homeostasis and
pattern generation. By sensing the degree of cell extension or compression,
individual cells may be able to monitor local changes in population crowding
or ECM compliance (e.g., due to enhanced remodeling or application of
mechanical stress). This mechanism for switching between life and death
could therefore serve to couple changes in cell mass with ECM extension and
thereby, tightly coordinate tissue growth and expansion. In the case of a
healing wound in which cells are lost due to injury, decompression of
195
Chapter 5 Conclusions
neighboring cells and associated cell spreading would promote growth and
repopulation of the denuded ECM until the original state of cell crowding is
restored. If cells become too dense or are forced to lose ECM contact as a result
of local overgrowth, destruction of ECM, or collapse of tissue, the cells would
be forced into a death program thereby ensuring maintenance of stable and
organized tissue form. In addition, having an intrinsic mechanism within
single cells to respond to its microenvironment, organisms can then locally
control whole societies of cells during tissue morphogenesis by regulating the
adjacent ECM environment.
Deregulation of this switching mechanism between growth and death
could lead to a piling up of cells and loss of cell-cell arrangements as is
observed during early stages of tumor formation [Ingber et al., 1981; Ingber et
al., 1985]. In fact, shape-sensitive regulatory mechanisms are lost during
malignant transformation, when cells gain the ability to survive and grow
independently of attachment to ECM [MacPherson and Montagnier, 1964;
Stoker et al., 1968] and cell spreading [Wittelsberger et al., 1981; Tucker et al.,
1981; Folkman and Greenspan, 1975]. This possibility is also supported by
recent transgenic studies which show that both tissue disorganization and
tumor formation can be induced by altering the proteolytic balance and
accelerating ECM turnover in situ [Sympson et al., 1995]. The fully defined in
vitro system we have developed could provide an important diagnostic
research tool to identify and characterize early steps in this transformation
process, by measuring shifts in the transition thresholds between apoptosis,
quiescence, and growth along the continuum of cell spreading.
Understanding the mechanism by which changes in cell shape and
cytoskeletal structure modulate apoptotic signaling by integrins also may
196
Chapter 5 Conclusions
open the window to rational design of new angiogenesis inhibitors and
hence, new forms of anti-cancer therapy.
The importance of cell shape in growth and apoptosis suggests that cell
shape must also be inherently important to many, if not all, cell processes.
Restricting cell spreading has been shown to induce the phenotypic
differentiation of many cell types, including hepatocytes [Mooney et al., 1992;
Singhvi et al., 1994], keratinocytes [Watt et al., 1988], and endothelial cells
[Ingber and Folkman, 1989]. Similarly, the density of ECM on a substrate can
control cell spreading [Ingber, 1990] and migration rates [DiMilla et al., 1993].
Perhaps, as was the case in the growth and apoptosis fields, scientists have
been witnessing the effects of cell shape their systems without consciously
controlling for this very important variable. Many effects have been observed
in in vitro systems that do not repeat in animals. Perhaps as we begin to pay
closer attention to the cellular microenvironment, many of these disparities
will vanish.
Cell-ECM interactions
Although past studies have shown that cell extension involves the
dynamic and cyclic interplay between the formation of new focal adhesions
and local cell protrusion, they have not been able to cleanly separate these two
processes from one another. Our data shows that while cells are restricted
from attaching to nonadhesive SAMs, they can spread and bridge across
narrow regions of nonadhesive SAMs that separate adhesive regions. Thus,
filopodial or lamellipodial protrusions can extend several micrometers
beyond the foremost adhesion at the cell border. This system thus could
provide a tool with which the process of cell protrusion can be examined in
the absence of new focal adhesion formation.
197
Chapter 5 Conclusions
Importantly, the ability of cells to spread across nonadhesive regions
has allowed us to fabricate substrates with small, focal adhesion-sized islands,
such that cells can spread across multiple islands but only form adhesions on
those engineered islands. Using these substrates, I demonstrated that these
adhesions act like large FACs: They contain the same structural and signaling
proteins of the FAC, and they guide and connect the actin stress fiber network.
The induction of "hypertrophic" FACs gives us the opportunity to examine
the anatomical structure of the focal adhesion as it forms and matures. By
pre-determining how cells must organize the geometry of their focal
adhesions, and observing their dynamic formation and dissolution by
automated monitoring, we can obtain longitudinal information regarding the
diversity of focal adhesion structures within a single cell. In addition, the
spatial relationship of other proteins and CSK filaments to the focal adhesion
can be easily studied in this system, since the focal adhesions lie in clearly
defined regions.
The adsorption of ECM proteins onto a hydrophobic SAM makes these
regions adhesive to specific cell surface integrin receptors. The inclusion of
this step in the fabrication process builds flexibility into the system to study
the role of specific ECMs and receptors in cell behavior: While this study
primarily focused on the fibronectin molecule, using collagen I, vitronectin,
and anti-integrin antibodies to coat substrates added a critical experiment that
demonstrated the general role of cell shape in apoptosis. We also
demonstrated that biospecific ligands (e.g., RGD peptide) can be introduced
directly into the alkanethiol before the SAM is made. The use of different
ECM molecules, fragments, and antibodies allows this system to be used with
any cell line, and should even be able to pan out specific cell types from a
suspension of mixed cell types.
198
Chapter 5 Conclusions
One general criticism of examining the role of specific ECM molecules
in cell processes is that cells rapidly degrade and redeposit ECM proteins onto
the culture substrate during the experiment. Thus, only the initial status of a
substrate is defined. To address this problem, we developed a substrate that
presented the covalently-linked RGD peptide on a protein-resistant, ethylene
glycol SAM. This substrate promoted the biospecific adhesion and spreading
of cells, while preventing the cell from depositing its own ECM molecules.
Thus, we can now investigate the role of integrin-RGD binding, independent
of other cell-ECM interactions, in cell function over a period of days. By
exchanging the peptide fragment, different ECMs also can be studied. In
addition, this system provides a baseline for studies that focus specifically on
the role of ECM turnover in cell processes, a field that has only recently
gained attention.
Potential for SAMs in engineering the cellular microenvironment
I have shown that SAMs can be used to generate surfaces containing
micrometer-scale regions that either promote or resist cell attachment. The
surfaces can be flat or contoured. The contoured surfaces themselves can be
generated with SAM technology. The adhesive regions can be tailored to
present specific ECM molecules or fragments, and even to prevent cells from
depositing their own adhesion molecules across the surface. While these
approaches were developed specifically to ask how shape regulates cell
function, their application to general science is quite broad, even with no
further advances in this technology.
The ability to place single cells into a grid opens up the possibility to
treat each cell as a single, isolated culture. Since the cells are restricted from
moving, they can be individually identified by their location within a grid,
199
Chapter 5 Conclusions
are prevented from forming colonies, and can be monitored separately. The
ability to monitor single cells longitudinally in a study, rather than an entire
population, could change our understanding of the diversity of biological
responses. For example, when one measures an increase in a particular
enzyme in a population, does every cell respond with a similar increase, or do
only some cells respond dramatically while others do not? The current
categorization of cell types is based on a relatively crude understanding of
their biology, and perhaps this new approach could revolutionize this field.
In addition to examining single cells, groups of 2 or 3 cells could be cultured
in small regions, again analogous to "micro" cultures. By changing the
geometry of these culture islands, the cell-cell interactions could be precisely
engineered. Additional complexity can be introduced by mixing different cell
types in such a system. In vivo, the interactions between multiple cell types is
critical for tissue biology (e.g., T-cells, B-cells, and macrophages in immunity;
endothelial cells, smooth muscle cells, and platelets in atherosclerosis;
hepatocytes, ito cells, and endothelial cells in hepatotoxicity and cirrhosis).
As basic biological processes become better defined, patterned substrates
will become an essential tool in the research and development of engineered
tissues. It has been demonstrated that cell function is regulated by cell shape,
specificity of cell-substrate interactions, cell-cell interactions, multicellular
tissue organization, and access to soluble factors. At all stages of tissue
engineering development, from basic research to geometric optimization to
final design, the defined patterning of cells will be critical. The development
of contoured substrates adds additional degrees of freedom to this end. These
surfaces increase bulk cell density of a substrate, add cell-cell interactions, and
provide spatial cues (e.g., contact guidance) that otherwise could not exist on a
flat substrate.
200
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ACKNOWLEDGMENTS
I would like to first thank my advisor, Don Ingber, for his dedication to
and confidence in my work. He has given me the education to approach
problems critically, the guidance to focus my efforts, and the freedom to be
creative. I also am indebted to George Whitesides, who kindly welcomed me
into his laboratory. His mentoring continues to challenge me to see things in
a broader perspective. Martha Gray and Doug Lauffenburger, the remaining
members of my thesis committee, who were always open and accessible
despite their incredibly demanding lives, provided invaluable insights that
focused my priorities on the most critical issues throughout the thesis.
I am grateful to several informal mentors, especially Martin Hemler,
Ning Wang, and Judah Folkman, whose thoughtful conversations have left
deep impressions on my paradigms for scientific progress; and I give many
thanks to many colleagues for having made this whole experience both an
intellectual adventure and a fun six years, especially members of the Ingber
laboratory; Milan Mrksich, Carmichael Roberts and Emanuele Ostuni of the
Whitesides laboratory; Deirdre Crommie of the Hemler laboratory; and
Sangeeta Bhatia, Jagesh Shah, and Tobi Nagel from the Medical Engineering
and Medical Physics program.
Lastly, I would like to thank my family and friends for all their
emotional support. Mom and Dad, thank you; my strength in persevering
stands upon the constancy of your confidence and love.
216