REVIEW
published: 29 March 2021
doi: 10.3389/fcell.2021.635263
Elucidating the Biomechanics of
Leukocyte Transendothelial
Migration by Quantitative Imaging
Amy B. Schwartz 1 , Obed A. Campos 1 , Ernesto Criado-Hidalgo 1 , Shu Chien 2,3 ,
Juan C. del Álamo 1,3,4,5* , Juan C. Lasheras 1,2,3* and Yi-Ting Yeh 1,2,3*
Department of Mechanical and Aerospace Engineering, University of California, San Diego, La Jolla, CA, United States,
Department of Bioengineering, University of California, San Diego, La Jolla, CA, United States, 3 Institute of Engineering
in Medicine, University of California, San Diego, La Jolla, CA, United States, 4 Department of Mechanical Engineering,
University of Washington, Seattle, WA, United States, 5 Center for Cardiovascular Biology, University of Washington, Seattle,
WA, United States
1
2
Edited by:
Hao Sun,
University of California, San Diego,
United States
Reviewed by:
Francis Luscinskas,
Harvard University, United States
Dietmar Vestweber,
Max Planck Institute for Molecular
Biomedicine, Germany
*Correspondence:
Yi-Ting Yeh
yiyeh@ucsd.edu
Juan C. del Álamo
juancar@uw.edu
Juan C. Lasheras
jlasheras@eng.ucsd.edu
Specialty section:
This article was submitted to
Cell Adhesion and Migration,
a section of the journal
Frontiers in Cell and Developmental
Biology
Received: 30 November 2020
Accepted: 09 March 2021
Published: 29 March 2021
Citation:
Schwartz AB, Campos OA,
Criado-Hidalgo E, Chien S,
del Álamo JC, Lasheras JC and
Yeh Y-T (2021) Elucidating the
Biomechanics of Leukocyte
Transendothelial Migration by
Quantitative Imaging.
Front. Cell Dev. Biol. 9:635263.
doi: 10.3389/fcell.2021.635263
Leukocyte transendothelial migration is crucial for innate immunity and inflammation.
Upon tissue damage or infection, leukocytes exit blood vessels by adhering to and
probing vascular endothelial cells (VECs), breaching endothelial cell-cell junctions,
and transmigrating across the endothelium. Transendothelial migration is a critical
rate-limiting step in this process. Thus, leukocytes must quickly identify the most
efficient route through VEC monolayers to facilitate a prompt innate immune response.
Biomechanics play a decisive role in transendothelial migration, which involves intimate
physical contact and force transmission between the leukocytes and the VECs. While
quantifying these forces is still challenging, recent advances in imaging, microfabrication,
and computation now make it possible to study how cellular forces regulate VEC
monolayer integrity, enable efficient pathfinding, and drive leukocyte transmigration.
Here we review these recent advances, paying particular attention to leukocyte adhesion
to the VEC monolayer, leukocyte probing of endothelial barrier gaps, and transmigration
itself. To offer a practical perspective, we will discuss the current views on how
biomechanics govern these processes and the force microscopy technologies that have
enabled their quantitative analysis, thus contributing to an improved understanding of
leukocyte migration in inflammatory diseases.
Keywords: leukocyte, vascular endothelial cell, transednothelial migration, biomechanics, force microscopy
INTRODUCTION
Leukocytes encompass a diverse group of white blood cells in the immune system, including
lymphocytes, monocytes, dendritic cells, and neutrophils, which exhibit a versatile and broad range
of migratory abilities. Leukocyte migration from the bloodstream to sites of injury or infection
is a primary component of the innate immune and inflammatory responses. Functioning as first
responders, leukocytes can efficiently overcome biophysical barriers in their response to proinflammatory stimuli, including the vascular wall and dense three-dimensional (3-D) extravascular
spaces. This efficient pathfinding is essential for leukocyte trafficking and provides potential
therapeutic targets for immune-related and inflammatory diseases.
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(Ley et al., 2007; Muller, 2014; Nourshargh and Alon, 2014;
Vestweber, 2015).
The scope and speed of the innate immune response are
primarily dictated by transendothelial migration (TEM). The
endothelium is formed by a monolayer of vascular endothelial
cells (VECs) lining the vessel walls and functions as a
physical barrier between the circulation and the underlying
interstitial tissue. During TEM, leukocytes adhere to the VECs,
transmigrate across the endothelium, and cross the vascular
basement membrane to the extravascular space (Ley et al.,
2007; Muller, 2014; Nourshargh and Alon, 2014; Vestweber,
2015; Figure 1). Pro-inflammatory stimuli such as TNF-α and
IL-8 can activate both leukocytes and VECs to initiate TEM
at the affected site (Middleton et al., 1997; Chandrasekharan
et al., 2007). Circulating leukocytes bind to selectin molecules
on the VECs via counter glycoprotein ligands, beginning a
rolling and adhesion process (Kansas, 1996). Immobilized IL8 chemokines on inflamed endothelial surfaces switch the
leukocytes’ integrins LFA-1 and VLA-4 to high-affinity states,
triggering the transition from rolling to firm adhesion and
lateral migration, followed by direct TEM. The concomitance
of high-affinity states in leukocyte integrins and increased
expressions of ICAM-1 and VCAM-1 on inflamed VECs
promotes cellular contractile forces, which regulate junctional
integrity, endothelial permeability, and ultimately leukocyte
trafficking (Cook-Mills and Deem, 2005; Stroka and ArandaEspinoza, 2010). TEM occurs via one of two routes: at endothelial
adherens junctions (paracellular migration) or through the VEC
itself (transcellular migration). Although the factors governing
route selection are not fully understood, both in-vitro and
in-vivo experiments have demonstrated that the paracellular
route is preferred, accounting for 90% of TEM events (Muller,
2003; Schulte et al., 2011; Woodfin et al., 2011). It remains
unclear whether and how leukocytes probe the endothelium to
find permissive sites for TEM and how leukocytes coordinate
the force generation with VECs to facilitate their passage
across the monolayer.
TEM involves several physical interaction cascades between
leukocytes and VECs, characterized by the sequences of
motions happening at the interfaces between the two cell
types. Receptor-ligand interactions govern leukocyte TEM by
modulating cellular functions, as mentioned above. For example,
the activation of cell surface proteins triggers cytoskeletal
rearrangements leading to increased cellular contractility and
force transmission between the leukocytes, VECs, and the
substrate. In this regard, TEM can be viewed as a biomechanically
regulated process with contributions from both leukocytes
and VECs. Recent advances in microfabrication, microscopy,
and quantitative analysis allowed researchers to measure the
mechanical forces involved in leukocyte-VEC interactions,
contributing to delineating their roles in deciding the TEM route
and driving cell motion. This review primarily focuses on two
phases that play a determinant role in leukocyte trafficking:
(1) adhesion and probing, and (2) direct TEM. Furthermore,
we discuss current advances in force microscopy techniques
for each phase and applications of force measurements in
elucidating biomechanical mechanisms of leukocyte TEM.
For additional background information on the biology of
leukocyte TEM, we recommend previous reviews on this topic
Frontiers in Cell and Developmental Biology | www.frontiersin.org
BIOMECHANICS OF LEUKOCYTE
ENDOTHELIAL ADHESION AND
PROBING
Almost immediately in response to pro-inflammatory signals,
circulating leukocytes roll on the endothelial monolayer and
then attach firmly to it (Figure 1A). Subsequently, they crawl
over the endothelium using integrin-dependent adhesions.
These interactions allow leukocytes and VECs to communicate
by well-regulated surface receptors and their counter ligands
on the opposing cell membrane. For example, the rolling
step is mediated by rapid interaction between leukocyte
selectin and P-selectin glycoprotein ligand-1 and endothelial
P- and E-selectins (Alon et al., 1995; Lawrence et al.,
1997; da Costa Martins et al., 2007; Hidalgo et al., 2007).
Chemokine-induced integrin activation facilitates firm adhesion,
spreading, crawling, and TEM by strengthening the leukocyteVEC bond via leukocyte integrins (CD11/CD18, VLA-4)
and their counter ligands, i.e., the adhesion molecules on
VECs (e.g., ICAM-1, VCAM-1). This cascade of interactions
has been characterized using specific blocking antibodies,
pharmacological manipulations, and genetic perturbations to
demonstrate each molecule’s role and downstream signaling
effects in vitro and in vivo (Berlin et al., 1995; Huo et al.,
2000; Singbartl et al., 2001; Chesnutt et al., 2006). This
interaction cascade is also highly mechanically regulated. For
example, during leukocyte rolling, the tensile forces on selectin
catch-bonds have been shown to activate leukocyte integrins
and facilitate leukocyte firm adhesion under shear stresses
(Morikis et al., 2017).
In addition to regulating biochemical receptor-ligand
interactions, leukocytes rely on mechanical forces to identify
endothelial sites with decreased barrier function and to
burrow through the endothelium. TEM does not occur with
equal probability at all locations within the endothelium.
Rather, it happens more often across the junctions between
adjacent VECs than across the cytoplasm of single VECs.
Moreover, it is observed more frequently at the confluence
of three VECs (tricellular junctions) (Lampi et al., 2017)
and between junctions loosened by inflammatory mediators
(Schaefer and Hordijk, 2015). Following rolling and firm
attachment to the endothelium, leukocytes spread out and
initiate the protrusion and retraction of podosome-like
structures that indent on endothelial membranes (Figure 1B).
These structures are speculated to continually probe the
underlying monolayer and play a decisive role in determining
the leukocyte TEM route (Burns et al., 1997; Martinelli et al.,
2014; Schaefer and Hordijk, 2015).
Podosomes are integrin-mediated adhesion structures
observed in cells originating from myeloid tissue such as
leukocytes and osteoclasts (Calle et al., 2006). These cells,
especially leukocytes, migrate on comparatively soft substrates
like endothelial or epithelial cells and their underlying interstitial
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FIGURE 1 | (A) Leukocyte extravasation: through the presence of inflamed VECs, circulating leukocytes localize themselves in the proximity of affected tissues.
Once in range, leukocytes use carbohydrate ligands to tether themselves to VECs that express specific selectins. Once tethered, the leukocyte is then able to roll
along the endothelium by creating and breaking bonds between the selectins and carbohydrate ligands. Upon the activation of integrins into a high affinity state,
triggered by chemokines binding to leukocyte’s chemokine receptors, the leukocyte can transition into a firm adhesion state that stops the rolling and allows the
leukocyte to spread out. The leukocyte then crawls and probes the vessel wall in search of VEC hotspots through which it is then able to transmigrate. This
maneuver allows for leukocytes to breach the endothelium and basement membrane, thus permitting them to reach the affected tissue area. (B) Crawling/probing:
leukocyte-VEC interactions, through high affinity integrins coupled with their respective CAMs, allow the leukocyte to migrate laterally, with the CAMs dictating the
migration pattern of the leukocyte along the vascular wall. Furthermore, the leukocyte can convert focal adhesions to invadosome/podosomes like protrusive (ILP)
structures, which are sensory organelles that they then utilize to search for TEM hotspots. (C) The transmigratory docking structure: once a hotspot is identified, a
cluster of ICAM-1 creates a cup formation to hold on to the transmigrating leukocyte. This docking structure allows the leukocyte to transition from lateral migration
to TEM. (D) TEM (paracellular): once in position at the sides of the VEC junction, leukocytes can increase VEC contractility, disrupting the local monolayer tension
and creating strong downward pushing forces, which allow for a junctional gap to form and increase in size, and for invasion of the basement membrane. This
widened gap allows for the leucocyte to push through the junction and break cellular bonds between VECs.
tissues (Zen and Parkos, 2003; Sabri et al., 2006; Carman
et al., 2007; Hidalgo and Frenette, 2007; Cougoule et al., 2010;
Dehring et al., 2011). They develop their focal adhesions
into specialized podosomes and invadosome/podosome-like
protrusions (ILPs), all similar and highly specialized subcellular
structures, when interacting with extracellular matrices and
endothelial membranes, respectively (Martinelli et al., 2014).
High-resolution microscopy revealed that the podosome
supramolecular organization consists of a central F-actin core
surrounded by a ring of integrins and focal adhesion molecules,
including talin, vinculin, and paxillin (Pfaff and Jurdic, 2001;
Vijayakumar et al., 2015; Foxall et al., 2019). The core and ring
structures are interconnected by F-actin networks containing
non-muscle myosin IIA (Pfaff and Jurdic, 2001; van den Dries
et al., 2013, 2014). F-actin polymerization in the podosome
core creates pushing forces, which are counterbalanced by the
lateral pulling forces generated through actomyosin contractility
in the cable-like structures connecting the core to adhesion
sites (Labernadie et al., 2014). This actomyosin apparatus
confers upon podosomes a highly dynamic behavior, including
fast turnover times of a few minutes (Destaing et al., 2003;
Frontiers in Cell and Developmental Biology | www.frontiersin.org
Evans et al., 2003) and the control over podosome growth,
stiffness, and protrusive force generation (Labernadie et al., 2010,
2014; Bouissou et al., 2017).
These results have raised the fundamental question of
how leukocytes utilize podosomes and ILPs to mechanosense
their microenvironment. Podosomes generate forces via their
actomyosin apparatus and sense their extracellular environments
via the integrin-based ring substructures in association
with mechanosensitive proteins, which activate downstream
mechanotransduction pathways to control various cell functions.
This process can be utilized to probe substrate topographies,
trigger extracellular matrix degradation, and sense the stiffness
of the surrounding matrix or underlying endothelium.
Studies on leukocyte adhesion to microfabricated substrates
have found that leukocyte podosomes align themselves along
substrate microgrooves (van den Dries et al., 2012). Because
conforming to 3-D microgroove topographies alters leukocyte
membrane curvature, this finding suggests that membrane
curvature could play a critical role in regulating both the
dynamics and spatial organization of podosomes. Given that
the microtopographic features of tricellular VEC junctions can
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Quantitatively Imaging Leukocyte Transendothelial Migration
seconds to ten of mins) (Carman, 2009). Furthermore, leukocytes
employ ILPs to probe the underlying VEC cytoskeleton and
preferentially migrate toward compliant areas with low F-actin
densities or loose junctions (Martinelli et al., 2014; Schaefer
and Hordijk, 2015). Also, while leukocyte migration has been
shown to vary with substrate compliance in substrates of uniform
stiffness (Stroka and Aranda-Espinoza, 2009), there is a lack of
data regarding leukocyte migration on substrates with stiffness
gradients. Existing data on other cell types, however, suggest
that integrin-mediated mechanosensing promotes durotaxis (i.e.,
migration toward stiffness gradient) rather than tenertaxis
(Choquet et al., 1997; Lo et al., 2000; Vincent et al., 2013).
Thus, there are still crucial outstanding questions regarding
mechanosensing by trafficking leukocytes and the role of ILPs in
this important cell function.
promote membrane curvature, specific subsets of protein and
lipids associating with membrane curvatures (e.g., the BAR
domain) might be involved in the podosome response to
substrate topographies during the leukocyte TEM process.
Super-resolution microscopy has revealed that the F-actin
podosome core is connected to a ventral F-actin module
bound by vinculin and a dorsal module, crosslinked by
myosin IIA and linked to other podosomes. Substrate
stiffness influences the balance between these two modules
allowing mesoscale podosome connections to collectively
switch between the explorative, degradative behavior and
the protrusive, non-degradative behavior (van den Dries
et al., 2019). This stiffness sensing behavior is crucial for
podosomes to explore spots compliant to protrusion. Moreover,
clustered podosome force oscillations have been associated
with expansion and retraction of the cell’s leading edge,
demonstrating the exploratory role of podosomes during
leukocyte migration (Kronenberg et al., 2017). The generation
of vertical protrusive forces from cancer cell invadopodia has
also been linked to cancer cell protease activity to degrade
extracellular matrices (Aung et al., 2014; Dalaka et al., 2020).
However, local disruption of integrin tensions in fibroblast
podosomes had no effect on distal podosomes (Glazier et al.,
2019), implying that collective podosome mechanosensing
may be cell-type dependent and/or more complex than
currently understood.
Given that integrins are a primary structural component of the
podosome ring, chemokines play an essential role in podosome
formation by promoting the high-affinity state of leukocyte
integrins (Carman et al., 2007; Shulman et al., 2009). Immobilized
or soluble chemokines bind to their receptors on leukocyte
surfaces to regulate both actin polymerization at the core and
integrin activation at the ring and promote the initiation of
specific podosome architectures (Hoshino et al., 2013). However,
there is no clear evidence showing any chemokine receptors
exist on the podosome structures, and the detailed interplay
between chemokine receptors and integrins will be needed for
further investigations.
Vascular endothelial cells can modulate ILP activities
by providing different ICAM-1 dependent ligand patterns
(Andersen et al., 2016), which could influence how leukocytes
select and migrate toward TEM hotspots. Conversely, ILPs have
also been implicated in sensing VEC junctional integrity and
cytoskeletal stiffness, and modulation of these factors has been
shown to affect the TEM route (Martinelli et al., 2014). However,
the precise nature of these biomechanical interactions is far from
understood (Vestweber, 2015). Leukocyte ILPs are not just a
sensory organelle and may, in fact, have additional functions.
VECs regulate endogenous tension to maintain monolayer
integrity and it is highly suspected that adhering leukocytes
can alter this tensional balance (Yeh et al., 2018). For example,
transcellular electron microscopy imaging suggests that ILPs
may distort and bend underlying actin filaments inside of VECs
by pushing directly on them (Martinelli et al., 2014).
In addition, ILPs display different characteristics from
podosomes. In particular, ILPs on VECs have shorter lifetime
than podosomes on extracellular matrices (seconds to mins vs.
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BIOMECHANICS OF DIRECT TEM
After locating a hotspot on the endothelium, leukocytes shift
from 2-D crawling to 3-D transmigration. Paracellular TEM
is the most common route through the monolayer, mediated
by the rapid disassembly of endothelial adherens junctions
in response to an adherent leukocyte. The biomechanical
interactions between leukocytes and VECs govern three crucial
steps in this process. Specifically, mechanical forces contribute
to opening endothelial gaps by destabilizing the junctions, help
pull the leukocyte across the monolayer, and mediate the closure
of the junctional gaps after TEM. This section discusses the
currently recognized mechanisms and outlines open questions
related to these three TEM steps.
The Initiation of TEM: The
Transmigratory Docking Structure
It has long been believed that VECs may play an active role
in facilitating leukocyte TEM. VECs protrude microvilli-like
projections perpendicular to the endothelium to form a specific
“transmigratory docking structure” shaped like a cup, which can
surround and hold an adherent leukocyte (Carman et al., 2003;
Carman and Springer, 2004; Yang et al., 2005; Gerard et al.,
2009; Teijeira et al., 2013). These structures are ICAM-1 or
VCAM-1 enriched after actively binding to leukocyte integrin
LFA-1 and VLA-4 (Barreiro et al., 2002; Carman and Springer,
2004; van Buul et al., 2007). Initially speculated to inhibit TEM
(Carman et al., 2003), the docking structure is now understood
to play an essential role in guiding leukocytes through the
initial stages of transmigration (Carman and Springer, 2004).
High-resolution time-lapse 3-D imaging has shown that ICAM1 clusters appearing at docking structures during early TEM
remain detectable surrounding the transmigrating leukocyte
through the late stages of TEM (Carman and Springer, 2004).
Disruption of these structures correlates with a reduction in TEM
events (Carman and Springer, 2004). Of note, this imaging data
revealed that ICAM-1 protrusions and docking structures align
perpendicular to the endothelium (i.e., parallel to the direction
of TEM). This spatial organization could help orient leukocyte
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causes the endothelial gap to enlarge for accommodating the
transmigrating leukocyte (Shaw et al., 2001; Alcaide et al., 2008;
Wee et al., 2009; Heemskerk et al., 2014). There is ample
evidence that endothelial tension regulates paracellular TEM.
Manipulating endothelial contractility by soluble inflammatory
or anti-inflammatory agents such as thrombin and angiotensin
I, biophysical cues such as stiff or soft subendothelial substrates,
or by activating or inhibiting the RhoA GTPase all respectively
increased or decreased the rates of leukocyte TEM (Hixenbaugh
et al., 1997; Saito et al., 1998, 2002; Adamson et al., 1999; Carman
et al., 2003; Yeh et al., 2018).
Apart from this VE-Cadherin-mediated junctional gap
formation mechanism, the homophilic interaction between
leukocyte PECAM-1 and VEC junctional PECAM-1 also plays
a crucial role in leukocyte TEM by recruiting the lateral border
recycling compartment (LBRC) to the site of TEM (Muller, 2003).
LBRCs are networks of dynamic VEC vesicle-like membrane
invaginations at cell-cell borders transported to TEM sites by
kinesin motors along microtubules (Mamdouh et al., 2008). The
primary molecule of LBRCs is endothelial PECAM-1, although
they also contain other junctional molecules such as JAM-A
and CD99 (Mamdouh et al., 2009). The LBRC surrounds the
leukocyte, clears junctional VEC-Cadherin to open junctional
gaps, and enlarges these gaps by contributing additional
membrane material, all of which facilitates the transmigration
process (Muller, 2014). Pharmacological perturbations inhibiting
the formation or recruitment of LBRCs prevent TEM.
To complete the picture provided by the above studies,
one must consider that leukocytes are mechanosensitive cells
that exert forces during migration and invasion (Huse, 2017;
Figure 1D). Recent 3-D traction force microscopy (TFM)
studies have shown that leukocytes exert large burrowing
stresses (∼1 KPa) to invade Matrigel substrates (Yeh et al.,
2018). Simultaneous quantification of cell shape changes,
position, and 3-D force exertion during leukocyte TEM
revealed that burrowing vertical forces increase significantly
during TEM events (Yeh et al., 2018). The tangential forces
also become stronger and display a vector pattern directed
inward toward VEC junctions, which lowers VEC monolayer
tension (Yeh et al., 2018). In contrast, VEC monolayer
tension raises when endothelial gaps form in response to
mechanically inert anti-ICAM-1-coated beads. Moreover, gap
formation mediated by these beads is significantly slower
(∼120 min) than leukocyte TEM (∼10 min) (Yeh et al.,
2018). Consistent with these findings, high-resolution correlative
microscopy imaging of VEC cytoskeletal remodeling during
TEM suggests that junctional gaps can be actively generated
by leukocytes squeezing between adjacent VECs (Barzilai et al.,
2017). In particular, the stiff leukocyte nucleus has been
suggested to act as battering ram that displaces nearby VEC
F-actin stress fibers to initiate and sustain junctional gaps
(Barzilai et al., 2017).
Altogether, these studies show that junctional gap
generation for TEM requires highly orchestrated biomechanical
contributions from both leukocytes and VECs. Future studies
are required to provide additional insight on exactly how these
forces work together to promote leukocyte TEM.
integrins so that leukocytes can shift from 2-D lateral crawling
and probing to an invasive 3-D migratory behavior.
The anchoring and embracing functions of VEC docking
structures are regulated by mechanosensitive ICAM-1-triggered
signaling, including recruitment of actin-binding proteins, an
increase in F-actin assemblies, and activation of Rho-ROCK
pathways, all of which result in increased actomyosin contractility
(Yang et al., 2006; Heemskerk et al., 2014; Figure 1C). F-actin
forms two types of assemblies with distinct functions in docking
structures: (1) F-actin filaments extending ventrally from the
apical side of endothelial membranes control VEC membrane
protrusions while (2) F-actin rich cable-like structures confine
transmigrating leukocytes at the basolateral side of VECs. In
the early stages of TEM, vertically protruding F-actin filaments
and VEC membrane fingers mediated by Myosin X activity
secure adherent leukocytes (Franz et al., 2016; Kroon et al.,
2018). The formation of these protrusions in inflamed VECs is
regulated by the ICAM-1 cluster-mediated Cdc42-myosin-PAK4F-actin signaling pathway, which generates mechanical forces
to hold the leukocyte in place and subsequently pull it toward
the VECs (Kroon et al., 2018). As TEM progresses, endothelial
pores form to accommodate transmigrating leukocytes. Pore
generation is regulated by the F-actin-rich cable-like structures
(Heemskerk et al., 2016), which exert contractile forces against
transmigrating leukocytes in order to maintain endothelial
barrier functions throughout the entire TEM process and assist
gap closure after it is complete (Mooren et al., 2014). Investigators
employed inert beads coated with ICAM-1 antibodies to mimic
adherent leukocytes, engage endothelial ICAM-1 clustering,
and demonstrate the active role of VECs in TEM. These
beads triggered a VEC process reminiscent of phagocytosis,
in which VEC membrane extensions protruded to dock and
embrace the beads (Carman et al., 2003; van Buul et al.,
2010; Kroon et al., 2018). In addition, the functionalized
beads were sufficient to induce strong localized VEC cellular
traction forces (Liu et al., 2010; Yeh et al., 2018). The
mechanical stresses created during docking structure formation
and those observed during phagocytosis share common features,
suggesting similarities between phagocytosis and leukocyte TEM
(Vorselen et al., 2020a).
The Crux of TEM: Junctional Gap
Formation
Because leukocyte sizes can be more than 20 times greater
than the size of endothelial cell-cell junctions (∼10 µm vs.
∼0.5 µm), transmigration must involve precise biomechanical
coordination between leukocytes and VECs. VECs actively
respond to the leukocyte’s presence by forming gaps to
accommodate paracellular TEM. The activation of endothelial
ICAM-1 through leukocyte binding can trigger a downstream
signaling pathway that promotes cytosolic calcium-mediated
myosin activity, resulting in increased endothelial contractility.
The resulting increase in the tensile force supported by the
F-actin cytoskeleton (i.e., endothelial tension, Figure 1D) is
transmitted to VE-Cadherin, which connects F-actin to the VEC
adherens junctional complex (Arif et al., 2021). This process
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et al., 2010; Hall et al., 2013; Style et al., 2014; Polacheck and
Chen, 2016; Figures 2A–C). This technique relies on measuring
the cell-induced deformations in a continuous elastic substrate
(e.g., a hydrogel) or an array of discrete elastic elements (e.g.,
microposts) of known mechanical properties. Using these data
as inputs, a mathematical inverse problem is solved to recover
traction forces (Style et al., 2014; Polacheck and Chen, 2016).
TFM experiments have revealed that, similar to other cell types
(Hur et al., 2009; Legant et al., 2013; Alvarez-Gonzalez et al.,
2015), neutrophils and macrophages in an adherent state exert
pulling forces around the cell edge and an unstructured bulk
pushing force under the main cell body (Kronenberg et al., 2017;
Yeh et al., 2018). The observation of subtle variations in the bulk
pushing force has been attributed to the force from podosomes
(Kronenberg et al., 2017). The joint requirements of fine spatial
resolution and high temporal sensitivity to measure such minute
forces make it challenging to quantify podosome exerted forces
by TFM. Thus, specialized methods based on atomic force
microscopy (AFM) and molecular sensors have been developed
for this application. Finally, VEC contractility contributes to
building tissue-level mechanical tension in monolayers, which
is modulated in response to external stimuli such as flow shear
stresses or the presence of an adhering leukocyte (Hur et al., 2012;
Yeh et al., 2018). Motivated by reports showing that monolayer
tension is known to regulate junctional integrity and endothelial
barrier function (Tornavaca et al., 2015; Andresen Eguiluz et al.,
2017), monolayer stress microscopy (MSM) techniques have
been developed to measure this tension. Below, we discuss
techniques developed to quantify the mechanics of leukocyte
TEM. These techniques are illustrated in Figure 2 and their
specific applications, strengths, and limitations are summarized
in Table 1.
The Resolution of TEM: Junctional Gap
Preservation of endothelial barrier function requires that VEC
junctional gaps be sealed once TEM concludes. The highly
dynamic VEC F-actin cytoskeleton plays a crucial role in
this process by continually polymerizing to form lamellipodial
protrusions that make contact with neighboring VECs to signal
gap closure (Mooren et al., 2014). Intact VECs exist under
isometric tensions with contractile forces balanced by the cellmatrix and cell-cell adhesions (Charras and Yap, 2018). Gap
formation during TEM is thought to disturb monolayer tension,
triggering the formation of the F-actin lamellipodial structures
that mediate gap closure (Phillipson et al., 2008; Martinelli
et al., 2013). Concomitantly, Rho GEF Ect2- and LARG-activated
RhoA signaling promotes actomyosin contractility in the F-actinrich cable-like structures surrounding the leukocyte, similar to
a purse string closure (Heemskerk et al., 2016). Inhibition of
RhoA activity in vitro and in vivo did not affect the rate of
leukocyte TEM events but caused leukocyte-induced vascular
leaks. Moreover, a recent in vivo study found another Tie-2
receptor/Cdc42 GEF FGD5-stimulated mechanism responsible
for preventing plasma leakage during leukocyte TEM. This study
identified that platelets recruited to endothelial VWF activate
the Tie-2 receptor by releasing Angiopoietin-1, reinforcing cablelike F-actin to close the endothelial pore (Braun et al., 2020).
Further supporting the contribution of a purse string mechanism
to maintaining barrier integrity during late TEM, the tangential
traction stress patterns during gap closure are consistent with
the presence of increased hoop tension (Yeh et al., 2018).
Overall, these results indicate that F-actin-mediated signaling
is essential for regulating gap closure, although there are not
yet any direct measurements of endothelial monolayer tension
during this process.
Quantifying Subcellular Forces
Early efforts to assess the dynamics and physical properties of
podosomes combined the application of AFM, micropatterned
substrates, and correlative fluorescent microscopy (Labernadie
et al., 2010; Figure 2D). Macrophages were plated on a
glass coverslip patterned with arrays of fibrinogen squares,
encouraging podosome formation in the protein spots in order
to restrict the size of the analysis field. AFM, in which a
cantilever applies a known force to the substrate in order to
determine its stiffness, was then used to identify the location
of membrane bumps corresponding to fluorescently labeled
F-actin rich structures (Henderson et al., 1992; Lu et al.,
2008). Time series of AFM topological images enabled accurate
measurement of podosome height (mean 578 ± 209 nm) while
force-distance mapping provided a wealth of information on
podosome stiffness. Not only was the mean Young’s modulus
of podosomes found to be approximately five times higher than
surrounding regions, but rapidly acquired force-distance curves
reproducibly demonstrated periodic oscillations in podosome
stiffness (Labernadie et al., 2010). This targeted application of
AFM permitted a refined analysis of podosome structure and
function. However, it also highlighted several key limitations
of standard AFM to quantify the biomechanics of podosomes.
Specifically, the impossibility of probing the basal tip of the
QUANTIFYING THE MECHANICS OF
LEUKOCYTE ENDOTHELIAL CRAWLING
AND TRANSMIGRATION
The trans-well assay has been widely used to quantify leukocyte
TEM for over two decades, unveiling several mechanisms crucial
to this process. For example, leukocytes can actively respond to
the exogeneous chemotactic gradient applied across the VEC
monolayers such C5a or fMLP (Cooper et al., 1995). Without
the application of exogeneous chemokines, the activation of
VECs by TNF-α or IL-8 can induce expression and secretion
of chemokines resulting in leukocyte integrin activation and
the subsequent firm adhesion, crawling and transmigration
(Adams and Lloyd, 1997). The trans-well device can easily
create a chemotactic gradient via a micropore-based membrane
separating the device’s upper and the lower chambers. However,
this assay is not ideally suited to quantify the mechanics of TEM
because it is not compatible with high magnification microscope
objectives, customized substrates, and real-time imaging of most
force microscopy methods.
The most common current method to quantify cell-generated
forces is TFM (Tan et al., 2003; Wang and Lin, 2007; Fu
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FIGURE 2 | (A,B) 2-D TFM: cells are seeded on top of a hydrogel substrate containing fluorescent beads (A) or elastic micropillars (B). Traction Forces can be
measured through the imaging of the beads’ displacements or deflections of micropillars. (C) 3-D TFM (top) and 3-D MSM (bottom): 3-D TFM takes vertical direction
displacement into account by comparing 3-D interrogate boxes between the deformed and undeformed state of substrates. 3-D MSM is derived from 3-D TFM and
takes bending effects on cellular tensions into account. (D) AFM: A cantilever beam with a probing tip at one end applies a force onto the cell. (E) PFM: cells are
seeded onto a formvar substrate, which is then stretched over a mesh grid. This formvar membrane is place upside, allowing for accurate topological information
about the podosomes to be recorded through AFM. With the use of mathematical modeling, forces exerted by the podosomes can be calculated. (F) ERISM: thin
transparent gold films are placed on top of and below an ultra-soft siloxane-based elastomer. The top gold film is protein coated to allow for cell adhesion. Local
deformation caused by the cell to the elastomer form resonance fringes that are captured using established imaging modalities. (G) MT-FILM: surface
functionalization of SLB with FRET-based DNA tension probe. When the integrin force applied is greater than F1/2 , the rated force the probe can handle, the linker
arms come together and open, causing the florescence of the probe increases. (H) microsphere-based TFM: Microspheres with fluorescent beads embedded are
placed within multilayer of cells. After a few days, the cells exert compressive forces that deform the microspheres.
fabricated, compliant formvar substrate (Labernadie et al., 2014;
Figure 2E). The formvar membrane, stretched over a square
mesh grid, was seeded with cells before being mounted upside
down in the microscope so that AFM could be performed
directly on the membrane bumps caused by podosomes. The
force generated by each podosome was calculated by fitting the
height and radius of each protrusion to a mathematical model of
podosome which is in contact with the substrate, the difficulty
of localizing podosomes from profiles of apical cell height,
and, most notably, the impossibility of measuring podosome
protrusive forces.
In order to address these limitations, protrusion force
microscopy (PFM) was developed as an extension of standard
AFM to measure indentations made by cells seeded on a specially
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TABLE 1 | Summary of cellular force measurement techniques.
System requirements
Post-processing
Advantages
Disadvantages
References
Traction force
microscopy (TFM)
Figures 2A–C
Depends mostly on
substrate properties
and microscopy setup
Hydrogel, PDMS,
elastomers, fibrillar
matrices (e.g.,
collagen); Arrays of
elastic micropillars
Standard fluorescent
microscope; Confocal
microscopy for out of
plane bead tracking
and 3-D tractions
Single particle tracking,
correlation tracking,
and/or particle image
velocimetry;
Theoretical/
computational solid
mechanics analysis
Cell substrate can be
physiologically realistic
(except micropillar
arrays); Image based –
highly versatile; Simple
experimental setup and
high throughput; Can
be extended to provide
collective cellular force
measurements (e.g.,
monolayer stress
microscopy)
Requires zero force
state (except micropillar
arrays) and calibrating
substrate elastic
properties; Limited
sensitivity to vertical
forces; Fluorescence
microscopy over long
periods can cause
phototoxic effects
Tan et al., 2003; Wang
and Lin, 2007; Fu et al.,
2010; del Alamo et al.,
2013; Hall et al., 2013;
Style et al., 2014;
Polacheck and Chen,
2016; Serrano et al.,
2019
Atomic force
microscopy (AFM)
Figure 2D
Resolution depends on
the imaging force and
probe geometries;
Lateral resolution
1–1.5 nm; Vertical
resolution 0.1 nm;
Force resolution 100
pN
Mica, glass, or glass
slides modified with
Silane to enhance cell
adhesions
Piezoelectric scanner
for mounting samples;
Proper probes attached
to pliable silicon or
silicon nitride cantilever;
Laser
beam/photodiode
setup for measuring
cantilever deflection
Cantilever deflection as
a function of vertical
displacements;
Conversion a
force-versus-separation
distance curve
Probes for molecular
interactions,
physiochemical
properties, surface
stiffnesses, and
macromolecular
elasticities
Requires careful
sample preparation and
data collection;
Requires physical
contact between the
AFM probe and the
sample – cannot probe
basal structures (e.g.,
podosomes tips)
Localizing specific cell
structures (e.g.,
podosomes) by AFM
alone is challenging
Labernadie et al., 2010
Protrusion force
microscopy (PFM)
Figure 2E
The same as AFM;
Vertical resolution 10
nm; Line rate on order
of 1 Hz; Force
resolution to the order
of nN
Compliant formvar
membranes
AFM system and
fluorescence
microscopy
The same as AFM, plus
mathematical model to
infer podosomes forces
from formvar
membrane deformation
Measures protrusive
forces applied
perpendicularly to the
substrate at a single
podosome level; High
spatiotemporal
resolution
The same as AFM,
except for localizing
podosomes; Narrow
range of applications.
Labernadie et al., 2014;
Bouissou et al., 2017
Elastic resonator
interference stress
microscopy
(ERISM)Figure 2F
Displacement
resolution 2nm (limited
by surface); Temporal
resolution <0.5 s;
Lateral resolution
∼1.6 µm;
Elastic optical
micro-cavity comprized
of a layer of ultra-soft
siloxane-based
elastomer sandwiched
between
semi-transparent gold
layers
Conventional wide-field
phase contrast or
fluorescent microscopy
with a tunable light
source capable of
providing
monochromatic
illumination
Each light fringe ∼
200 nm = ≥ count
fringes to determine
size of deformations;
Conversion of forces by
utilizing substrate
mechanical properties
Unlike many TFM
methods, no zero-force
state required; No
phototoxic effects;
Versatile, and
compatible with other
microscopy methods;
Excellent vertical and
lateral force sensitivities
Experimental setup and
fabrication of ERISM
cavities are relatively
involved; 2D soft
substrate may not be
physiologically realistic
for some applications
Kronenberg et al.,
2017; Liehm et al.,
2018
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Quantitatively Imaging Leukocyte Transendothelial Migration
Cell substrate
8
Resolution (x/y/z/F/t)
Standard fluorescent
microscope; Confocal
microscopy for tracking
3-D shape
deformations
Comparison between
deformed and
undeformed states
from 3-D shape
reconstructions
Suitable for studying
forces in environments
with complex
mechanical properties,
where TFM and ESRIM
would be challenging
Intensive image
processing
requirements
Resolution is limited by
spatial distribution of
microspheres in sample
Girardo et al., 2018;
Mohagheghian et al.,
2018; Kaytanli et al.,
2020; Vorselen et al.,
2020b
a clamped membrane subject to a point force (Labernadie et al.,
2014). By increasing the thickness of the formvar sheet and thus
the substrate stiffness sensed by cells, PFM results demonstrated
that leukocyte podosomes increase their protrusive forces in
response to stiffer substrates, suggesting that podosomes have
a mechanosensing function (Labernadie et al., 2014). Periodic
oscillations in podosome protrusive forces concomitant with the
aforementioned periodic stiffness oscillations were uncovered
using time-lapse PFM, which functions by keeping the AFM
cantilever tip at a constant force in contact with the top of a
protrusion as it moves in real-time (Labernadie et al., 2014).
The similarities between periods of podosome protrusive forces
(40 ± 14s), core stiffness (37 ± 20s) as measured by PFM,
and F-actin intensities at the podosome core (44 ± 11s) as
measured by total internal reflection fluorescent imaging indicate
a correlation between the generation of oscillating protrusion
forces and the stiffness and F-actin content of the podosome core.
This correlation was validated by pharmacological perturbations,
demonstrating that both F-actin polymerization and actomyosin
contractility regulate periodic protrusion forces.
While PFM provides invaluable information about the
biomechanics of podosomes, it is a highly specialized technique.
For instance, it is not straightforward to integrate PFM
measurements of protrusive forces with long-term, whole-cell
measurements of tangential traction stresses and motility. The
emergence of elastic resonator interference stress microscopy
(ERISM, Figure 2F) now allows for quantifying wholescale lateral and protrusive cell-substrate forces, with enough
resolution to detect the minute forces exerted by podosomes
and over extended periods of time. ERISM implements
interferometric detection of cell-induced deformations in an
elastic microcavity, allowing for high sensitivity to weak forces
in a non-destructive manner such that cells can be retained on
the substrate after imaging for subsequent measurements and
assays (Kronenberg et al., 2017; Liehm et al., 2018). ERISM
measurements could in principle be analyzed to recover collective
cellular stresses such as endothelial monolayer tension. Thus,
although it cannot currently be applied to 3-D physiologically
relevant environments like, (e.g., Matrigel or collagen matrices),
ERISM constitutes a promising technique to quantify the
biomechanics of leukocyte TEM.
Pursuant to these developments, specialized microscopic
methods were developed to investigate the contributions of
specific proteins to podosome force generation. Specifically,
DNA-based subcellular tools were developed to explore the role
of integrin tensile forces in podosome formation and illustrate the
mechanical link between integrin tension at the podosome ring
and actin protrusion at the podosome core (Glazier et al., 2019;
Figure 2G). Molecular Tension-Fluorescence Lifetime Imaging
Microscopy (MT-FLIM) allows for precise, piconewton (pN)
resolution measurement of integrin tensile forces. MT-FLIM
relies on the surface functionalization of a laterally fluid, selfassembling phospholipid membrane on glass (supported lipid
bilayer, or SLB) with FRET-based DNA tension probes (Wang
and Ha, 2013; Blakely et al., 2014; Zhang et al., 2014; Brockman
et al., 2018; Glazier et al., 2019). The probes consist of a
binary DNA hairpin with an internal loop structure and two
Force resolution to the
order of nN
Microsphere-based
traction force
microscopy
Figure 2H
Hydrogel-based
microspheres;
Fabricated by water-oil
emulsions
Force threshold F1/2 is
a measure of the
applied force at which
50% of probes are
open, estimating
applied force ranges;
Force resolution to the
order of pN exerted by
individual integrins
Supported lipid bilayer
(SLB) – phospholipid
membranes; Confined
in the Z-direction but
are laterally fluid
Inverted microscopes
with perfect focus
capabilities and
appropriate lasers for
excitation; Specific
software required;
Matlab Bioformats
Toolbox and
semiautomated custom
scripts; FIJI plugins,
including
MultiKymograph and
TrackMate
Highly specific
observations of integrin
behavior and force
generation as it relates
to podosome formation
and mechanosensing
Highly technical in the
development and
implementation of
molecular tension
probes, and in
microscopy set-up
Misses forces
transmitted via
non-specific
interactions
Wang and Ha, 2013;
Blakely et al., 2014;
Zhang et al., 2014;
Brockman et al., 2018;
Glazier et al., 2019
Quantitatively Imaging Leukocyte Transendothelial Migration
Molecular tensionfluorescence
lifetime imaging
microscopy
(MT-FLIM)
Figure 2G
TABLE 1 | Continued
Resolution (x/y/z/F/t)
Cell substrate
System requirements
Post-processing
Advantages
Disadvantages
References
Schwartz et al.
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Quantitatively Imaging Leukocyte Transendothelial Migration
VECs and adherent leukocytes and play a role in TEM (Rabodzey
et al., 2008; Liu et al., 2010).
The 2-D TFM is relatively straightforward on standard
fluorescent microscopes, has excellent resolution, and can be
adapted to a wide variety of applications (Style et al., 2014;
Polacheck and Chen, 2016). However, leukocyte TEM is an
inherently 3-D process involving significant forces in the vertical
direction of invasion. TFM measurement of these vertical
forces requires more involved imaging setups and careful
postprocessing to balance resolution with phototoxic effects.
Motivated by the fact that cells generate both lateral and vertical
traction forces while adhering to and migrating over planar
substrates (Hur et al., 2009; Maskarinec et al., 2009), TFM
methods to measure these 3-D forces have been developed
(Hur et al., 2009; Maskarinec et al., 2009). These techniques
are loosely referred to as 2.5-D TFM because they provide 3-D
traction forces in the 2-D plane of cell attachment, differentiating
them from volumetric TFM experiments where cells are fully
embedded inside 3-D matrices (Legant et al., 2010). In 2.5-D TFM
experiments the substrate is the same as in 2-D TFM, but confocal
imaging is required to record 3-D substrate deformations. To
minimize the phototoxicity generated by laser radiation when
acquiring a z-stack of confocal images, del Alamo et al. developed
a methodology that inputs the 3-D deformation at the top plane
of the substrate into the solution of the 3-D elastostatic equation
(del Alamo et al., 2013), thus requiring only ∼10-slice z-stacks
(or approximately 10 µm in depth). By adding deformation data
from additional planes, this methodology can be extended to
substrates of unknown mechanical properties or used to detect
substrate degradation (Aung et al., 2014; Alvarez-Gonzalez et al.,
2017). Overall, 2.5-D TFM constitutes a powerful tool to study
the biomechanics of TEM and delineate the distinct roles of VEC
and leukocyte forces in coordinating this process.
linker arms hybridized to include (1) a Cy3/BHQ FRET pair,
which is brought into proximity when a force greater than the
tunable F1/2 threshold is applied to the probe, opening the
hairpin and (2) cyclic Arg-Gly-Asp-D-Phe-Lys (cRGD), which is
localized on the upper arm and whose depletion is a proxy for
podosome protrusive forces as measured by percentage decrease
in fluorescence. Application of a force greater than or equal to
F1/2 will open the probe, increasing both fluorescence intensity
and lifetime (Glazier et al., 2019). Using tunable F1/2 DNA
probes, MT-FLIM enabled the identification of a narrow range
of integrin mediated forces and used time-course imaging to
establish a picture of the spatiotemporal evolution of podosome
force generations at a subcellular level.
Quantifying Single Cell Forces
Two-dimensional (2-D) TFM, which measures the lateral
traction forces parallel to the surface of cell attachment, was first
applied by Dembo and Wang to fibroblasts migrating on flat
substrates (Dembo and Wang, 1999). This technique has been
widely used to quantify the biomechanics of leukocyte adhesion
and crawling on substrates of varying stiffness of 0.05–8 kPa,
providing traction stress maps with a lateral resolution of 1–
5 µm (Rabodzey et al., 2008; Liu et al., 2010; Yeh et al., 2018).
The experimental assay used in this type of experiments has
been refined over the years and consists of a protein-coated gel
(e.g., polyacrylamide) containing fluorescent microspheres near
its surface (Figure 2A). Substrate deformations are quantified
from the movement of the fluorescent beads by image correlation
techniques, using reference images obtained after treating the
cells (e.g., by trypsin) to detach from the substrate or after
the cells move away from the region of interest. The partial
differential equation of elastic equilibrium for the substrate (i.e.,
the elastostatic equation) can be solved to determine the traction
stresses from the measured deformations using a variety of
inversion and regularization procedures (Schwarz et al., 2002;
Sabass et al., 2008). Notably, the computationally efficient Fourier
analysis of the elastostatic equation proposed by Butler et al.
(2002) makes it possible to calculate traction stresses from raw
microscopy images virtually in real time.
In micropost-based TFM, protein-coated arrays of
microscopic pillars made from the deformable elastomer
polydimethylsiloxane (PDMS) serve as a substrate (Figure 2B).
Cells attached to these arrays induce pillar deflections that
can be converted into force vectors using the known height,
width, and material properties of the pillars (Tan et al., 2003;
Fu et al., 2010; Polacheck and Chen, 2016). In principle,
micropost-based TFM does not require a reference image since
the undeflected positions of the pillars are known theoretically
(Lemmon et al., 2009), which is advantageous. On the other
hand, microfabrication and imaging constraints limit the spatial
resolution of this technique (Amato et al., 2012). Furthermore,
the highly particular substrate topography created by the
micropost arrays differs from physiologically relevant scenarios
and can affect cell adhesion. Pioneering micropost-TFM studies
of leukocyte endothelial crawling showed that VECs exert
increased tangential forces in response to a firmly adherent
leukocyte, uncovering that biomechanical interactions between
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Quantifying Tissue-Level Cell Forces
In order to maintain homeostatic barrier function, VECs must
regulate their monolayer tension to balance the biomechanical
stability of cell-cell junctions and cell-substrate adhesions.
This balance prevents cell adhesion forces from tearing the
endothelium apart or detaching it from the substrate (Charras
and Yap, 2018). During inflammatory responses, both the
magnitude and fluctuations of VEC monolayer tension tend to
increase, leading to inherently unstable junctions (Yeh et al.,
2018). Measurements of endothelial monolayer tensions over
time suggest that the rate of leukocyte TEM correlates with
tension fluctuations, which can be actively induced by leukocytes
at TEM sites (Yeh et al., 2018). Recent mathematical models
also support the idea that monolayer tension fluctuations
play a crucial role in monolayer integrity and leukocyte
TEM (Escribano et al., 2019). However, joint quantitative
measurements of endothelial traction forces, monolayer tensions,
and the forces exerted by leukocytes on the endothelium
are still scarce.
In comparison to AFM or TFM, the development of
experimental, image analysis, and computational tools to
quantify collective cellular forces has been recent. A salient
technique is MSM, an extension of TFM that quantifies the
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Quantitatively Imaging Leukocyte Transendothelial Migration
Finally, it is important to note that, although shear stress is an
important regulator in inducing leukocyte TEM (Cinamon et al.,
2001), most existing force microscopy studies of leukocyte TEM
have not considered shear flow conditions so far. While it requires
a more complicated experimental setting, it is not unfeasible to
consider shear flow in force microscopy assays and, in fact, there
are established TFM and FRET imaging assays that include shear
flow (Hur et al., 2012; Perrault et al., 2015; Heemskerk et al.,
2016). Future efforts shall exploit these tools to study how shear
affects the mechanics of leukocyte TEM by directly measure the
forces involved in the process.
collective distribution of intracellular stresses in thin confluent
cell layers (Trepat et al., 2009). Of note, MSM can measure
monolayer tension, which is the tensile intracellular stress. Most
MSM methods calculate intracellular stresses in the monolayer
from 2-D measurements of in-plane traction stresses by applying
Newton’s third law in differential (Trepat et al., 2009) or
integral (Hur et al., 2012) form after averaging across monolayer
thickness. The differential formulation provides significantly
better lateral spatial resolution than the integral one, although it
relies on a number of simplifying assumptions such as linearly
elastic material behavior, constant elastic moduli, and known
Poisson ratio. For the most part, these assumptions do not seem
to severely affect the recovered intracellular stresses (Trepat et al.,
2009; Tambe et al., 2013). Furthermore, they can be relaxed
using particle dynamics simulations (Zimmermann et al., 2014)
or Bayesian inference analyses (Nier et al., 2016). However,
these 2-D approaches do not consider that cell monolayers
respond to not only in-plane tangential stresses, but also outof-plane stresses that induce monolayer bending. Confluent
VECs adhering to soft substrates can generate strong out-ofplane traction stresses that bend the monolayer, particularly near
the monolayer edges (Serrano et al., 2019). The invasive forces
exerted by leukocytes during TEM also cause monolayer bending,
leading to significant perturbations in intracellular tension (Yeh
et al., 2018). To overcome the limitations of 2-D MSM, Serrano
et al. recently developed a new MSM method (Serrano et al.,
2019) that uses 2.5-D TFM measurements to calculate the
contributions of lateral and bending deformations to monolayer
tension (Figure 2C).
An inherent difficulty in quantifying the biomechanics of
TEM is to tease out the forces exerted by the leukocytes
from those exerted by the VECs. To this end, ICAM-1
antibody-coated polystyrene beads mimicking firmly adherent
leukocytes have been used in combination with TFM and
MSM methods (Liu et al., 2010; Yeh et al., 2018; Serrano
et al., 2019). Given that the microbeads are mechanically
inert, these experiments provide useful information about
how VECs regulate monolayer tension during TEM. However,
polystyrene beads are rigid, which makes it impossible to
quantify the forces that VECs exert on the beads, and the
recent development of methods to quantify mechanical forces
in vivo via deformable hydrogel microspheres could overcome
this limitation. Inspired by the seminal use of functionalized
oil droplets to measure anisotropic stresses within 3-D cell
aggregates (Campas et al., 2014), emerging microfabrication
methods can now produce deformable hydrogel-based spherical
force sensors, with sizes ranging from a few µm up to hundreds of
microns (Girardo et al., 2018; Mohagheghian et al., 2018; Kaytanli
et al., 2020; Vorselen et al., 2020b; Figure 2H). These elastic
microspheres can be employed to study cellular forces induced
by specific ligand-receptor interactions, known as microspherebased TFM. Comparisons between the deformed and the stressfree state of microspheres allows force measurements to be
performed using analysis methods similar to those employed
in TFM. These techniques are anticipated to generate novel
quantitative insights about the mechanical progression of VEC
docking structure formation during the initial TEM process.
Frontiers in Cell and Developmental Biology | www.frontiersin.org
CONCLUDING REMARKS
Leukocyte recruitment is a hallmark of all acute and chronic
inflammatory disorders. Understanding leukocyte TEM to
inflammation sites could help identify therapeutic targets to
boost immune defense and minimize inflammatory tissue
damage. Currently, medical treatments of chronic inflammation
employ general immune suppressors, which have numerous
adverse side effects. This lack of success is attributed to
the complexity of and the multitude of redundancies and
interdependencies in the molecular pathways involved. Although
many molecules, some of which are discussed above, have
been implicated in TEM, their specific roles remain elusive.
In particular, we still know little about how VECs and
leukocytes orchestrate pathfinding and localized endothelial
barrier modulation. These processes can depend on direct
receptor-ligand biomechanical interactions and more intricate,
collective mechanosensitive pathways. Their understanding
requires advanced experimental techniques to detect subcellular,
single-cell, and tissue-level deformations during leukocyte TEM,
along with the corresponding development of computational
models to analyze data streams of increasing complexity and size.
During the past ten years, the progress in the development of
soft materials have brought in vitro assays close to reproducing
physiological microenvironments under controlled conditions.
In parallel, advances in 3-D imaging and force quantification
have opened a window to data of unprecedented richness and
quality. Indeed, these methodologies have significantly extended
our current understanding of leukocyte trafficking. The current
technological frontier is methods to allow investigation of these
biomechanical events in vivo. Of note, emerging microscopy
techniques such as two-photon, light sheet, and super-resolution
microscopy will most likely play a transformative role in bridging
the gap going from in vitro conditions to more realistic animal
models. Heavy interdisciplinary efforts involving engineers,
physicists, and mathematicians will certainly be required to
overcome these formidable challenges.
AUTHOR CONTRIBUTIONS
AS, EC-H, SC, JÁ, JL, and Y-TY wrote the manuscript. AS and
OC composed the figures and table. All authors contributed to
the article and approved the submitted version.
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Association Grant 18CDA34110462 (to Y-TY), Fundación
Bancaria ‘la Caixa’ (ID 100010434), and the partial financial
support through a ‘la Caixa’ Fellowship LCF/BQ/US12/
10110011 (to EC-H).
FUNDING
This work was supported by National Institutes of Health
Grants GM084227 (to JL and JÁ), American Heart
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Conflict of Interest: The authors declare that the research was conducted in the
absence of any commercial or financial relationships that could be construed as a
potential conflict of interest.
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