I the Role of Mechanically Gated Ion Channels in Dorsal Closure During Drosophila Morphogenesis by Ginger Hunt

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The Role of Mechanically Gated Ion Channels in Dorsal Closure During Drosophila

Morphogenesis

Ginger Hunter

Department of Biology Duke University

Date:______Approved:

______Dan Kiehart, Supervisor

______Terry Lechler

______Danny Lew

______Dave McClay

______Zhen‑Ming Pei

Dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department of Biology in the Graduate School of Duke University

2012

ABSTRACT

The Role of Mechanically Gated Ion Channels in Dorsal Closure During Drosophila

Morphogenesis

Ginger Hunter

Department of Biology Duke University

Date:______Approved:

______Dan Kiehart, Supervisor

______Terry Lechler

______Danny Lew

______Dave McClay

______Zhen‑Ming Pei

An abstract of a dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department of Biology in the Graduate School of Duke University

2012

Abstract

Physical forces play a key role in the morphogenesis of embryos. As cells and

tissues change shape, grow, and migrate, they exert and respond to forces via

mechanosensitive proteins and protein complexes. How the response to force is

regulated is not completely understood.

Dorsal closure in Drosophila is a model system for studying cell sheet forces

during morphogenesis. We demonstrate a role for mechanically gated ion channels

(MGCs) in dorsal closure. Microinjection of GsMTx4 or GdCl3, inhibitors of MGCs,

blocks closure in a dose‑dependent manner. UV‑mediated uncaging of intracellular Ca2+

causes cell contraction whereas the reduction of extra‑ and intracellular Ca2+ slows

closure. Pharmacologically blocking MGCs leads to defects in force generation via

failure of actomyosin structures during closure, and impairs the ability of tissues to

regulate forces in response to laser microsurgery.

We identify three genes which encode candidate MGC subunits that play a role

in dorsal closure, ripped pocket, dtrpA1 and nompC. We find that knockdown of these

channels either singly or in combination leads to defects in force generation and cell

shapes during closure.

Our results reveal a key role for MGCs in closure, and suggest a mechanism for

the coordination of force producing cell behaviors across the embryo.

Contents

Abstract ...... iv

List of Tables ...... viii

List of Figures ...... ix

Acknowledgements ...... xi

1. Introduction ...... 1

1.1 Why is mechanotransduction relevant? ...... 1

1.2 Actomyosin contractility ...... 3

1.3 Force sensing: outside‑in signaling ...... 5

1.3.1 Mechanically gated ion channels ...... 5

1.3.2 Role of Adherens Junctions ...... 14

1.4 A mechanical view of morphogenesis...... 18

1.4.1 Cell and tissue behaviors that generate forces in morphogenesis...... 19

1.5 Dorsal closure is model system for studying forces in morphogenesis...... 21

1.5.1 Tissue movements in dorsal closure...... 21

1.5.2 Actin and myosin in closure...... 27

1.5.3 Forces in dorsal closure...... 29

1.5.4 Mechanotransduction in dorsal closure...... 34

1.6 Central hypothesis...... 36

2. Mechanically gated ion channels regulate cell shapes and movements during dorsal closure...... 38

2.1 Results ...... 42

2.1.1 Pharmacological inhibitors of mechanically gated ion channels disrupt closure...... 42

2.1.2 Dorsal closure requires extracellular and intracellular calcium...... 59

2.1.3 Observing endogenous calcium dynamics during dorsal closure...... 63

2.1.4 Uncaging calcium in dorsal epithelia ...... 68

2.2 Discussion ...... 69

3. Identification of candidate mechanically gated ion channel subunits for dorsal closure...... 75

3.1 Results ...... 78

3.1.1 RNAi screen for MGC candidates ...... 78

3.1.2 RNAi knockdown of RPK and dTRPA1 disrupts dorsal closure...... 80

3.1.3 GAL4 and RNAi expression patterns...... 89

3.1.4 The effect of mutations in MGC candidates on dorsal closure...... 91

3.1.5 The effect of rpk mutations on dorsal closure...... 94

3.1.6 The role of nompC (TRPN) in dorsal closure...... 99

3.1.7 An integrated approach to MGC knockdown in closure...... 105

3.2 Discussion ...... 112

4. Conclusions ...... 119

4.1 Specific conclusions and future directions...... 120

4.1.1 Identification of MGCs for dorsal closure: electrophysiological analysis...... 120

4.1.2 Identification of MGCs for dorsal closure: heterologous expression systems...... 121

4.1.3 Site directed mutation of MGC candidates...... 122

4.1.4 RNAi screen for MGC candidates: dmPeizo...... 125

4.1.5 Calcium signaling during Drosophila embryogenesis...... 127

4.2 Implications of this work...... 131

5. Methods ...... 133

5.1 Drosophila strains...... 133

5.2 Pharmacology and microinjection of Drosophila embryos...... 133

5.3 Molecular Biology ...... 134

5.4 Confocal microscopy and laser microsurgery...... 138

5.5 Immunoblotting and Immunofluorescence ...... 139

5.6 Reagents ...... 140

Appendix A ...... 141

A.1 rpk53 allele sequence analysis...... 141

References ...... 144

Biography ...... 166

vii

List of Tables

Table 1. GdCl3 inhibits dorsal closure in a dose‑dependent manner...... 44

Table 2. GsMTx4 blocks dorsal closure in a dose‑dependent manner...... 45

Table 3. GsMTx4 inhibition of channels leads to defects in seam formation during closure...... 54

Table 4. Microinjection of calcium chelators inhibits dorsal closure...... 60

Table 5. Calcium indicators used in this study...... 64

Table 6. RNAi screen for MGC candidates in dorsal closure...... 79

Table 7. Rpk‑ and trpA1RNAi lethality and GAL4 driver strength...... 83

Table 8. Embryonic lethality associated with mutations in dtrpA1 or rpk...... 92

Table 9. Embryonic lethality associated with nompC mutants...... 103

Table 10. HC‑030031 disrupts dorsal closure in a dose‑dependent manner...... 106

Table 11. Embryonic lethality associated with combinations of nompC, rpk and dtrpA1 alleles...... 110

viii

List of Figures

Figure 1. A model of MGC response to mechanical stimuli ...... 7

Figure 2. Basic polarity and organization of key adhesions in epithelial cell sheets...... 15

Figure 3. Dorsal closure during Drosophila embryogenesis...... 22

Figure 4. MGC inhibitors block dorsal closure in a dose‑dependent manner ...... 43

Figure 5. Role of apoptosis in epithelial integrity of GsMTx4 treated embryos ...... 47

Figure 6. MGC‑inhibited dorsal epithelia fail to regulate forces in response to laser microsurgery ...... 50

Figure 7. MGC inhibition interferes with F‑actin localization and dynamics ...... 52

Figure 8. Cadherin localization is disrupted in MGC inhibited embryos...... 56

Figure 9. MGC inhibition during germband retraction...... 58

Figure 10. Chelating extra‑ and intracellular calcium disrupts dorsal closure ...... 61

Figure 11. Effects of ionomycin on GCaMP3 reporter and cell contraction ...... 65

Figure 12. Endogenous calcium flux in amnioserosa cells ...... 67

Figure 13. Uncaging calcium triggers rapid cell contractions ...... 69

Figure 14. Model of mechanically gated ion channel activity in dorsal closure epithelial ...... 74

Figure 15. RNAi knockdown of RPK and dTRPA1 disrupts closure ...... 81

Figure 16. JNK signaling in RPK and dTRPA1 knockdown embryos ...... 84

Figure 17. MGC knockdown in the lateral epidermis during dorsal closure ...... 86

Figure 18. Laser wounding of MGC knockdown embryos ...... 88

Figure 19. Expression pattern of GAL4 drivers used in this study ...... 90

Figure 20. Effect of rpk and dtrpA1 mutants on dorsal closure ...... 93

Figure 21. RPK localization in dorsal closure tissues ...... 96

Figure 22. GFP‑RPK localization ...... 97

Figure 23. Characterization of nompC expression in dorsal closure ...... 100

Figure 24. Microinjection of the TRPA1 inhibitor, HC‑030031, during dorsal closure ... 108

Figure 25. Immunoblot analysis of RPK expression ...... 109

Figure 26. Embryos mutant for combinations of nompC, rpk, and dtrpA1 display dorsal closure defects ...... 112

Figure 27. Southern blot analysis of the rpk53 allele ...... 143

Acknowledgements

Thank you Dan, for your constant mentorship, advice, critique, support, and

encouragement. Your enthusiasm for exciting and meticulous science is inspiring.

Thanks to my labmates for your advice and collaborative spirit; for editing,

listening, and tea time; for your technical support.

Thank you also to Ruth, Eric, Sam, Gao, Anna, Selina, and members of my

committee for your generous advice and technical support.

1. Introduction

Where does the diversity of organismal structure come from? The life cycle of

nearly all organisms involves shape changes: the development of a vertebrate embryo,

or the transition from larvae to pupae in the life cycle of an insect are dramatic examples

of these processes. Over the last thirty years, genetic and molecular studies have thrown

open the doors on embryonic development, providing tools and a new understanding of

how shape change is controlled. Development is carefully driven by a largely conserved

and complex network of genes, morphogens, and signaling pathways that specify

cellular identity, growth, death, and behavior. We have extensive observations of cell

shape changes from various model systems of morphogenesis, and from cell culture.

Now the challenge is to better understand how biomechanical information is integrated

into the molecular and biochemical pathways that drive morphogenesis.

1.1 Why is mechanotransduction relevant?

Mechanotransduction is the process by which mechanical stimuli are translated

into biochemical signals. All cells are mechanosensitive, but the mechanisms by which

they sense force remain elusive (Orr et al., 2006). Recent research has revealed the

molecular identities of several key mechanosensitive proteins and protein complexes,

which provides a parts list that will be useful for future research on how such protein

complexes function in mechanotransducing pathways.

Force sensing plays a critical role in the function of sensory organs, in tissue

homeostasis, and in development (Jaalouk and Lammerding, 2009). For example, our

sense of hearing relies on the ability of inner ear cells to transduce sound waves into

neural signals via the mechanical activation of tip links (Corey, 2006). Mutations in the

proteins that maintain tension in these cells (including unconventional myosins and

cadherins) can result in deafness. Several non‑sensory organ tissues also rely on

mechanical stimuli to maintain homeostasis (e.g., arteries, nephrons, and muscles), and

the failure of mechanotransduction in these systems can lead to disease (atherosclerosis,

FSGS, and muscle dystrophies, respectively). A remarkable finding of the cellular

requirement for mechanotransduction is the effect of the mechanical environment on

tumorigenesis (Butcher et al., 2009; Schedin and Keely, 2011). Studies show that tumors

remodel their 3D environment as they transform, and that cellular tension as mediated

by small GTPases contributes significantly to invasive phenotypes. A central theme

regarding these diseases is that while we can clearly understand the effects of force and

the requirement of tension in these cells and tissues, the signaling pathways that

mediate mechanotransduction for these situations are not known.

The major question addressed by this introductory chapter is how embryonic

tissues sense and respond to mechanical forces generated as the embryo develops its

final form. We briefly introduce how cells generate force, and focus on known

mechanisms of cellular mechanotransduction. Then we examine research that has

identified the mechanical forces which contribute to developmental processes, and what

mechanosensors, if any, have been identified. Finally, we address the specific case of

dorsal closure during Drosophila development. We review how mechanical forces drive

this process and what we know about force sensing in closure.

1.2 Actomyosin contractility

Cellular contractility in non‑muscle cells is mediated by the ATP‑dependent

association of myosin II with the actin cytoskeleton (unless otherwise noted, ‘actin’

refers to filamentous F‑actin). When we discuss the forces for morphogenesis, the basic

provider of most, if not all of these forces is the motor activity of myosin II. Myosin II

produces force through conformational changes in the head domain due to ATP

hydrolysis that allows the processive binding and release of actin (Sellers, 1991; Vicente‑

Manzanares et al., 2009). Bipolar minifilaments of myosin II associated with actin has the

potential to be a contractile network; the properties of this network depend largely on

the organization of actin (i.e., bundles or branches) and the presence of actin crosslinkers

(Bendix et al., 2008).

Another important property for understanding the transmission of forces in

morphogenesis is mechanical stiffness (Wozniak and Chen, 2009). For a single cell, we

can think of mechanical stiffness in its immediate environment as the degree to which

the scaffolds that it is attached to – the extracellular matrix and other cells – is

deformable by the contractile forces that the single cell generates. In the case of a single 3

cell, local stiffness is mediated by cytoskeleton deformability and contractility, cellular

adhesion, and extracellular matrix composition. Stiffness can also be a property of an

entire tissue, distinct from adjacent tissues, as in the case of dorsal tissue during Xenopus

neurulation (Rolo et al., 2009; Zhou et al., 2009).

The emergent properties of actin organization and myosin II contractility are

spatially and temporally regulated in cells. For example, in migrating cells the

organization of actin cytoskeleton differs at the leading edge protrusions (primarily

branched networks of F‑actin) compared to the retracting posterior of the cell (primarily

anti ‑parallel bundled actin). Furthermore, little to no local myosin II function is needed

for leading edge protrusions, whereas myosin II activity at the rear of the cell aids its

retraction as the cell moves forward (Vicente‑Manzanares et al., 2009). In cells that

comprise tissue sheets, actin and myosin organization often reflects apico‑basal cell

polarity (Levayer and Lecuit, 2012). An example of this is in Drosophila epithelia cells

during gastrulation, when the apical region of the cell is dominated by apical‑medial

networks and junctional belts of actomyosin, and the baso‑lateral regions by cortical

actomyosin. Forces that are generated at the sub‑cellular scale (e.g., actomyosin

contraction), are organized in individual cells (apical constriction), and ultimately

translate into tissue movements across the embryo (gastrulation).

1.3 Force sensing: outside-in signaling

Here, we focus on how cells can sense changes in their mechanical environment

as a consequence of physical contact with neighboring cells or the extracellular matrix

(ECM). Such changes are due to cell motility, contraction, or tissue dynamics, such as

those during wound healing. Mechanical stimuli can be transmitted to the cell in any

number of ways, but the three means by which we understand the most about cellular

mechanotransduction are focal adhesions (i.e., forces transmitted through cell‑ECM

contacts), cell‑cell adhesions, and transmembrane receptors, including mechanically

gated ion channels (MGCs). Since the focus of this paper is mechanosensing in

morphogenesis with special attention to the role of MGCs, we discuss mechanosensing

by MGCs and then introduce the paradigms of junctional mechanosensing as they relate

to MGC activity.

1.3.1 Mechanically gated ion channels

MGCs are integral membrane proteins that transduce forces exerted through the

membrane into biochemical signals (Hamill, 2006). The consequence of applying force to

MGCs is change in gating, i.e., the probability that the channel is open or closed. When

open, these channels exhibit a range of permeability, typically to ions (e.g., Ca2+, K+, Na+,

and Cl‑). MGC‑based mechanosensing is thought to have evolved very early, given that

many unicellular dynamics (e.g., cell division) require changes in cell shape, but also

because MGCs have been identified in bacteria, archaea, and eucarya (Martinac and

Kloda, 2003).

There are two principle models that explain mechanical gating: a membrane

model in which the channel is sensitive to forces parallel to the plasma membrane, and a

tethered model, in which a channel domain(s) interacts with accessory proteins (such as

the cytoskeleton or extracellular matrix) and force applied to the tether leads to changes

in gating (Figure 1, also Christensen and Corey, 2007). Channels that are mechanically

sensitive are not necessarily mechanically gated, as there are instances of channels

involved in mechanosensing processes but appear to be downstream of a primary

signaling event (Becchetti et al., 2010). For example, the mechanosensory complex

responsible for mammalian hair cell mechanoreceptor currents involves the MGC

TRPA1 (Corey et al., 2004). However, TRPA1 knockout mice are not deaf indicating that

TRPA1 is not the primary mechanosensor in hair cells, but likely downstream of the

mechanosensor (Kwan et al., 2006). Channels are considered directly gated if the delay

between stimulus and ion flux is at least faster than known second‑messengers and if

knockdown of their expression leads to loss of a mechanosensory response (Christensen

and Corey, 2007).

1 ECM membrane 2 cytosol 3

Applied force Ions

Figure 1. A model of MGC response to mechanical stimuli. In this model, MGCs open, allowing ion flux, in response to forces generated in 1) the extracellular space, 2) the plasma membrane, and/or 3) the cytoplasm (e.g., the cytoskeleton).

1.3.1.1 A paradigm of MGC activity: MscL.

One of the best understood MGCs is the E. coli mechanosensitive channel of large

conductance, MscL. This is one of three classes of channels in E. coli that function in the

cellular response to osmotic shock (Haswell et al., 2011). MscL is a homopentamer with

non‑specific ion selectivity. Voltage‑clamping experiments show that MscL conductance

is ~3 nS, with fast kinetics (Berrier et al., 1996; Sukharev et al., 2001). The rate of ion flow

through the channel is measured in Siemens (S). Kinetics refers to the rate of change in

conductance that ion channels undergo when transitioning from closed to open

(sometimes through intermediates). The molecular interactions between the channel and

the membrane are critical for MscL force transduction. Briefly, the opening of MscL is

primarily due to force‑dependent conformational changes in transmembrane helix

packing, and stabilized by the hydrophobic mismatch of helix resides and the lipid

bilayer (Sukharev et al., 2001; Perozo et al., 2002b). Another important modifier of MscL

activity is the geometry of the plasma membrane, as the open‑state of the channel is

altered by the asymmetric incorporation of cone‑shaped lipids (e.g.,

lysophosphatidylcholine) in liposomes (Perozo et al., 2002a).

1.3.1.2 Methods for studying MGCs.

A number of techniques have been critical in our understanding of MscL activity,

and are proving to be important in the discovery and study of MGCs in eukaryotes

(Haswell et al., 2011). When biochemically homogenous MscL is reconstituted into

purified liposomes, it retains mechanosensitivity under patch clamp conditions (Hase et

al., 1995) . The patch clamp method allows the detection of single channel currents in

plasma membranes (Neher et al., 1978; Hamill et al., 1981). Channel activity in purified

liposomes indicates that mechanotransduction by these channels is direct, without any

requirement for interactions with other proteins, the cytoskeleton, or the extracellular

matrix. Since its introduction, patch clamp recordings of MGCs expressed in liposomes,

heterologous systems (especially Xenopus oocytes), or in whole cells represent the ‘gold

standard’ for determining if a channel is mechanosensitive.

The mechanism of MscL mechanosensing was advanced by the discovery of its

crystal structure (Chang et al., 1998; Steinbacher et al., 2007). This work demonstrated

that MscL is a homopentamer of subunits. Each subunit has two transmembrane

domains, one that faces the plasma membrane (TM1) and one that faces the pore (TM2).

Closed, the pore is 2 – 18 Å in diameter, but is occluded on the cytoplasmic side. The

study on the effects of lysophosphatidylcholine on MscL gating further applied this

strategy to stabilize and analyze the crystal structure of MscL in its open state (Perozo et

al., 2002a). Open, the MscL pore is > 25 Å in diameter. Additional work has been done to

determine the structure of the channel in its intermediate conformations (Sukharev et al.,

2001). These structures are used to model the transition between closed and open MscL,

and established the dependence of this transition on interactions between TM1 the lipid

bilayer.

Finally, mutagenesis (both site‑directed and random) has identified specific

residues in MscL helices that enhance or inhibit mechanosensitivity of the channel (Ou

et al., 1998; Sukharev et al., 2001; Perozo et al., 2002a; Yoshimura et al., 2004). A number

of amino acid residues in both TM1 and 2 are now known to contribute to the gating

mechanism of MscL. Together, these methods elucidate the structure and molecular

mechanisms of MscL activity in response to force, and represent standard methods that

are used to understand the general activity of MGCs.

1.3.1.3 TRP channels

Members of the transient receptor potential (TRP) family of MGCs in eukaryotes

have been widely implicated in mechanosensing in various organs, tissues and

organisms (Christensen and Corey, 2007). Although they are expressed in sensory

neurons, TRP family channels do not preferentially express in sensory neurons. TRP

family channels are widely expressed in many tissues, often at low protein levels (Desai

and Clapham, 2005). Sequence and topology divide the TRP superfamily into two

groups, each characterized by 7 subfamilies, and one distant subfamily consisting of the

yeast TRPY. In this study, we will focus on Group 1, which represent channels that are

more closely related to the founding TRP channel in Drosophila (Montell and Rubin,

1989). All group 1 TRP channels are tetrameric, cationic selective channels, which share

sequence homology in the six transmembrane domains and the pore loop between the

fifth and sixth transmembrane domains. These channels largely lack other conserved

sequences, domains or function (Venkatachalam and Montell, 2007). Despite this

variation, all TRP channels appear to be involved in sensing external stimuli.

TRP channels are activated by stimuli including light, temperature, chemicals,

and touch; some channels are activated by multiple mechanisms. At least one member of

each subfamily and TRPY has been shown to mediate mechanical stimuli in cells

(Castiglioni and Garcia‑Anoveros, 2007). Many of these channels have been associated

with defects in mechanical nociception, bristle touch, and hearing in the Drosophila

model system (Walker et al., 2000; Tracey et al., 2003; Gong et al., 2004). In bristle touch,

for example, transduction currents in neurons associated with adult bristle movement

were nearly abolished in flies lacking TRPN1 (encoded by the gene nompC) expression

(Walker et al., 2000). TRPN1 is also required for touch in larvae and hearing in the adult

fly (Kamikouchi et al., 2009; Cheng et al., 2010a). A major problem with of all of the

Drosophila mechanosensitive TRPs studied to date is that none of them have been shown

to be mechanosensitive under patch clamp conditions when expressed in heterologous

systems. There are several reasons that could explain this – the heteromeric constitution

of channel complexes, lack of other interactors – but because of this, further examination

is necessary to determine whether these channels function as direct, primary

mechanosensors.

In non‑sensory cells and tissues, mechanotransduction and TRP channels

contribute to contractility and directional movement. TRPC, TRPV and TRPM channels

have been associated with cell motility and morphogenesis (Thodeti et al., 2009; Wei et

al. , 2009; Tian et al., 2010; Liu et al., 2011). In these studies, knockdown of channel

expression by siRNA or morpholino eliminates the mechanoresponse. In the examples

from cell culture studies, the mechanoresponse to includes Rho‑GTPase dependent

reorganization of the actin cytoskeleton, calcium‑dependent cytoskeletal reorientation

during the application of cyclical stretch, and directional migration during chemotaxis.

In the last example, knockdown of the stretch sensitive TRPM7 in fibroblasts led to loss

of Ca2+ ‘flickers’ (transient, microdomains of increased Ca2+) and the inability of

migrating fibroblasts to change direction during chemotaxis (Wei et al., 2009). On the

other hand, there are far fewer examples of roles for TRP channels in embryonic

morphogenesis. In the study mentioned (Liu et al., 2011), morpholino suppression of

TRPM7 expression blocked cell movements associated with convergent extension during

gastrulation. A persistent observation in these studies that perturb TRP channel function

in non‑sensory cells is the tight coupling of channel activity with regulation of the

cytoskeleton.

No high‑resolution structure for a full‑length TRP channel exists (Li et al., 2011).

Numerous studies have introduced point mutations and domain deletions to TRP

channels to study channel activity in vivo (Christensen and Corey, 2007). Some of these

will be discussed in Chapter 4. Many of these mutations are proposed to affect pore

selectivity, conductance, and channel kinetics, to name a few, and contribute to the

characterization of TRP channel function in mechanosensory processes. However, to

date the only TRP channel that has been thoroughly analyzed under patch cla mp

conditions is yeast TRPY, encoded by TrpY1 (Zhou et al., 2003; Arnadottir and Chalfie,

2010). TRPY is activated in vacuolar membranes in response to osmotic shock. In

particular, a forward genetic screen for TrpY1 gain‑of‑function mutants in combination

with patch clamping contributes to our understanding of the mechanism of this TRP

channel’s activity (Su et al., 2007).

1.3.1.4 DEG/ENaCs

The other class of MGC considered here are the Degenerin/Epithelial sodium

channels (DEG/ENaC). DEG/ENaC channels are expressed in a wide variety of tissues,

including epithelia, neurons, and muscle (Mano and Driscoll, 1999). Two major

characteristics of these channels are that they are permeable mainly to sodium (Na+) ions

and can be blocked by amiloride. DEG/ENaC proteins exhibit more sequence similarity

to one another than TRPs, with two transmembrane domains, intracellular amino and

carboxyl termini, and a large extracellular loop (Kellenberger and Schild, 2002). Several

C elegans DEG/ENaCs are sensitive to a mutation in the extracellular loop near the N‑

terminal transmembrane domain, termed the DEG mutation, which leads to constitutive

channel activity (Huang and Chalfie, 1994). This finding supports the role of the loop in

channel gating, which is largely mediated by residues in the N‑terminal transmembrane

domain. Channel subunits can form homo‑ or hetero‑multimers, like TRPs, but subunit

stoichiometry varies among channel complexes.

Some of the first DEG/ENaC proteins identified resulted from a screen for touch

insensitivity in C elegans (Chalfie and Sulston, 1981; Chalfie and Au, 1989). MEC‑4 and

MEC‑10 are DEG/ENaC proteins that form the pore of the MGC complex in C elegans

touch receptor neurons, as demonstrated through knockdown and directed mutation

analyses (Goodman et al., 2002; OʹHagan et al., 2005). Interestingly, when MEC‑4 is

expressed alone in the Xenopus oocyte heterologous expression system it is not

mechanically activated, but demonstrates a mechanoresponse when co‑expressed with

MEC‑10 (Goodman et al., 2002). In vivo patch clamping of C elegans touch receptor

neurons showed that MEC‑4 and MEC‑10 are required for currents induced by touch.

The mechanism by which this MGC complex transmits force is the focus of intense

research, and together these findings are an example of how we can apply the methods

that were established in MscL research to the study of MGCs in general.

A mechanosensory role has been hypothesized for DEG/ENaCs in other systems,

including Drosophila and vertebrates. Mammalian ENaCs are thought to mediate

endothelial response to shear stress in the kidney and blood vessels (Carattino et al.,

2004; Wang et al., 2009). In Drosophila, among a number of genes predicted to be

homologues, the two best characterized DEG/ENaC proteins are Pickpocket and Ripped

pocket (Adams et al., 1998). Pickpocket (PPK) expression in specific sensory neurons is

required for larval response to noxious mechanical stimuli (Zhong et al., 2010), and

Ripped pocket (RPK) may be involved in oocyte activation and are sensitive to the DEG

mutation (Adams et al., 1998; Horner and Wolfner, 2008a). RPK can form amiloride and

GdCl3‑sensitive homomultimers when heterologously expressed in Xenopus oocytes

(Adams et al., 1998), supporting the hypothesis that it functions as an MGC.

These examples of TRP channels, DEG/ENaCs, and prokaryotic MSCs

demonstrate some of the diversity of mechanically gated ion channels that exist in cells.

1.3.2 Role of Adherens Junctions

Cells in the Drosophila embryo express four types of junctions: gap junctions,

septate junctions, cell‑cell and cell‑ECM junctions (Tepass and Hartenstein, 1994). It is

not known if gap and septate junction proteins are mechanosensitive or if their function

is regulated by mechanosensory pathways. The role of cell‑cell and cell‑ECM junctions

in mechanosensing, while difficult to determine in developing embryos, is well

established in cell culture. 14

Figure 2. Basic polarity and organization of key adhesions in epithelial cell sheets.

1.3.2.1 Junctional mechanosensing: focal adhesions.

Cells typically maintain traction on their substrate via integrin‑based focal

adhesion complexes (FAs). The purpose of FAs is to anchor the cell to the ECM (Figure

2). They are the site of cytoskeleton regulatory signaling and the transmission of forces

to and from the ECM (Zamir and Geiger, 2001). Both the maturation and function of FAs

are sensitive to force: as focal complexes become less associated with rapidly turning

over actin cytoskeleton characteristic of the leading edge of cellular protrusions and

more associated with actin stress fibers and myosin II, they mature into FAs (Gardel et

al., 2010). The proteome of FAs varies, but many associated proteins have been shown to

respond to force, including p130Cas and talin (Sawada et al., 2006; del Rio et al., 2009).

Integrin adhesions have been shown to be associated with and regulated by ion

channels (Arcangeli and Becchetti, 2006). A relationship between integrin

mechanosensing and MGCs was observed in magnetic twisting cytometry experiments,

in which magnetic beads coated with protein are bound to cells in culture and pulled on

through application of magnetic pulses (Sen and Kumar, 2010). One magnetic twisting

cytometry study of integrin mechanosensing in endothelial cells applied beads coated

with arginine‑glycine‑aspartic acid (RGD) peptide, which binds specifically to integrins.

The cellular response to bead pulling included an increase in cytoplasmic Ca2+ levels,

actomyosin ‑dependent stiffening of the cytoskeleton, and sensitivity of this cytoskeleton

stiffening to the MGC inhibitor GdCl3 (Matthews et al., 2006). Studies have also shown

that FA activity in the leading edge of migrating fibroblasts is GdCl3‑sensitive and

requires Ca2+ (Munevar et al., 2004).

1.3.2.2 Junctional mechanosensing: cell‑cell adhesions.

Compared to how we understand mechanotransduction at FAs, much less is

understood about how forces are transmitted at cell‑cell adhesions. Cell‑cell adhesions

based on the transmembrane, calcium‑dependent adhesion protein cadherin have been

characterized in terms of their function in adhesion homeostasis (Nelson, 2008). The

current model is that the intracellular domain of cadherin interacts with at least β‑

catenin and α‑catenin (Figure 2). This complex interacts with the actomyosin

cytoskeleton, directly or indirectly, and experiences a resting tension along cell‑cell

contacts. Previous models of the cadherin‑catenin complex describe direct and stable

interactions between cadherin, β‑catenin, α‑catenin, and actin cytoskeleton (Kobielak

and Fuchs, 2004). The cadherin‑catenin complex can be biochemically isolated, and

mutational analysis demonstrates the interdependence of the complex and the actin

cytoskeleton (Ozawa and Kemler, 1992; Quinlan and Hyatt, 1999). However, an

alternative model was proposed that questioned the stability of this linkage (Drees et al.,

2005; Yamada et al., 2005). In this model, the relationship between the cadherin‑catenin

complex and the actin cytoskeleton is mediated by transient interactions with α‑catenin.

This model proposes that m onomeric α‑catenin binds to the cytoplasmic domain of

cadherin, whereas α‑catenin homodimers bind to actin and antagonize the activity of

Arp2/3 (an actin regulator). Recently, α‑catenin was shown to be a mechanosensor that

specifically binds to vinculin upon stretch and exposure of cryptic binding sites

(Yonemura, 2011).

Mechanosensing by cadherin molecules was demonstrated by culturing cells on

deformable substrates coated with the extracellular domain of cadherin (Ladoux et al.,

2010). This research demonstrated a positive correlation between substrate rigidity and

traction forces exerted by the cells, supporting a mechanosensory role for cadherin

adhesions. Magnetic twisting cytometry experiments, this time with cadherin‑coated

beads, demonstrated a relationship between cadherin mechanosensing and channel

activity (Ko et al., 2001). Specifically, forces sensed at cadherin‑based adhesions activate

GdCl3‑dependent Ca2+ signaling. This subsequently leads to Ca2+‑dependent

reorganization of the actomyosin cytoskeleton and Ca2+‑dependent reinforcement of the

adhesion, through recruitment of additional cadherin and catenin.

Together, research addressing mechanotransduction at cellular junctions (both

cell‑cell and cell‑ECM) supports crosstalk with MGC activity.

1.4 A mechanical view of morphogenesis.

Mechanical questions drove many early studies in the field of developmental

biology, which has benefited greatly by the genetic and molecular dissection of

developmental patterning that has dominated the field in the last 30 ‑ 40 years. These

tools, in combination with new techniques to study the mechanics of cells, tissues, and

embryos, has driven our current knowledge of the role that mechanical forces play in

embryogenesis. Mechanotransduction is hypothesized to play a role in morphogenesis

and cell shape changes across several model systems (Zhou et al., 2007; Hamant et al.,

2008; Haswell et al., 2008; Sherrard et al., 2010). Progress is being made to identify and

demonstrate molecular mechanisms of force sensing in developmental processes. Not

surprisingly, the protein complexes we have described here (i.e., integrin adhesions,

cadherin adhesions, MGCs) have been shown to be key players.

1.4.1 Cell and tissue behaviors that generate forces in morphogenesis.

Studies of morphogenesis indicate that a wide range of cell shape changes and

tissue behaviors underlie organismal shape change and growth. These include apical

constriction, cell elongation, single or collective cell migration, and intercalation (Keller

et al., 2003; Wu et al., 2007; Gorfinkiel et al., 2010; Martin, 2010). Furthermore, cell shape

changes associated with cell growth, division and apoptosis can also influence

morphogenesis (Lecuit and Le Goff, 2007; Toyama et al., 2008).

1.4.1.1 The example of convergent extension.

Convergent extension (CE) is a process by which tissues narrow and lengthen

usually through the intercalation of individual and groups of cells. A series of

experiments have focused on CE movements associated with vertebrate neurulation and

gastrulation, Drosophila germband extension, and zebrafish epiboly (Adams et al., 1990;

Irvine and Wieschaus, 1994; Beloussov and Luchinskaia, 1995; Moore et al., 1995;

Wallingford et al., 2002; Keller et al., 2003; Benko and Brodland, 2007; Zhou et al., 2009).

During CE, cells intercalate mediolaterally to form the notochord and drive elongation

of the embryo. These studies use ex vivo measurements of tissue deformation to

determine the force generated in dorsal ectoderm during neurulation. These

experiments work because explanted tissues are able to largely recapitulate in vivo tissue

movements. Additionally, it was shown that these explants could react to and resist

applied stretching forces (Beloussov and Luchinskaia, 1995). Actomyosin based cell

contractility is required for these explant behaviors (Rolo et al., 2009).

We are beginning to understand additional cell signaling pathways that regulate

this contractility (Zhou et al., 2009; Weber et al., 2011). We know that integrins,

fibronectin, and cadherins are all required for CE cell movements (Dzamba et al., 2009;

Rozario et al., 2009; Theveneau et al., 2010). From these studies it is clear that there is

crosstalk between adhesion pathways, but the extent to which they are functioning as

mechanosensors is not known.

1.4.1.2 The examples of Drosophila anterior midgut differentiation and GBE.

Germband extension (GBE) is the process by which the Drosophila embryo

elongates on the anteroposterior axis. The formation of the foregut occurs concurrently

with GBE. Studies have demonstrated that force provided by GBE movements is

necessary and sufficient to induce expression of the mesoderm specific gene twist and

redistribution of myosin II activity during foregut differentiation and formation (Farge,

2003; Desprat et al., 2008; Pouille et al., 2009). In these experiments, tension was released

by laser wounding and reintroduced by application of force with magnetic tweezers on

intracellularly microinjected ferromagnetic fluid. They determined that an applied force

of 60 ± 20 nN is sufficient to rescue twist expression in embryos in which the connection

between the developing foregut and the extending germband was ablated. The

mechanosensitive regulator of twist activation is not known.

Further work employing cell tracking and strain rate modeling, in combination

with traditional fly genetics, focused on endogenous cell shape changes that drive GBE

itself (Blanchard et al., 2009; Butler et al., 2009). These studies used image segmentation

of wild type and mutant embryos to analyze the individual contribution of cell

intercalation and cell elongation to tissue deformation. They found that that cell

elongation drives early GBE and cell intercalation drives late GBE, and that this behavior

is further influenced by mechanical forces in the mesoderm.

Together, these types of experiments indicate that force production by a range of

cell behaviors is coordinated between tissues during embryogenesis.

1.5 Dorsal closure is model system for studying forces in morphogenesis.

1.5.1 Tissue movements in dorsal closure.

Dorsal closure is a model system of epithelial cell sheet morphogenesis, and is an

important system for the study of wound healing. Dorsal closure is an event that starts

in the middle of Drosophila embryogenesis (~stage 11) and is completed in 2 – 3 hours (at

stage 16; Figure 3A). By the end of germband retraction, the event immediately

preceding closure, the dorsal surface of the embryo is covered in an extraembryonic

tissue called the amnioserosa (Figure 3A, B). During dorsal closure the amnioserosa is

replaced with a continuous dorsal epithelium that can secrete cuticle, the tough

exoskeleton that protects hatched larvae from the environment. The bulk movements of

closure are the dorsal‑ward movement of the lateral epidermis, the reduction in area of

the amnioserosa tissue, and seam formation between the two fronts of the lateral

epidermis at the anterior and posterior corners of the dorsal opening, termed the canthi.

In wild type embryos, the movement of the leading edge towards the embryonic midline

occurs at a rate of 6.0 nm/s on average (labeled with sGMCA, at 22 °C; Hutson et al.,

2003; Toyama et al., 2008).

Figure 3. Dorsal closure during Drosophila embryogenesis. (A) Movements of germband retraction (left panel, arrow), dorsal closure (center), and end of closure (right). (B) Relevant dorsal closure tissues and structures, visualized by sGMCA (F‑ actin). (C) Distribution of amnioserosa cells, central (pink) and peripheral (green). (D) Time‑lapse of an sGMCA embryo, time in minutes. Scale bar, 25 μm; D, dorsal; V, ventral; A, anterior; P, posterior.

1.5.1.1 Lateral epidermis.

The dorsal movement of the lateral epidermis is associated with the dorsoventral

elongation of the first row of cells, termed the leading edge. During early closure, the

leading edge develops from a scalloped border to a straight border. This transition is

concomitant with the accumulation of F‑actin and non‑muscle myosin II (zip/MyoII) at

the leading edge (Kiehart et al., 2000; Solon et al., 2009). The development of a straight

leading edge is thought to be due, in part, to increased tension along its curvature. When

the leading edge is scalloped, individual lateral epidermal cells are polygonal; they

gradually elongate dorsoventrally as actin accumulates at the leading edge. The

accumulation of actin at the leading edge is organized in a supracellular purse string.

While F‑actin appears to extend from one cell boundary to the next, zip/MyoII co‑

localizes with actin in the space between cell boundaries at the leading edge giving a

‘bars on a string’ appearance (Franke et al., 2005). Several proteins localize to the

tricellular junctions (where the purse string meets cell‑cell boundaries) at the leading

edge, and most are associated with cellular junctions (e.g., α‑actinin and zyxin;

Rodriguez‑Diaz et al., 2008). The expression of these proteins and of others that regulate

leading edge cell behaviors is dependent on signaling cascades (e.g., Jun N‑terminal

kinase signaling, TGFβ signaling). In the absence of Dpp, a Drosophila TGFβ‑related

secreted factor, leading edge cells do not elongate and generate a defective purse string

(Fernandez et al., 2007).

Cells with a leading edge‑like fate appear to be specified by physical interaction

with the amnioserosa, as ‘islands’ of amnioserosa tissue formed in closure mutant

embryos are surrounded by Jun kinase expressing cells (Stronach and Perrimon, 2001).

Recently, the homophilic adhesion molecule Echinoid (Ed) was shown to be essential for

purse string formation (Chang et al., 2011; Laplante and Nilson, 2011). Ed is initially

expressed in all dorsal epithelia, but is lost in the amnioserosa as closure initiates. This

loss of Ed expression coincides with purse string formation. These findings underline

the importance of the physical interaction between the lateral epidermis and the

amnioserosa for dorsal closure.

1.5.1.2 Amnioserosa.

The amnioserosa is the epithelial tissue that covers the dorsal surface of the

embryo at the end of germband retraction, but its developmental patterning is a result of

signaling that occurs as early as axis specification during oogenesis (OʹConnor et al.,

2006; Pope et al., 2008; Ray et al., 1991). The reduction of total amnioserosa area during

dorsal closure is associated with spatially regulated apical constriction of individual

cells and apoptosis (Kiehart et al., 2000; Toyama et al., 2008; Gorfinkiel et al., 2009).

Peripheral amnioserosa cells adjacent to the leading edge constrict first and central cells

constrict last (Figure 3C). Amnioserosa cells near either canthi also contract earlier or

faster than central cells. As dorsal closure progresses, peripheral amnioserosa cells not

only apically contract but also extend their cell‑cell boundary with adjacent leading edge

cells, in a manner that is Dpp and integrin dependent (Narasimha and Brown, 2004;

Fernandez et al., 2007; Sokolow et al., 2012).

Amnioserosa cells primarily constrict orthogonal to the embryonic

anteroposterior axis over the course of dorsal closure (Blanchard et al., 2010). This

constriction is associated with dynamic oscillations in apical cell area with a period of ~

200 ‑ 300 seconds (Ma et al., 2009; Solon et al., 2009; Sokolow et al., 2012). These

oscillations are driven by the contraction of both dynamic apical actomyosin networks

and junctional belts of F‑actin (Kiehart et al., 2000; Fernandez et al., 2007; Gorfinkiel et

al., 2009; Ma et al., 2009; Solon et al., 2 009; Blanchard et al., 2010; David et al., 2010;

Sokolow et al., 2012). The dynamic localization of actin and zip/myoII to the apical

surface of amnioserosa cells correlates with constriction (Blanchard et al., 2010). These

patches of actomyosin travel across the cell at a rate of 0.2 μm/s (Ma et al., 2009). The

signaling pathways that regulate the apical networks include activity of the PAR

polarity complex (David et al., 2010). Mechanical oscillations are a well documented

phenomenon in morphogenetic events (for review see (Kruse and Riveline, 2011) and

(Martin, 2010), but the role of these oscillations in amnioserosa cell behavior is not clear.

For example, a recent study hypothesized that amnioserosa cell oscillations allow

passive reduction in amnioserosa area (Solon et al., 2009); another proposed that

amnioserosa cell oscillations are required for the coordination of cell contraction across

the tissue (Blanchard et al., 2010).

As dorsal closure progresses, apoptosis also contributes to the decrease in

amnioserosa tissue area. Initially consisting of ~200 cells, ~10% of these cells delaminate

from the amnioserosa epithelium and undergo apoptosis (Kiehart et al., 2000; Hutson et

al., 2003; Toyama et al., 2008; Sokolow et al., 2012). By the end of dorsal closure, the

ultimate fate of the amnioserosa cells appears to be programmed cell death and

phagocytosis by hemocytes.

1.5.1.3 Seam formation.

Finally, at both canthi of the dorsal opening seam formation occurs. The specific

interaction of actin protrusions from the leading edge and contraction of the underlying

amnioserosa cells together constitute zipping, which is the process of seam formation

(Jacinto et al., 2000; Millard and Martin, 2008). One purpose of zipping is to physically

bring the two lateral epidermal fronts together. Before dorsal closure occurs,

developmental patterning of Drosophila embryos in the anteroposterior axis establishes

unique parasegments that ultimately contribute to the patterning of the larvae and adult

fly (Lawrence and Struhl, 1996). By the end of dorsal closure, these segments are

continuous across the dorsal surface. Another purpose of zipping, then, is to regulate the

process of segment matching. Along the entire length of the leading edge we observe the

generation of actin protrusions, including filopodia and lamellopodia. Similar to

observations in cell culture, filopodia and lamellopodia are regulated by small GTPases

(especially Rho, Rac and Cdc42) and actin regulators (e.g., enabled; Harden, 2002; Jaffe

and Hall, 2005; Gates et al., 2007). The mechanism of segment alignment during closure

is not clear, but it appears to require the presence of actin protrusions, zip/myoII

contractility, and cell‑cell adhesion regulators (Bloor and Kiehart, 2002; Franke et al.,

2005; Gates et al., 2007; Gorfinkiel and Arias, 2007).

The coordination of these three tissue movements ultimately generates a

seamless dorsal epidermis.

1.5.2 Actin and myosin in closure.

1.5.2.1 Non‑muscle myosin II produces force in dorsal closure.

The heavy chain of non‑muscle myosin II is encoded by the Drosophila gene

zipper (Young et al., 1993). Together with the actin cytoskeleton, zip/myoII generates the

forces required for dorsal closure and is a key component of the force producing

structures described above (Young et al., 1993; Kiehart et al., 2000; Franke et al., 2005). In

the absence of zygotic zip/myoII (myosin protein is maternally loaded) the purse string

is disorganized, dorsal closure slows, and adhesion within the amnioserosa or between

the amnioserosa and lateral epidermis fails (Franke et al., 2005). zip/myoII localizes to

bars along the purse string that coincide with the leading edge but are excluded from

cell‑cell junctions (Young et al., 1993; Franke et al., 2005). In the amnioserosa, zip/myoII

localizes to cell boundaries and apical actin networks (Franke et al., 2005; David et al.,

2010). Mosaic rescue of zip/myoII in dorsal closure tissues lacking functional

endogenous zipper expression demonstrated that zip/myoII is required for the 27

generation of tension in the purse string and in amnioserosa cells, as non‑rescued cells

were stretched and failed to contract as closure progressed (Franke et al., 2005).

1.5.2.2 Other myosins in dorsal closure.

Other myosins are expressed in Drosophila embryos, and at least two have been

documented to have a dorsal closure phenotype. Myosin XV is encoded by the gene

sisyphus (Liu et al., 2008). Knockdown of sisyphus expression affects filopodia activity

and affects segment matching during dorsal closure. There is conflicting evidence that

myosin VI, encoded by the gene jaguar, plays a role in dorsal closure (Millo et al., 2004;

Morrison and Miller, 2008) . Alleles of jaguar with disrupted promoter regions and loss of

myosin VI expression have dorsal closure defects, including defects in amnioserosa and

lateral epidermal cell shapes, purse string formation and epithelial integrity (Millo et al.,

2004). However, subsequent genetic characterization proposed that defects associated

with these and other jaguar alleles is due to disruption of a neighboring gene, and that

loss of myosin VI function is not lethal in flies, consistent with results in other species

(Morrison and Miller, 2008).

1.5.2.3. Actin regulation

F‑actin, as visualized by the GFP‑tagged actin binding domain of moesin

(sGMCA; Edwards et al., 1997; Kiehart et al., 2000) or by GFP‑actin localizes to the

periphery of all cells in the amnioserosa and the lateral epidermis in a manner consistent

with the characteristic localization of actin to cell‑cell junctions and cortical membranes

in other epithelia (Papusheva and Heisenberg, 2010). There is a strong localization of

sGMCA to the leading edge of the lateral epidermis, consistent with the location of the

purse string. As mentioned, filopodia and lamellopodia are produced at the leading

edge of the lateral epidermis, but filopodia‑like structures are also observed at the apical

surface of amnioserosa cells.

Non‑muscle cytoplasmic actin in Drosophila is encoded by two genes, which are

expressed at different points in development from embryo to adult fly (Tobin et al.,

1990). The two actin genes, 5C and 42A, are expressed in cell lines and during

embryogenesis (Cherbas et al., 2011; McQuilton et al., 2012). Actin regulators expressed

during dorsal closure include Arp2/3 and profilin, and the expression of some of these

genes is promoted by JNK signaling in the leading edge (Jasper et al., 2001; Harden,

2002). The role of the cytoskeleton regulating small GTPases (RhoA, Rac, and Cdc42) in

dorsal closure has been studied extensively (Harden et al., 1999; Bloor and Kiehart, 2002;

Woolner et al., 2005).

1.5.3 Forces in dorsal closure.

The cell sheet movements and cell shape changes described above ultimately

require the generation of forces in order to achieve them. The evidence for forces in

dorsal closure comes from a series of laser ablation experiments (Kiehart et al., 2000;

Hutson et al., 2003; Peralta et al., 2007; Rodriguez‑Diaz et al., 2008; Toyama et al., 2008;

Ma et al., 2009; Solon et al., 2009) in which recoil and other behaviors after induced 29

lesions served as a readout for the distribution of tension in cells or the epithelium. From

these studies, a force balance equation has emerged that describes the contributions of

individual forces towards dorsal closure, at the symmetry point on the leading edge: σLE

– σAS – Tκ = b dh/dt. dh/dt is the rate of dorsal closure, where h is the distance of the

leading edge to the embryonic midline. b represents viscous drag, which is a

consequence of both intracellular properties and connective (adherent) properties of the

tissues that participate in closure. The rest of these terms will be described below.

1.5.3.1 Force production in the purse string.

In order to determine if the purse string in the leading edge is under tension,

small wounds were introduced at the leading edge and the subsequent behavior of cells

was observed (Kiehart et al., 2000; Rodriguez‑Diaz et al., 2008). These results showed

that the purse string behaves like a spring under tension. After a laser wound was made,

purse string on either side of the wound rapidly recoiled away from the site of

wounding. Over time, recoil slows, then the tissue recovers and dorsal closure

completes. Further analysis indicates that the purse string may not produce uniform

tension along its length (Peralta et al., 2008). There is evidence suggesting that leading

edge cells oscillate to a degree, but the significance of this, if any, is not known. Instead,

it appears that concerted contraction (i.e., irreversible shortening) is localized to zipping

events at the canthi. What drives the transition towards contraction is not understood.

Taken together with the laser ablation studies, this suggests that the bulk of the purse

string is in a dynamic state of tension without net contraction.

Quantitative modeling of the forces that drive dorsal closure represent the force

in the purse string that contributes to the movement of the leading edge towards the

embryonic midline as Tκ (Hutson et al., 2003; Peralta et al., 2008). T is the tension in the

purse string, and κ is the curvature of the leading edge. Compared to other forces

generated in closure, the tension in the purse string is the largest, approximately 2‑5

times larger than the force per unit length of the amnioserosa (assuming the average cell

is 10 μm in diameter). However, this tension is generated along the curvature of the

leading edge, and thus contributes less to movements towards the embryonic midline

approximately equivalent to forces from the amnioserosa.

1.5.3.2 Force production in the amnioserosa.

Forces in the amnioserosa were investigated by a series of laser ablation studies:

small (1‑2 cells) laser ablations (Kiehart et al., 2000); ‘A1’ cuts, laser incision from one

canthi to the other across the amnisoerosa (Hutson et al., 2003); and edge cuts, laser

incision from one canthi to the other along the amnisoerosa side of the leading edge

(Toyama et al., 2008).

Small laser ablations in the amnioserosa, like those made in the purse string,

resulted in rapid recoil away from the site of ablation indicating that the amnioserosa

tissue is under tension (Kiehart et al., 2000). In this work, it was observed that repeated

ablations in the amnioserosa appeared to increase the rate of closure compared to

internal controls (i.e., only one half of the amnioserosa had been targeted for laser

ablation and it closed faster than the other half). A1 and edge cuts were designed to

determine the contribution of amnioserosa contraction towards dorsal closure (Hutson

et al., 2003; Toyama et al., 2008). The edge cuts were developed to avoid the effects of the

remaining, attached amnioserosa on the leading edge observed during A1 cuts. Using

snaking algorithms to follow the movement of the purse strings before and after A1 or

edge cuts, the inital recoil of the purse string away from the cut amnioserosa is

determined. The initial recoil velocity is proportional to σAS, the force per unit length

produced by the amnioserosa on the leading edge (Hutson et al., 2003; Toyama et al.,

2008). Together these experiments determined that the amnioserosa contributes a

contractile force that is comparable in magnitude to Tκ.

The generation of forces that drive closure have recently been examined in

greater detail at the cellular level. All amnioserosa cells apically constrict, but in ~10% of

these cells apical constriction is associated with ingression followed by apoptosis

(Kiehart et al., 2000; Toyama et al., 2008). In the absence of apoptosis the rate of closure

decreases, and in the presence of increased apoptosis the rate of closure increases. This

contribution appears to be about one‑third of the total force produced in the

amnioserosa (Toyama et al., 2008). Recently, it was shown that cells that ingress and

undergo apoptosis can be identified prior to changes in cell shape or F‑actin

organization by observation of mitochondrial fragmentation (Muliyil et al., 2011).

In the remaining amnioserosa cells, there appear to be two kinds of apical

constrictions. The first is associated with dynamic oscillations in apical cell area and are

primarily observed prior to and at the onset of closure, described above (Gorfinkiel et

al., 2009; Ma et al., 2009; Solon et al., 2009; Blanchard et al., 2010; David et al., 2010). The

second is a more sustained apical constriction observed after the onset of closure

(Sokolow et al., 2012).

The dynamic apical cell oscillations observed in dorsal closure correspond to the

formation and dissipation of contractile apical actomyosin networks (Ma et al., 2009).

Cell boundaries recoil in response to repeated laser ablation of the apical surface

demonstrating that these structures generate force. The nature of the physical

relationship and how forces are coordinated between junctional and apical actomyosin

arrays is unclear.

1.5.3.3 Force production in the lateral epidermis.

The bulk of the lateral epidermis is under tension during closure (Kiehart et al.,

2000; Hutson et al., 2003; Rodriguez‑Diaz et al., 2008). However, the lateral epidermis

contributes a resisting force (i.e., favoring movement away from the embryonic midline)

to dorsal closure (σLE) . Currently it is not known if this tissue is being passively pulled

by forces in the amnioserosa and the purse string, or if it actively senses and (nearly)

balances these forces in opposition of closure.

1.5.4 Mechanotransduction in dorsal closure.

1.5.4.1 Dorsal closure forces are sensed and regulated: implications of the force

balance equation.

Laser incision experiments demonstrated several key findings. First, they defined

the two contractile elements that drive dorsal closure (the purse string and the

amnioserosa), one that resists closure (the lateral epidermis), and the relative

contribution of each. Second, subsequent quantitative analysis determined that the

individual forces that contribute to the progression of closure are large compared to the

net force that drives closure (b dh/dt). The individual forces in closure are 2 orders of

magnitude larger than the net force that drives closure (b dh/dt). These large individual

forces are nearly balanced such that closure proceeds at only ~6.0 nm/s (Hutson et al.,

2003; Toyama et al., 2008), but how they are coordinated to achieve this is unknown.

Finally, laser ablation studies demonstrate that closure is robust and resilient, such that

the removal of one (but no more than one) of the contributing forces does not

permanently impede the progression of closure. In fact, the removal of one process (e.g.,

seam formation) leads to the upregulation of force production in another (in this case,

amnioserosa constriction; Peralta et al., 2007).

1.5.4.2 Dorsal closure forces are sensed and regulated: experimental evidence. 34

A key experiment for understanding that forces are sensed and regulated during

dorsal closure is canthus nicking. In this protocol, lesions introduced in the amnioserosa

at either or both canthi inhibits seam formation (Hutson et al., 2003; Peralta et al., 2007).

Canthus nicking experiments were initally performed to show that zipping is not

required for the movement of the leading edges towards the midline, but rather

influences the completetion of dorsal closure through seam formation. When one

canthus is inhibited, the rate of zipping is upregulated at the other canthus (> 50 μm

across the tissue; Peralta et al., 2007).

When both canthi are inhibited, closure proceeds, but a local constriction with

reverse curvature of the leading edge develops where the remaining amnioserosa spans

the dorsal opening. This change in geometry is indicative of increased force production

in the remaining amnioserosa. Subsequent release of this tension by laser cutting and

measurement of the new σAS demonstrates that the force in the amnioserosa increases by

~37% in the double canthus nicking experiment. Together, these experiments

demonstrate the ability of both the amnioserosa and leading edge to react and

compensate for the loss of processes that contribute to closure. These consequences

imply that force sensing is occuring in the amnioserosa and the leading edge.

1.5.4.3 Mechanisms of mechanosensing and regulation during dorsal closure.

In order for forces to be regulated, they must be sensed at some biological level.

Some of the models of mechanotransduction outlined here – cell‑cell adhesions, focal

adhesions, MGCs – may participate in force sensing and regulation during dorsal

closure. The contribution of E‑cadherin based cell‑cell adhesion and integrin based cell‑

ECM adhesion has been established for closure (Tepass et al., 1996; Narasimha and

Brown, 2004; Gorfinkiel and Arias, 2007). However, dissecting the cellular requirement

for adhesion and the mechanosensing properties of junctional complexes is an imposing

challenge in embryos. E‑cadherin is required for embryos to develop, therefore null

mutations are difficult to study (Tepass et al., 1996). Integrins, specifically βPS integrin

encoded by myospheroid, are required to maintain tissue integrity during closure

(Narasimha and Brown, 2004) . Before the tissues fail in myospheroid mutant embryos, the

rate dorsal closure has been observed to be decreased to ~60% of wild type (Hutson et

al., 2003). But it is difficult to interpret this phenotype as being a direct result of integrin

function, its potential mechanosensory role, or the role of other proteins at the integrin‑

based adhesion complex. Non‑junctional proteins that have force‑dependent functions

(e.g., vinculin, α‑catenin) are expressed in Drosophila embryos, and so provide additional

targets for understanding how mechanotransduction signaling could affect closure. No

role has yet been established in the literature for MGC function in dorsal closure.

1.6 Central hypothesis.

One of the central motivations of this dissertation is to begin to address the

mechanism of cellular mechanosensing in a multicellular organism undergoing

morphogenesis. The following work builds upon the our understanding that dorsal 36

closure is a morphogenetic process that requires the generation and precise regulation of

forces. We hypothesize that forces for dorsal closure are regulated at the cellular level by

a mechanosensory circuit involving more than one mechanosensitive proteins. We

further provide biophysical, pharmacological, and genetic evidence that a key subset of

these mechanosensory proteins consists of mechanically gated ion channels.

2. Mechanically gated ion channels regulate cell shapes and movements during dorsal closure.

During morphogenesis, embryonic cells and tissues respond to genetic and cell signaling networks, generate physical forces, then change shape, move and/or proliferate (Wozniak and Chen, 2009; Lecuit et al., 2011). Such forces can in turn impact the course of development through regulation of gene expression, signal transduction and progression through the cell cycle (Farge, 2003; Hutson et al., 2003; Martin, 2010).

Intracellular (e.g., those due to activity of the cytoskeleton), extracellular (e.g., those due to adhesion), cell autonomous and non‑autonomous forces impact morphogenesis, providing a biomechanical circuit subject to regulation and feedback. An increased understanding of the molecular mechanisms that confer mechanosensitivity in vivo is therefore of wide interest.

How forces drive morphogenesis has been addressed extensively in Drosophila embryogenesis: during gastrulation, dorsal closure, and imaginal disc morphogenesis

(Hutson et al., 2003; Hufnagel et al., 2007; Martin, 2010). The kinematics and dynamics of dorsal closure, a cell sheet morphogenetic event, have been extensively described

(Harden et al., 2002; Hutson et al., 2003; Franke et al., 2005; Peralta et al., 2007; Solon et al., 2009). Closure is robust and resilient: individual forces for closure, generated in the amnioserosa or purse string, are approximately 2 to 3 orders of magnitude in excess of the net force that drives closure (Hutson et al., 2003; Peralta et al., 2007). Upon removal

of one contributing force by laser microsurgery, the remaining forces are upregulated to resume closure at nearly wild type rates (Peralta et al., 2007; Layton et al., 2009). When seam formation is blocked during closure by the laser ablation of tissue, the tension generated in the remaining amnioserosa increases. Since upregulation is observed to occur distant (≥ 10 cell diameters) from the site of laser perturbation, it is hypothesized that mechanical feedback is involved. However, the molecular basis by which tension is sensed and upregulated is not known.

Individual amnioserosa cells exhibit dynamic oscillations in apical cell area before and during closure. In closure, oscillations are associated with an overall decrease in apical area and contraction orthogonal to the embryonic midline (Fernandez et al.,

2007; Gorfinkiel et al., 2009; Ma et al., 2009; Solon et al., 2009; Blanchard et al., 2010;

David et al., 2010; Sokolow et al., 2012). The coordination of these behaviors across the epithelia towards closure requires a combination of cell signaling and mechanical feedback (Fernandez et al., 2007; Solon et al., 2009). However, it is unclear how or whether these two pathways are independent during closure. Mechanical feedback is hypothesized from a few key observations of contractility during closure and in silico.

First, there appears to be a supracellular organization of individual amnioserosa cell contraction across the tissue (Blanchard et al., 2010). These experiments tracked the apical cell contractions of >50% of amnioserosa cells through early‑mid dorsal closure.

Their results indicate that while there does not seem to be a predictable pattern to the

oscillations, the apical cell constriction of a given cell can be influenced by adjacent cells or even distant neighboring cells. Second, the in silico observation that in vivo behavior can be recapitulated when a cell’s contraction is triggered by its stretching past a critical point by neighboring, contracting cells (Solon et al., 2009).

Mechanically gated ion channels (MGCs) transduce mechanical force into intracellular signals. Pharmacological tools are used to manipulate MGC activity in vitro where these channels can be subjected to mechanical stresses through patch clamp approaches. MGCs were initially observed when researchers developed the patch clamp technique, in which a pipette (of varying diameter, but ~0.5 ‑ 5 μm is typical; Hamill,

2006) draws up cell membrane and proceeds to apply pressure while simultaneously recording current (Hamill et al., 1981; Neher and Sakmann, 1992). Using this technique it became clear that MGCs are prevalent in cell membranes. Application of the patch clamp remains the standard for determining if a channel is mechanically gated.

Early pharmacological tools to study MGCs included amiloride, gadolinium, lysophosphatidylcholine, and aminoglycoside antibiotics (Hamill et al., 1981). When these inhibitors are supplied in the patch clamp probe buffer, the activity of different types of MGCs is affected. Less is understood about how pharmacologically inhibiting

MGC activity affects developmental processes in vivo (Yang and Sachs, 1989; Wilkinson et al., 1998; Hamill, 2006).

None of the inhibitors listed above specifically inhibit MGC activity. Until recently, the best option was gadolinium (GdCl3) which interacts with negatively charged phospholipids (e.g., phosphatidylserine) leading to compaction of the plasma membrane (Ermakov et al., 2010). Increasing the rigidity of the membrane in this way is thought to increase the probability that MGCs will shift towards a closed state.

However, it is known that GdCl3 binds to a number of substrates, including other (non‑ mechanically gated) ion channels (Hamill and McBride, 1996), so the precise mechanism by which it affects channels and other biological targets is not clear.

In recent years, GsMTx4 has been developed as a specific MGC inhibitor

(Gottlieb et al., 2004). GsMTx4 is an inhibitory cysteine knot peptide inhibitor derived from the venom of the tarantula spider Grammostola spatulata (Bowman et al., 2007;

Kamaraju et al., 2010a). This inhibitor is characterized by a hydrophobic face surrounded by positively charged amino acids, and is thought to function by partially inserting itself into the plasma membrane. GsMTx4 is hypothesized to affect MGC gating indirectly through binding and penetration into the lipid bilayer, leading to deformations in the plasma membrane that are then conveyed to the channel. This is consistent with the finding that GsMTx4 acts as a gating modifier, rather than a traditional ‘lock and key’ inhibitor (Suchyna et al., 2004). GsMTx4 has been used to probe for MGC function in E. coli, neurons, muscle cells and in cell culture (Follonier et al., 2008; Kamaraju et al.,

2010b; Bae et al., 2011). Sensitivity to GsMTx4 is now frequently used as an assay for

determining the presence of MGC‑based current in patch clamp or cell culture conditions (e.g., Follonier et al., 2008; Bae et al., 2011)

Here, we investigate the molecular mechanism of force regulation during dorsal closure. We determine that pharmacological inhibition of MGCs by the peptide toxin

GsMTx4, or gadolinium (GdCl3), blocks dorsal closure in a dose‑dependent manner. We show that inhibiting MGCs leads to changes in actomyosin dynamics and subsequent cell behaviors, including oscillating apical constrictions. In support of the relevance for force regulation, we find that MGC‑inhibited embryos fail to recover from laser microsurgery. We show that dorsal closure responds to manipulation of both extra‑ and intracellular Ca2+ levels, consistent with the pharmacological evidence for MGCs. These results demonstrate a role for MGCs in dorsal closure, and indicate a mechanism for mechanosensing during morphogenesis.

2.1 Results

2.1.1 Pharmacological inhibitors of mechanically gated ion channels disrupt closure.

To determine whether MGCs are required for dorsal closure, we microinjected pharmacological inhibitors of channel function into embryos at dorsal closure stages.

Both the peptide toxin GsMTx4 and GdCl3 are inhibitors of MGC activity (Hamill and

McBride, 1996; Bowman et al., 2007; Arnadottir and Chalfie, 2010). We followed bulk tissue movements and analyzed cellular behavior in live embryos expressing a GFP‑ moesin construct that labels F‑actin (sGMCA; Figure 4B).

Figure 4. MGC inhibitors block dorsal closure in a dose‑dependent manner. (A) Schematic of dorsal closure. (B) Control microinjection of sGMCA embryos labeling F‑actin. (C) GsMTx4 injected sGMCA embryos (5 mM microneedle tip concentration) fail to close. Embryos in B and C were injected approximately 60 minutes before t = 0. T = 0 min is selected to compare two embryos at approximately the same total height, microns. (D) The progression of closure in control injected (dashed line) and GsMTx4 injected (solid line) embryos in B and C. (E) GdCl3 injected sGMCA embryos (50 mM tip concentration) fail to close. Scale bar, 25 μm; time in minutes. 43

2.1.1.1 Gadolinium (GdCl3) inhibits dorsal closure in a dose‑dependent manner.

GdCl3 is a non‑specific MGC inhibitor, nevertheless GdCl3 has been used in a number of studies as a preliminary indicator of MGC function. Microinjection of GdCl3 into dorsal closure staged embryos leads to a range of epithelial defects, which culminate in a failure to close (Table 1). At the highest doses (≥ 20 mM tip concentration), holes develop within the amnioserosa and at the junction of the amnioserosa and the leading edge of the lateral epidermis (Figure 4E). We do not observe a consistent cause of tissue failure (i.e., delaminating cells, tearing at the junctions between the leading edge and lateral epidermis). At lower doses, seam formation is delayed, and the overall morphology of the dorsal opening becomes abnormal (data not shown). This supports previous evidence that GdCl3 affects other stages of Drosophila development (Horner and Wolfner, 2008a).

Table 1. GdCl3 inhibits dorsal closure in a dose‑dependent manner.

Range of Effects (% of total) GdCl3 Tip injected Holes and Failure concentration N No Abnormal volume zipping to (mM) effect opening (pL) defects close 100 277 – 588 20 15 15 5 65 50 180 – 507 15 13.3 13.3 60 13.3 25 190 – 245 13 30.8 15.4 46.2 7.6 20 241 – 840 18 44.4 22.2 33.3 0 10 190 – 1560 40 55 30 15 0 1 230 ‑ 2157 16 56.3 43.7 0 0

2.1.1.2 GsMTx4 inhibits dorsal closure in a dose‑dependent manner.

The peptide toxin GsMTx4 is the most specific inhibitor of MGCs to date, and has no other known targets (Suchyna et al., 2004). We find that microinjection of GsMTx4 into dorsal closure staged embryos leads to closure defects and failure to close in a dose‑ dependent manner (Table 2). These defects essentially phenocopy effects seen with

GdCl3.

Table 2. GsMTx4 blocks dorsal closure in a dose‑dependent manner.

GsMTx4 tip Range of injected % Dorsal closure N concentration (mM) volume (pL) failure

20 137 ‑ 143 88.9 9 5 77 – 356 51.6 31 2.5 127 – 248 34.6 26 1 239 – 244 23 12 0 40 – 893 0 16

Failure of dorsal closure is ~52% penetrant (n = 31) at microneedle tip concentrations of 5 mM GsMTx4. Such tip concentrations result in an estimated final concentration between 100 and 150 μM, assuming free diffusion into a volume the size of the Drosophila embryo, and no sequestration of toxin. The actual concentration of

MGC‑inhibitor required to disrupt closure is not known, because to what extent the peptide is sequestered and to what extent not all volume is accessible is unknown. In embryos treated with 5 mM GsMTx4 epithelial integrity ultimately breaks down and

closure fails (Figure 4C). Together these data indicate that the activity of MGCs contributes to the progression of dorsal closure.

2.1.1.3 Etiology of closure failure in GsMTx4 treated embryos.

We further investigated the cause of closure failure due to GsMTx4 microinjection. We found that the dorsal epithelium becomes disrupted when apoptotic amnioserosa cells delaminate. In wild type embryos, approximately 10% of amnioserosa cells undergo apoptosis as dorsal closure progresses (Kiehart et al., 2000; Toyama et al.,

2008). These cells undergo an apical constriction until they are extruded from the cell sheet. In GsMTx4 treated embryos, apoptotic amnioserosa cells apically constrict and attempt to delaminate from the epithelium (Figure 5A). However, the epithelium becomes disrupted because the neighboring cells are unable to successfully maintain epithelial continuity while extruding the apoptotic cell (Figure 5A, blue star). As a result a hole forms (red star), which expands and leads to embryo failure.

We explored whether the failure of the epithelium in GsMTx4 treated embryos is due to processes associated with apoptosis or a failure of cell‑cell adhesion in general. To distinguish these possibilities, we blocked apoptosis in the amnioserosa by expressing

UAS‑p35 under the c381 amnioserosa GAL4 driver. In p35 expressing embryos, no apoptosis is observed but dorsal closure completes (Figure 5B). We find that GsMTx4 treatment of p35 expressing embryos still leads to dorsal closure cell shape defects but fewer embryos develop holes in the amnioserosa (Figure 5C). These results suggest that

MGC function plays a role in maintaining the integrity of cell‑cell adhesions, but that apoptosis is not required to generate the GsMTx4 phenotype.

Figure 5. Role of apoptosis in epithelial integrity of GsMTx4 treated embryos. (A) Time lapse of an apoptotic cell (blue star) in the amnioserosa of sGMCA embryos treated with 5 mM GsMTx4. Apoptotic cell attempts to delaminate, but contributes to a hole (red star) in the amnioserosa. Yellow star, reference cell. (B) Control injected embryo expressing p35 in the amnioserosa, with E‑cadherin‑GFP. (C) Embryo of the same genotype as (B), treated with 5 mM GsMTx4, closure arrests and amnioserosa integrity fails. Time in minutes, scale bar (A) 10 μm, (B) 25 μm.

2.1.1.4. Channel inhibition antagonizes force production in the amnioserosa

We investigated how inhibiting MGCs influences the forces that drive dorsal closure. The rate of closure, vnative = dh/dt, reflects the net force that drives dorsal closure, where h is the height of the dorsal opening above the dorsal midline, and t is time. In 47

uninjected or control injected embryos, closure occurs at a reproducible, linear rate of 5.9

± 0.9 nm/s (n = 7) consistent with published data (Hutson et al., 2003; Peralta et al., 2007;

Toyama et al., 2008). In embryos treated with 20 mM tip concentration GdCl3, the rate of dorsal closure is decreased, and vgadolinium = 4.4 ± 1.6 nm/s, n =5. The rate of closure in

GsMTx4 treated embryos is non‑linear (Figure 4D). We measured the average rate of closure after the initiation of seam formation but before reaching 0 nm/s and tissue failure and find that dh/dt is slower than wild type, vtoxin = 4.0 ± 1.8 nm/s (n = 6, p = 0.03).

It is not clear to us why GdCl3 treated embryos maintain a linear, but reduced, rate of closure while GsMTx4 treated embryos do not. Our results indicate that the force production during dorsal closure is affected by the inhibition of MGCs via GdCl3 or

GsMTx4.

Since dh/dt is related to the total balance of forces at the symmetry point (Hutson et al., 2003), we quantified force production in the amnioserosa after GsMTx4 treatment.

We performed mechanical jump experiments (Kiehart et al., 2000; Hutson et al., 2003;

Peralta et al., 2007) by using a steered microbeam laser to cut the amnioserosa away from the leading edge during closure (Figure 6). Following laser severing of the

Figure 6. MGC‑inhibited dorsal epithelia fail to regulate forces in response to laser microsurgery. (A) Control injected sGMCA embryo, edge cut time‑lapse. Panel labels 1) pre‑wounding, 2) immediately after wounding (dashed line indicates path of laser incision), 3) turning point, 4) closure. (B) 2.5 mM GsMTx4 injected embryo, edge cut time‑lapse. Panels labels 1, 2 correspond to those in (A), with 3’) corresponding to maximum height in (A), and 4) as the final time point. (C) The progression of closure for embryos in A, B. Labels 1‑4, 3’‑4’ correspond to panels in (A, B). Scale bar, 50 μm.

amnioserosa, the lateral epidermis rapidly recoils away from the dorsal midline. The recoil velocity (vrecoil) of the leading edge after the incision is directly proportional to σAS

(Toyama et al., 2008). The average vrecoil in embryos treated with 2.5 mM GsMTx4 is decreased from controls (629 ± 460 nm/s, n = 7 versus 1290 ± 300 nm/s, n = 7, respectively; p = 0.008). Together these results demonstrate that MGCs play a role in regulating the actomyosin structures and cell behaviors that produce force in the amnioserosa during closure.

2.1.1.5 MGC inhibition causes defects in amnioserosa actomyosin dynamics.

Effect of GsMTx4 on apical medial arrays.

To understand how MGC function affects the net force balance during closure, we examined the effect of channel inhibitors on the individual force generating tissues.

The amnioserosa contracts during dorsal closure, due to the apical constriction of individual cells in a coordinated manner via junctional belts and apical actomyosin networks (Kiehart et al., 2000; Hutson et al., 2003; Peralta et al., 2007; Toyama et al., 2008;

Gorfinkiel et al., 2009; Ma et al., 2009; Solon et al., 2009; Blanchard et al., 2010; David et

al., 2010). While the belts appear stable, the actomyosin networks rapidly assemble and disassemble across the apices of amnioserosa cells (Figure 7A). In the presence of

GsMTx4 the frequency at which these networks form is suppressed (Figure 7A’‑B) and the amplitude of apical area oscillations is dampened (Figure 7C). Some cells in treated embryos continue to display apical actin networks, presumably due to an uneven distribution of GsMTx4. Approximately 46.4 ± 10.1% of amnioserosa cells (n = 99 cells, see Methods) in 5 mM GsMTx4 treated embryos exhibited oscillating apical actin networks, compared to 92.7 ± 6.4% of cells in the amnioserosa of control‑injected embryos (n = 127 cells). We conclude that MGC function in the amnioserosa modulates apical actin network formation and function.

Figure 7. MGC inhibition interferes with F‑actin localization and dynamics. (A) Control injected sGMCA embryos exhibit apical actin meshwork (arrows), which are inhibited in the presence of 5 mM GsMTx4 (A’). Both panels are z‑projections. (B) Quantification of actin network formation in control or 5 mM GsMTx4 conditions. (C) Representative data indicating that amnioserosa cells from control injected sGMCA embryos (blue). Amnioserosa cells from 5 mM GsMTx4 injected embryos exhibit dampened oscillations (red). (D) The leading edge of a control injected sGMCA embryo. (D’) The purse string in the leading edge is disorganized in the presence of GsMTx4. (E) Organization of actomyosin at the junctional belts of amnioserosa cells in control injected embryos. (E’) Actomyosin at the junction belts of amnioserosa cells in embryos treated with 5 mM GsMTx4. Scale bars, 10 μm; *p < 0.0001.

Effect of GsMTx4 on junctional belts.

The other component of contractile behavior in the amnioserosa is the actomyosin activity at the junctional belt. Junctional belts are associated with cadherin adhesion, and thus are basal to the apical networks mentioned above. During dorsal closure in control embryos, some F‑actin is always associated with the junctional belt

(Figure 7E). Blocking MGCs by the addition of GsMTx4 leads to a reorganization of F‑ actin at the junctional belts (Figure 7E’). The distribution of F‑actin in MGC‑inhibited cells broadens, and the width of boundaries in GsMTx4 treated embryos increases compared to controls (1.45 ± 0.56 μm versus 0.8 ± 0.20 μm, respectively; n = 20 boundaries for each condition). Whether this reflects the additional recruitment of F‑ actin to the junctions or the failure to organize (e.g., bundle) wild‑type levels of F‑actin at junctions is unclear. Together, these data support a role for MGCs in regulating amnioserosa contractility.

2.1.1.6 Channels are required to upregulate forces for contractility and zipping in the lateral epidermis

We find that blocking MGCs with GsMTx4 also leads to defects in the lateral epidermis. GsMTx4 microinjection causes the supracellular purse string to become disorganized (Figure 7D‑D’). The distribution of actomyosin in the purse string broadens and seam formation via zipping is attenuated in GsMTx4 treated embryos compared to controls. We quantified changes in zipping in control and MGC‑inhibited

embryos at both the anterior (wa) and posterior (wp) canthi (Toyama et al., 2008). The average seam length in embryos displaying a GsMTx4 phenotype is less than controls

(Table 3). One hypothesis for why the seam length is disproportionally affected at the posterior canthus is that the distribution of GsMTx4 is not equal across the embryo. In fact, in the standard protocol for microinjection reagents are injected at the posterior end of the embryo. Previous studies have shown that zip/myoII contractility in the purse string is required to pull the leading edges into the canthus (Franke et al., 2005).

Therefore, the effect of GsMTx4 on purse string organization and seam formation suggests that regulation of actomyosin may be a general characteristic of MGC function during dorsal closure.

Table 3. GsMTx4 inhibition of channels leads to defects in seam formation during closure.

Control (μm) GsMTx4 (μm) p

wa 50.8 ± 19.9 36.8 ± 12.5 0.17

wp 38.5 ± 7.26 26.3 ± 10.1 0.03

We next determined the effect of MGC inhibition on force production in the lateral epidermis and purse string by analyzing the recovery of embryos from the edge cut protocol. Laser severing of the connection between the amnioserosa and the lateral epidermis leads to rapid recoil of the lateral epidermis away from the embryonic midline. Dorsal closure does not fail as a result of this severing: closure recovers, and progresses as a consequence of a new balance of forces generated in the lateral

epidermis, and potentially by wound healing in the amnioserosa (Peralta et al., 2007).

Following the incision, the leading edge recoils away to a turning point, hmax. After a brief recovery period, closure in untreated controls resumes in two distinct phases, vrecovery1

(17.0 ± 2.1 nm/s, n = 7) and vrecovery2 (3.9 ± 1.2 nm/s, n = 7; Figure 6C). Embryos treated with

2.5 mM GsMTx4 exhibit a range of defects. Severely affected embryos fail to resume closure (Figure 6B). Less affected embryos exhibit a single‑phase vrecovery = 7.1 ± 3.8 nm/s (n

= 7), indicating that forces are misregulated during the resumption of closure. GdCl3 injected embryos also exhibit decreased σAS (σAS = 800 nm/s ± 300, n = 6; 20 mM tip concentration) and failure to recover (data not shown). Taken together, these data indicate that MGCs contribute to force regulation in the dorsal epidermis.

2.1.1.7 GsMTx4 treatment affects the organization of cell‑cell junctions during closure.

GsMTx4 treatment leads to dramatic failure of tissue cohesion. We treated embryos ubiquitously expressing E‑cadherin GFP during dorsal closure with 5 mM

GsMTx4 to observe if MGC inhibition affects cadherin dynamics during closure. We find that GsMTx4 treatment leads to dramatic changes in cell‑cell boundaries in the amnioserosa and defects in E‑cadherin localization.

The first observed defect in E‑cadherin‑GFP embryos treated with GsMTx4 occurs at the cell‑cell boundaries of amnioserosa cells. In control injected embryos, individual cell‑cell boundaries in the amnioserosa display a degree of persistence, such that most boundaries appear as straight or gently curved lines (Figure 8A). Within

minutes after microinjection of MGC inhibitor, cell‑cell boundaries in GsMTx4 treated embryos become ‘wiggly’; the boundaries lack persistence and develop invagination‑ like curves in all amnioserosa cells (Figure 8B). Since the transition from straight to wiggly cell‑cell boundaries is indicative of decreased cortical tension (Lecuit and Lenne,

2007; Blanchard et al., 2010), we hypothesize that loss of wild‑type cell‑cell boundaries here reflects the loss of cortical tension in response to MGC inhibition.

Figure 8. Cadherin localization is disrupted in MGC inhibited embryos. (A) E‑ cadherin‑GFP localizes to the cell membrane in amnioserosa cells. (B) Embryos treated with 10 mM GsMTx4 develop irregular cell‑cell boundaries throughout the amnioserosa. (C) E‑cadherin‑GFP puncta develop in the cytoplasm of amnioserosa cells in embryos treated with 5 mM GsMTx4. (D) Example of a cell‑cell boundary in the amnioserosa of control (upper panel) and GsMTx4 treated (lower panel) embryos. GFP signal is lost from the cell‑cell boundaries after GsMTx4 treatment. Scale bar (A, C) 10 μm, (B) 25 μm.

As closure progresses in E‑cadherin‑GFP embryos treated with GsMTx4, cell‑cell junctions lose E‑cadherin. In control embryos, the localization of E‑cadherin to junctions is such that there are no gaps that lack GFP (Figure 8A, D). MGC inhibited embryos progressively lose GFP from cell junctions, resulting in boundaries with gaps. Moreover, cells in both the amnioserosa (Figure 8C, D) and lateral epidermis (data not shown) concomitantly accumulate GFP puncta in the cytoplasm. Given the observation that closure fails in MGC inhibited embryos when delaminating cells fail to maintain a continuous epithelium, these data suggest that cross talk exists between MGC activity and cadherin‑based cell‑cell junctions.

2.1.1.8 Early GsMTx4 treatment.

We next microinjected GsMTx4 into embryos prior to dorsal closure to determine if an earlier dose of MGC inhibitor could completely block the onset of closure. The developmental event that directly precedes dorsal closure is germband retraction (GBR), in which the caudal end of the embryo moves from a dorsal, anterior position to its final posterior position (Figure 9A, see also Figure 3A). At the end of GBR, the amnioserosa is exposed on the dorsal surface and early closure events begin.

Figure 9. MGC inhibition during germband retraction. (A) Embryo control injected during germband retraction. (B) Embryo injected with 5 mM GsMTx4 during germband retraction. T = 0 was selected for staging, both embryos were injected ~10 min prior to T = 0. Time in minutes; scale bar, 25 μm.

When we treat GBR embryos with 5 mM GsMTx4, we find that GBR initiates, but does not complete (Figure 9B). We observe movement of the germband towards the posterior, but the caudal end stops retracting just short of reaching the posterior, leaving part of the embryo folded over itself. These embryos start dorsal closure despite incomplete GBR, as we observe the dorsalward movement of the lateral epidermis and contraction of the amnioserosa. The observation that GsMTx4 treated embryos start

closure suggests that MGC function may not be required for the genetic and cell signaling program that initiates closure. However, dorsal closure is defective in early‑ injected embryos. We observe irregular apical cell shapes in the amnioserosa and failure to generate a robust purse string (Figure 9B). The terminal phenotype of embryos injected during GBR is due to epithelial holes that develop in the amnioserosa, followed by the failure of dorsal closure. Failure of early‑injected embryos phenocopies embryos injected at the onset of closure. In contrast, control injection does not affect GBR completion or the initiation of dorsal closure (Figure 9A). These results indicate that

MGCs may function in at other developmental time points in Drosophila embryogenesis, and that the progression of, but not the initiation of, dorsal closure requires MGC function.

2.1.2 Dorsal closure requires extracellular and intracellular calcium.

2.1.2.1 The effects of extracellular calcium chelation on dorsal closure.

Given that many MGCs conduct Ca2+ (Christensen and Corey, 2007), we next addressed whether the presence of free calcium (Ca2+) is required for dorsal closure. We hypothesized that microinjection of calcium chelators should partially phenocopy

GsMTx4 or GdCl3 treatment. The ionic composition of the fluid that occupies the perivitelline space (the extracellular space between the epithelium and vitelline membrane) surrounding the Drosophila embryo has been determined; the concentration of Ca2+ is ~ 5.0 ± 0.3 mM (van der Meer and Jaffe, 1983). We find that microinjection of

BAPTA, a cell‑impermeable Ca2+ chelator, in the perivitelline space causes dorsal closure defects. BAPTA treatment causes vnative to slow and the integrity of the amnioserosa to fail in a dose‑dependent manner (Table 4).

Table 4. Microinjection of calcium chelators inhibits dorsal closure.

Tip Chelator concentration N % DC defects % lethality (mM) 5 18 76.9 72.2 NP‑EGTA AM 1 13 69.2 30.8 0.5 18 28.5 38.9 75 33 75.8 87.9 50 43 34.4 74.4 BAPTA 25 13 0 0 10 16 0 0

Severely affected embryos at ≥ 50mM tip concentration of BAPTA fail to complete closure and exhibit disrupted epithelia (Figure 10A). In these embryos, the dorsal opening is elongated suggesting a zipping defect, and BAPTA injection also leads to a weak purse string in the lateral epidermis (Figure 10A’). Comparing BAPTA and

GsMTx4 microinjection conditions, the purse string is less organized in GsMTx4 treated embryos than in BAPTA treated embryos. However, the purse string in BAPTA embryos does not generate a smooth curve along the leading edge: with time, the leading edge bunches in places.

Figure 10. Chelating extra‑ and intracellular calcium disrupts dorsal closure. (A) sGMCA embryo injected with 50 mM BAPTA (tip concentration) in the perivitelline space. (A’) Leading edge and amnioserosa in a BAPTA injected embryo. (B) sGMCA embryo injected with 5 mM NP EGTA AM. (B’) Leading edge and amnioserosa in an NP EGTA AM injected embryo. Arrowhead indicates tissue failure. (C) E‑cadherin‑GFP embryo injected with 50 mM BAPTA. Time in minutes, where t = 0 is the time of injection. Scale bar (A) 25 μm, (B’) 10 μm.

Since epithelial failure may also be due to sequestration of Ca2+ from cadherin‑ based cell‑cell junctions, we observed embryos expressing E‑cadherin‑GFP during closure in the presence of BAPTA. Intracellular GFP puncta appear over time in BAPTA injected embryos, suggesting that junctions are affected. Nevertheless, cadherin‑GFP remains continuous at the cell‑cell membranes as closure slows and does so until the tissue fails (Figure 10C). Sites where cells are undergoing apoptosis do not appear to be

the origin of tissue failure in BAPTA treated embryos. Amnioserosa cells can undergo normal apoptosis and delamination in the presence of BAPTA, although rosettes (an arrangement of cells formed by the remaining neighboring cells when an apoptotic cell delaminates) persist, unlike controls (data not shown). Rosettes that persist do not expand into holes in the amnioserosa, unlike tissue failure in GsMTx4 treated embryos.

2.1.2.2 The effects of a cell‑permeable calcium chelator on dorsal closure.

To determine the contribution of intracellular Ca2+ in addition to extracellular

Ca2+, we microinjected the photolabile, cell‑permeable Ca2+ chelator NP‑EGTA AM

(NEA). We find that NEA also affected dorsal closure in a dose‑dependent manner

(Table 4). At lower concentrations of NEA (≤ 1 mM tip concentration), the morphology of dorsal closure is wild type. However, at these concentrations, vnative is decreased (4.90 ±

0.65 nm/s, n = 6; p = 0.04) relative to controls (5.90 ± 0.9 nm/s, n = 7). Higher concentrations of NEA induce epithelial disruption, disorganized purse strings and irregular amnioserosa cell shapes (Figure 10B, B’). These findings are consistent with

BAPTA findings, although NEA appears to be a more effective blocker of dorsal closure

(i.e., 5 mM NEA is sufficient to disrupt closure whereas 50 mM BAPTA is required to disrupt closure, Table 4). We conclude from these experiments that extracellular and intracellular Ca2+ is required for dorsal closure, and that one of the primary defects of chelating Ca2+ from this system is defects in actomyosin contractility.

2.1.3 Observing endogenous calcium dynamics during dorsal closure.

2.1.3.1 Generating a system to analyze endogenous Ca2+ during closure.

The net result of activating an MGC in cells is a change in conductance, or ion flux across the membrane. Changes in intracellular ion concentration can have profound effects on protein function, gene transcription and/or further ion flux (e.g., Ca2+‑induced

Ca2+ release). In cell culture, force‑induced MGC activity is associated with changes in intracellular Ca2+ (Matthews et al., 2006; Wei et al., 2009). Given the effect of Ca2+ chelators on dorsal closure epithelia, we hypothesized that Ca2+ flux is associated with contractile events in these cells. Specifically, we focused on the dynamic cell shape changes that occur in the amnioserosa as closure proceeds. We hypothesized that the oscillations in apical cell area observed in amnioserosa cells during early dorsal closure is associated with calcium flux.

We applied various fluorescent indicators to observe Ca2+ dynamics in the amnioserosa (Table 5). We found that cell permeable, microinjectable calcium indicators

(Oregon Green BAPTA‑1AM, FuraRed) sequestered in the yolk. This caused background fluorescence that obscured signals from the overlying amnioserosa and lateral epidermis. Therefore we generated or obtained a number of transgenic stocks of inducible, genetically encoded Ca2+ indicators. Second generation cameleons (D2, 3, 4‑ cpv) and GCaMP3 are Ca2+ reporters based on the interaction between the Ca2+ binding domain of calmodulin (CaM) and its target, the M13 domain of myosin light chain

kinase (Palmer et al., 2006). Upon binding Ca2+, the two domains physically interact each other. In the case of cameleons, this brings a CFP and a circularly permuted Venus into proximity with each other, leading to FRET (fluorescence resonance energy transfer). In the case of GCaMP3, a split GFP separates the CaM and M13 domains, and the presence of Ca2+ allows for GFP fluorescence.

Table 5. Calcium indicators used in this study.

Ca2+ sensitive Kd for Indicator Strategy References element Ca2+

Oregon green 0.17 Increased emmission (Paredes et

BAPTA 1 AM μM at ex488nm al., 2008) Decreased 0.13 (Takahashi FURA‑red emmission at μM et al., 1999) ex488nm CaM and MLCK FRET between (Miyawaki et cameleon 2.1 2 μM domains CFP/YFP al., 1997) CaM and MLCK FRET between (Palmer et D4cpv 64 μM domains CFP/cpVenus al., 2006) CaM and MLCK 0.66 (Tian et al., GCaMP3 Split GFP domains μM 2009) C2 domain of un‑ Interaction with PS (Oancea and C2 PKCβ known at membrane Meyer, 1998)

We were able to demonstrate the functionality of GCaMP3, via microinjection of the calcium donor ionomycin. We microinjected 3 mM ionomycin (tip concentration) into embryos expressing GCaMP3 and observed rapid activation of the reporter (Figure

11A). We also observed rapid contraction of cells in the embryo in response to ionomycin microinjection (Figure 11B), followed by widespread tissue failure (data not

shown). We conclude from this experiment that GCaMP is sensitive to rapid changes in

Ca2+ when expressed in dorsal closure tissues.

We expressed D4cpv or GCaMP3 reporters using the GAL4‑UAS system in the amnioserosa along with UAS‑RFP moesin in order to visualize cells (data not shown).

Using these reporters, we were unable to determine the relationship between Ca2+ flux and cell contractility. A high signal‑to‑noise ratio persisted in D4cpv or GCaMP3 expressing amnioserosa, in part due to high background signal from yolk auto‑ fluorescence.

Figure 11. Effects of ionomycin on GCaMP3 reporter and cell contraction. (A) An embryo expressing UAS‑GCaMP3 in the amnioserosa using c381‑GAL4 before (top panel) and after (lower panel) microinjection of 3 mM (tip concentration) ionomycin. Signal in top panel is due to yolk auto‑fluorescence. Arrow indicates amnioserosa cells that increase in GFP fluorescence as a result of microinjection. (B) An embryo ubiquitously expressing E‑cadherin‑GFP before (top panel) and after

(lower panels) microinjection of 3 mM (tip concentration) ionomycin. Cells rapidly contract, stretching neighboring cells. Time in seconds; scale bar, 25 μm.

2.1.3.2 Observing intracellular calcium with C2‑GFP.

Yu and Bement (2007) developed a novel calcium indicator based on the Ca2+ binding domain of Xenopus laevis PKCβ, called C2 (Oancea and Meyer, 1998). In the presence of Ca2+, this reporter is recruited to the plasma membrane through its interaction with phosphatidylserine. We generated UAS‑inducible C2‑GFP transgenics for reporter expression in Drosophila embryos. We expressed this reporter under an amnioserosa GAL4 (c381), and as a control we expressed the Ca2+‑insensitive mCD8‑GFP in the amnioserosa.

We were able to successfully observe dynamic localization of C2‑GFP fluorescence at the cell membranes in oscillating amnioserosa cells (Figure 12A). The combination of the c381‑GAL4 driver and the C2‑GFP responder is expressed mosaically, fortuitously providing contrast that aids in image interpretation. In a subset of cells, we observe increases in GFP localization to cell borders in contracting cells (n =

10/34 cells, 8 embryos). We also see GFP dynamics associated with contraction of individual boundaries (n = 16/34 cells, 8 embryos). These dynamics are not observed in amnioserosa cells expressing mCD8‑GFP (n= 12 cells, 3 embryos; Figure 12B). This data provides evidence that endogenous Ca2+ flux is associated with cell shape change during dorsal closure, but the signal‑to‑noise ratio is not optimal and the correlation between

increased membrane fluorescence (and therefore Ca2+) and contractility is not as robust as it might be (see discussion).

Figure 12. Endogenous calcium flux in amnioserosa cells. (A) Amnioserosa cell of interest expressing C2‑GFP is outlined in red. (A’) Time lapse of this amnioserosa cell through one cycle of contraction and relaxation. LUT key: blue is low GFP signal; yellow/white is high GFP signal. (B) Time lapse of an amnioserosa cell expressing mCD8‑GFP. Time in seconds; scale bar 10 μm.

2.1.4 Uncaging calcium in dorsal epithelia

We then asked how raising the level of Ca2+ in subpopulations of epithelial cells affected cell behavior. We uncaged NEA by targeting 3 ‑ 4 amnioserosa cells with a UV laser in embryos expressing both GCaMP3 to follow Ca2+ dynamics (Tian et al., 2009) and E‑Cadherin‑GFP to follow cell shapes. We observed rapid contraction of amnioserosa cells within seconds of uncaging (Figure 13A). Contraction was measured as the loss of area measured at the plane of the junctional belts. Cells in which Ca2+ was uncaged lost area at a rate (dA/dt) of 94.9 ± 35.5 nm2/s (n = 5; p = 0.002). The maximum rate of area loss in control oscillating cells was 3080 ± 1780 nm2/s (n = 16 cells), consistent with published values (Sokolow et al., 2012). Therefore, contraction due to uncaging Ca2+ is within physiological ranges, and the lower rate of contraction in cells targeted for uncaging may reflect the efficiency of Ca2+ release. Amnioserosa cells were observed to contract without respect to the embryonic anterior‑posterior axis, unlike wild type cells, which preferentially contract orthogonal to the embryonic midline (Blanchard et al.,

2010). Lateral epidermal cells display mild cell constrictions and produce apical actin projections subsequent to uncaging Ca2+ (Figure 13B). Following uncaging, embryos continue closure essentially normally, but cells do not relax back to their initial cell area or location observed prior to uncaging. Nevertheless they appear healthy, maintain junctions with neighbor cells, and do not undergo apoptosis. These results demonstrate

that dorsal closure epithelia require both extra‑ and intracellular Ca2+ and are capable of responding to experimentally induced increases in Ca2+.

Figure 13. Uncaging calcium triggers rapid cell contractions. (A) Uncaged Ca2+ induces apical constriction in the amnioserosa of an embryo expressing E‑cadherin GFP and c381‑GAL4>UAS‑GCaMP3. Cells are numbered for reference. (B) Uncaged Ca2+ induces weak apical cell constriction in the lateral epidermis and apical actin projections (red, arrows) in an embryo expressing E‑cadherin GFP and e22c‑ GAL4>UAS‑RFP moesin, UAS‑GCaMP3. Dashed line indicates cells targeted for uncaging. Scale bars, 10 μm.

2.2 Discussion

Successful development requires the coordination of forces across and among embryonic cells and tissues. Most cells have the capacity for sensing mechanical stimuli, notably through cell adhesive and integral membrane protein complexes (Janmey and

McCulloch, 2007). Through a combination of pharmacological and biophysical techniques, our study reveals a key role for ion channels during morphogenesis. We find that MGCs contribute to the regulation of the actomyosin cytoskeleton, a key mediator

of cellular force during dorsal closure. In support of the pharmacological evidence for

MGCs, we show that dorsal epithelia respond to changes in extra‑ and intracellular Ca2+.

Loss of channel function via pharmacological inhibition leads to disorganization of the actomyosin purse string and defects in amnioserosal apical actomyosin networks, structures known to be involved in force production during dorsal closure (Kiehart et al., 2000; Hutson et al., 2003; Peralta et al., 2007; Toyama et al., 2008; Ma et al., 2009;

David et al., 2010). MGCs also regulate tissue response to changes in tension generated by laser microsurgery, indicating a role in coordinating force across the dorsal epidermis.

Our data support a close relationship between channel function and adhesion in vivo: MGC inhibition leads to a dramatic loss of epithelial integrity. A key question that remains is whether the activity of MGCs influences actomyosin and cell‑cell junctions in parallel or in series, and if in series, in what order. Our findings are consistent with studies in cell culture showing that three components, MGCs, cell adhesions, and actomyosin cytoskeleton, function together in response to force. There is evidence that application of force via either adhesion molecule‑coated magnetic beads or general mechanical stimuli activates MGC‑dependent ion flux (Ko et al., 2001; Matthews et al.,

2006; Hayakawa et al., 2008). Additionally, loss of MGC function by pharmacological inhibitors or targeted mutations leads to defects in actomyosin contractile behaviors

(Guilak et al., 1999; Follonier et al., 2008; Wei et al., 2009).

Buffering endogenous Ca2+ and uncaging Ca2+ during closure revealed two important findings. First, increasing Ca2+ throughout the cell leads to large contractions in the amnioserosa and real, but subtler responses in the lateral epidermis, suggesting roles for Ca2+ in these tissues. Second, that increasing Ca2+ leads to isotropic amnioserosa cell contraction. We speculate that localized, sub‑cellular domains of ion flux contribute to the directional contraction of amnioserosa cells. Studies in cell culture illustrate how

Ca2+ flickers confer directionality to migrating cells by driving ion‑responsive cell contractile processes (Gomez et al., 2001; Fabian et al., 2008; Wei et al., 2009). It remains to be seen whether these flickers exist in dorsal epithelial cells, and to what extent normal intracellular flux influences cell behaviors in closure. Very few studies have addressed the role of calcium in Drosophila morphogenesis.

Cell shape changes associated with force production in the dorsal epidermis fail when MGC activity is reduced. The forces in closure must to be coordinated since individual tissue forces are orders of magnitude larger than the net force that drives closure (Hutson et al., 2003), and in response to laser microsurgery loss of one force leads to the upregulation of others (Peralta et al., 2007). σAS is reduced by GsMTx4 treatment; residual contractility seen after toxin treatment suggests the presence of additional regulators of actomyosin activity in the amnioserosa, and may reflect the dose used in these experiments. Our experiments demonstrate that functional MGCs are required to coordinate forces in a reproducible manner in the lateral epidermis during

wound healing. Although we do not yet understand the basis of the biphasic recovery we observe, MGC activity appears to regulate these transitions during the resumption of closure. These channels may have additional roles in wound healing, as shown with both TRP and DEG/ENaC channels in cell culture based wound‑healing assays (Berridge et al., 2003; Del Monaco et al., 2009). We hypothesize that these possibilities reflect a role in balancing force production between cells in native dorsal closure tissues. Given that morphogenesis throughout Drosophila development requires the assembly and regulation of force producing structures (Butler et al., 2009; Martin et al., 2010) it will be interesting to determine how other processes are affected by MGC inhibition.

Based on our observations, we hypothesize that MGCs function in a mechanical circuit serving to coordinate forces across dorsal closure epithelia. We propose that the contractile behavior of the epithelial cells and MGC activity are interdependent (Figure

14). As a single amnioserosa cell or domain of amnioserosa cells contract, it exerts a pulling force on neighboring amnioserosa cell(s). The neighbor domain is stretched, which triggers ion transients via MGC activity. This feedback loop could drive overall amnioserosa contraction by activating force producing molecules downstream of MGCs.

The intracellular ion response to applied force is known to promote cytoskeletal and junctional organization (Ko et al., 2001; Chan et al., 2004; Hayakawa et al., 2008;

Kobayashi and Sokabe, 2010). Several regulators of the contractile actomyosin cytoskeleton are ion‑responsive (e.g., myosin light chain kinase, gelsolin; Janmey, 1994),

and could mediate the cellular response to localized changes in ion concentrations.

Previous research suggests that actomyosin contractility during closure can act in a cell non‑autonomous manner, implicating a positive reinforcement of force producing activities or structures between and within the embryonic tissues (Franke et al., 2005).

Specifically, in the instance where two adjacent zipper‑/‑ cells express different levels of a zipper rescue construct (i.e., mosaicism), the contractility of the rescued cell has an effect on the behaviors and structures of the non‑expressing cell. Similar feedback loops are proposed for the oscillatory behavior of other mechanically coupled, contractile cell types (Follonier et al., 2008; Kruse and Riveline, 2011; Schillers et al., 2011). Mechanically gated ion channels therefore appear to mediate force production via the actomyosin cytoskeleton, providing a novel, molecular understanding of how physical forces impact morphogenesis.

Figure 14. Model of mechanically gated ion channel activity in dorsal closure epithelia. A schematic interpretation of MGC function in the amnioserosa (AS). In response to the contraction of a single amnioserosa cell (cell 1, T0) or domain of synchronized amnioserosa cells, neighboring cells are stretched (cell 2, T0), impinging on MGC activity in those cells. We hypothesize that at least some of those MGCs are stretch‑activated such that stretching leads to ion flux (cell 2, T0). In particular, Ca2+ ion flux leads to intracellular response by Ca2+‑binding proteins or other Ca2+ effectors, which in turn leads to actomyosin contractility (cell 2, T1).

3. Identification of candidate mechanically gated ion channel subunits for dorsal closure.

As discussed in Chapter 1, the three primary mechanisms of mechanosensing involve focal adhesions, cell‑cell adhesions and sensing along the plane of the plasma membrane. MGCs have been implicated in early developmental events associated with egg activation in Drosophila (Horner and Wolfner, 2008b). Research demonstrates a significant role for mechanical forces in generating shape change in later developmental stages. Dorsal closure during Drosophila embryogenesis requires the generation and regulation of actomyosin‑based forces (Kiehart et al., 2000; Hutson et al., 2003; Franke et al., 2005; Peralta et al., 2007). In Chapter 2, we demonstrated that dorsal closure staged embryos are sensitive to MGC‑specific pharmacological inhibitors. No known MGC subunits have been shown to be required for dorsal closure, although some MGC subunits are expressed in Drosophila embryos at other stages of development (Horner and Wolfner, 2008a; Graveley et al., 2011).

The lack of conserved genetic sequences or motifs that confer mechanosensitivity to specific MGCs presents a challenge for the identification of these channels (Arnadottir and Chalfie, 2010). For example, the Drosophila genome encodes approximately 140 presumptive ion channel subunits (Littleton and Ganetzky, 2000), and few are associated with mechanosensing (nompC, nanchung, inactive, painless, pickpocket, and dmpiezo;

Walker et al., 2000; Gong et al., 2004; Tracey et al., 2003; Adams et al., 1998; Kim et al.,

2012). 75

The genetics of dorsal closure is well established. Many genes have been identified which, when mutated, leads to dorsal closure defects. These genes can be grouped by the processes that they affect and the signaling pathways that they contribute to (Harden, 2002), and thus include genes associated with U‑shaped signaling, Jun N‑terminal kinase signaling, TGFβ signaling, Wnt signaling, and

Notch/Delta signaling. Additionally, several genes encode non‑receptor tyrosine kinases, small GTPases, pathways associated with junctional complexes (e.g., PDZ domain proteins) and cytoskeleton regulation (e.g., profilin). For many of these genes, removing or altering the function of the zygotic protein is sufficient to cause dorsal closure defects. For example, loss of function of the Wnt signaling receptor dishevelled leads to failure to accumulate actin and myosin at the leading edge of the lateral epidermis and elongation of these cells along the anteroposterior axis (Kaltschmidt et al.,

2002). However, determining the role of all of the genes that regulate dorsal closure is complicated by other factors. First, removing the zygotic contribution of a gene may not be sufficient to disrupt closure. For example, mutations in shotgun, the gene that encodes

E‑cadherin, do not cause closure defects when the zygotic contribution alone is affected

(Tepass et al., 1996). This is due to maternal load, and indeed when the maternal copy of shotgun is removed by germline clonal analysis, development cannot proceed. Therefore, in order to research the role of E‑cadherin in closure, germline clones of weak alleles have to be used (Tepass et al., 1996; Uemura et al., 1996).

A second challenge of identifying genes for closure is the potential for redundant function. This can be a problem for identifying genes in cell biological pathways in general. If the removal or mutation of a gene can be compensated for by other gene products, then it is possible for the penetrance of defects to be obscured. For example, we can consider the discovery of the bacterial MscL and MscS channels discussed in

Chapter 1. Initial studies of MscL were conflicting because although patch clamp performed on giant spheroplasts and purified liposomes demonstrated a channel conductance consistent with its presence and function, E. coli lacking MscL are capable of growing in hypotonic solutions (Sukharev et al., 1994). Further research demonstrated that mutants lacking the genes encoding both MscL and MscS fail to survive a transition to medium of low osmolarity (Levina et al., 1999). Subsequently, our understanding of

MscL, ‑S and ‑M is that these channels represent transmembrane valves of different sensitivities to stretch forces in the membrane, consistent with the force required to activate them under patch clamp. In a range of hypotonic challenges, MscM is activated first, then McsS, followed by MscL; MscM activity alone is insufficient to compensate for loss of MscS and MscL (Martinac, 2004). Although functional redundancy and maternal load represent two of the potential hurdles that can complicate the identification of genes products involved in dorsal closure, we also have some tools and strategies that can address these problems.

Previously we observed that the forces that drive dorsal closure are precisely regulated and sensitive to the pharmacological inhibition by the MGC specific inhibitor

GsMTx4 (Chapter 2). We hypothesize that individual MGC subunit(s) are expressed

(maternally, zygotically, or both) to function as force sensors in the embryo. Further, we propose that these proteins contribute to the overall morphology and distribution of actomyosin‑based forces in dorsal closure tissues via regulation of specific tissue behaviors and dynamics. Here, we use genetic and molecular geneic approaches to identify three loci that encode candidate MGC subunits that play a role in regulating cell and tissue behavior during dorsal closure: ripped pocket, dtrpA1, and nompC. We analyze their function in dorsal closure, alone and in combination with each other. We use biophysical techniques to evaluate their contribution to the force producing behaviors during closure. Finally, we determine their localization and expression pattern in dorsal epithelia. These studies identify and characterize the first MGC subunits to be implicated in dorsal closure.

3.1 Results

3.1.1 RNAi screen for MGC candidates

To identify MGC candidates that might be involved in dorsal closure, we performed a screen to test candidate genes that encode channel subunits previously implicated in mechanosensing in Drosophila and other systems (Table 6; Dietzl et al.,

2007). Previously, we showed that MGC inhibitors and Ca2+ chelators have distinct, yet

significant effects on the contraction of the amnioserosa and its attachment to the lateral epidermis (Chapter 2). In this screen, we focused on identified and predicted members of the transient receptor potential (TRP) channel family and degenerin/epithelial sodium channel family (Arnadottir and Chalfie, 2010). For each candidate, we drove expression of individual UAS‑RNAi constructs in the amnioserosa using MJ33a‑GAL4 and analyzed dorsal closure (Hrdlicka et al., 2002). Initially, we tested embryonic lethality, analyzed dorsal closure morphology, and determined the rate of closure (dh/dt), where possible

zTable 6. RNAi screen for MGC candidates in dorsal closure.

MGC Closure N RNAi Target dh/dt (nm/s) Family Defects (imaged) TRP ‑ 5.5 ± 0.8 6 Painless (TRPA) ‑ 5.1 ± 0.7 5 TRPA1 + n/a Nanchung (TRPV) ‑ 5.6 ± 1.4 7 TRP Inactive (TRPV) ‑ 5.5 ± 0.8 8 TRP‑γ ‑ 6.8 ± 0.5 3 TRPL ‑ 7.1 ± 1.3 4 TRPM ‑ 6.4 ± 0.8 3 nompC (TRPN) ‑ 5.2 ± 0.9 5 Ripped pocket + n/a Pickpocket ‑ 4.9 ± 1.0 6 CG8546 ‑ 5.3 ±1.2 5 DEG/ENaC CG15555 ‑ 6.2 ± 1.0 8 CG13278 ‑ 6.1 ± 1.4 6 CG33289 ‑ 6.1 ± 1.0 5 Control ‑ 5.9 ± 0.9 7

3.1.2 RNAi knockdown of RPK and dTRPA1 disrupts dorsal closure.

Our screen identifies two RNAi targets that result in both increased embryonic lethality and dorsal closure phenotypes: Ripped pocket (RPK) and dTRPA1 (Table 6).

RPK is a DEG/ENaC subunit expressed early in Drosophila embryogenesis (Adams et al.,

1998). dTRPA1 is a Ca2+ permeable TRP channel subunit required for larval thermosensing and locomotion (Rosenzweig et al., 2005; Hamada et al., 2008).

Knockdown of RPK and dTRPA1 subunits were verified as targets of RNAi knockdown via immunoblotting (Figure 15D, E).

Figure 15. RNAi knockdown of RPK and dTRPA1 disrupts closure. Dorsal closure (A) is severely affected by knockdown of (B) trpA1 and (C) rpk expression in the amnioserosa by RNAi. Arrowheads in (C) indicate the presence of segmental grooves. (A’) Lateral epidermal cells elongate and organize a contractile actomyosin purse string at their leading edge. Embryos expressing (B’) trpA1 and (C’) rpk RNAi in the amnioserosa have lateral epidermal cells that do not elongate and fail to generate a robust purse string. (D) Immunoblot with affinity purified anti‑RPK antibody demonstrates that RPK expression (~65 kD) is lost in RNAi expressing 81

embryos. 1, tubGAL4>UAS‑FL GFP RPK; 2, w1118; 3, tubGAL4>UAS‑rpk RNAi. (E) Immunoblot with anti‑trpA1 (to the fourth extracellular loop) demonstrates that dTRPA1 expression (~150 kD) is lost in RNAi expressing embryos. 1, sqhGAL4>UAS‑ dTRPA1 (endogenous TRPA1 runs at ~150 kD, construct runs at ~100 kD, arrowheads), 2, w1118; 3, sqhGAL4>UAS‑trpA1 RNAi. Scale bar, 5 μm.

3.1.2.1 General developmental defects associated with RNAi knockdown.

Expression of rpkRNAi in the amnioserosa led to defects in closure (see below), as well as other morphogenetic defects and increased embryonic lethality, thereby phenocopying key aspects of GsMTx‑4 treatment (Figure 15B; Table 7). Expression of trpA1RNAi in the amnioserosa led to similar dorsal closure defects (Figure 15C, Table 7).

In both cases, the dorsal opening contracts with a rectangular shape (compare to Figure

15A). In addition to the overall decrease in amnioserosa area, individual cells in this tissue decrease in apical cell area, suggesting that contractility is to some extent intact.

Wild type amnioserosa cells contract anisotropically, such that by late closure cells are elongated along the anteroposterior axis. Furthermore, wild type cells appear to contract first nearest the leading edge of the lateral epidermis resulting in a gradient of apical cell area, with the largest cell areas in the central amnioserosa and smaller cell areas towards the periphery of the tissue (Gorfinkiel et al., 2009). Both rpk‑ and trpA1RNAi expressing amnioserosa display the radial gradient of apical cell areas as in wild type (compare

Figure 15B and C to 15A), however individual cells appear to contract isotropically as dorsal closure progresses.

The lateral epidermis in rpk‑ and trpA1RNAi expressing embryos is clearly specified and distinguishable from the amnioserosa by a relatively straight boundary; however, the leading edge cells of the lateral epidermis fail to dorsoventrally elongate as in wild type and appear polygonal (Figure 15B’ and C’, compared to 15A’). Canthus formation is blocked, although it is unclear whether this is defect in seam formation, decreased σAS near the canthi, or a defect secondary to other morphogenetic events – in particular head involution and germband retraction (note that the defective canthus in

Figure 15C is the anterior canthus). Segmental grooves are pronounced in RNAi expressing embryos (Figure 15C, arrowheads) suggesting a delay in developmental progression.

Table 7. Rpk‑ and trpA1RNAi lethality and GAL4 driver strength.

GAL4 driver UAS‑RNAi Embryonic Lethality N MJ33a 69.5 ± 15% 462 c381 Rpk 31.2 ± 16% 196 sqh 16.7 ± 3% 284 MJ33a 42.0 ± 24% 514 c381 trpA1 30.2 ± 17% 430 sqh 6 ± 5% 234 MJ33a 50 ± 5% 305 c381 Rpk; trpA1 51.0 ± 6% 260 sqh 57.7 ± 7.1% 385 UAS‑GFP 11.5 ± 11% 265 MJ33a UAS‑Grim 20.7 ± 5.8% 260 UAS‑GFP 10.1 ± 5.4% 305 c381 UAS‑Grim 54.6 ± 6.1% 315

3.1.2.2 Specification of dorsal closure tissues via Jun N‑terminal kinase signaling.

The severe phenotype of RNAi expressing embryos made it unclear if the cells and tissues that participate in dorsal closure were specified for their correct fates. To verify that these dorsal closure tissues had been properly specified in RNAi expressing embryos, we investigated Jun N‑terminal kinase (JNK) signaling by evaluating the expression levels of puckered (puc). JNK signaling is a key component of dorsal closure whose activity is both an indication of and a requirement for the progression of closure.

In wild type embryos puc is expressed in the leading edge cells of the lateral epidermis

(Figure 16A). We observed puc expression in the leading edge of the lateral epidermis in embryos expressing rpk‑ or trpA1RNAi in the amnioserosa, comparable to wild type

(Figure 16B, C). Due to the nature of the RNAi phenotypes, it was unclear if the range of puc expression had expanded ventrally from the leading edge. Taken together, these data suggest that the channel activity of RPK or dTRPA1 contributes to the morphology and behavior of dorsal closure epithelia without necessarily affecting their developmental patterning.

Figure 16. JNK signaling in RPK and dTRPA1 knockdown embryos. (A) JNK signaling, visualized by expression of the β‑Gal reporter puce69, is active in the single 84

row of leading edge cells in the lateral epidermis (lacZ, green; F‑actin, red). (B) trpA1RNAi and (C) rpkRNAi expressing embryos display JNK signaling despite morphological defects. Scale bar, 25 μm.

3.1.2.3 Actomyosin defects associated with RNAi knockdown.

The partial progression of the lateral epidermal sheets toward the dorsal midline in embryos which express RNAi in the amnioserosa and display severe tissue morphological defects suggest that at least some of the actomyosin contractile elements that drive closure remains partially functional. In order to visualize actomyosin structures during closure, we time‑lapse imaged rpkRNAi or dtrpA1RNAi expressing embryos. We observe that amnioserosa cells in these embryos have dynamic apical actomyosin networks and junctional belts comparable to control embryos (Figure 15).

Actomyosin is also present along the cell‑cell boundaries in the lateral epidermis. These data suggest that the knockdown of a single MGC subunit is not sufficient to completely block actomyosin localization or kinematics.

The fourth notable actomyosin structure in dorsal closure tissues is the contractile purse string in the leading edge of the lateral epidermis. Although expression of RNAi is restricted to the amnioserosa, cells in the lateral epidermis of these embryos fail to generate a robust actomyosin purse string (Figure 15B’, C’). To determine the role of RPK and dTRPA1 in the lateral epidermis, we expressed RNAi in epidermal stripes using engrailed‑GAL4. Expression of rpk‑ or trpA1RNAi in the lateral epidermis alone decreases dh/dt (the rate of dorsal closure, defined in Chapter 1) without other closure 85

defects (Figure 17). We do not observe changes in actomyosin kinematics or cell shape changes in either the amnioserosa or lateral epidermal cells. However, we find that the rate of closure is decreased in both rpkRNAi (4.3 ± 0.6 nm/s, n = 8; p < 0.0001) and trpA1RNAi (4.1 ± 1.9 nm/s, n = 9; p = 0.009) expressing embryos compared to engrailed

GAL4 controls (6.6 ± 0.5 nm/s, n = 6). This data suggests that inhibiting channel function in this tissue compromises purse string contractility.

Figure 17. MGC knockdown in the lateral epidermis during dorsal closure. (A) A control embryo demonstrating the expression pattern of the lateral epidermal enGAL4, driving UAS‑GFP moesin in stripes. (B) enGAL4>UAS‑rpk RNAi and (C)

enGAL4>UAS‑trpA1 RNAi embryos do not display dorsal closure defects. Visualized with sGMCA; scale bar, 25 μm.

3.1.2.4 Effect of MGC knockdown on force production in the amnioserosa.

Since actomyosin contractile elements provide the bulk of the force that drives dorsal closure, we investigated how channel knockdown in the amnioserosa affects contractile forces for closure using laser surgery. Control embryos subjected to the edge cut protocol (described in detail in Chapter 2) exhibit recoil immediately after the laser incision, followed by a wound healing process that seals the hole in the epidermis, and finally the resumption of closure.

In response to a single 30 ‑ 40 μm lesion generated in the amnioserosa, embryos expressing either rpk or trpA1 RNAi in the amnioserosa exhibit tissue recoil (Figure 18A,

B). Recoil indicates the presence of force in this tissue. Since the snaking algorithm we use to determine σAS cannot follow the altered morphology of the dorsal opening, we cannot use this method to generate a value for σAS in these embryos.

Figure 18. Laser wounding of MGC knockdown embryos. (A) MJ33aGAL4>UAS‑rpkRNAi embryo targeted for wounding (top panel, dashed line), but embryo fails to heal. (B) MJ33aGAL4>UAS‑trpA1RNAi embryo similarly targeted for wounding also fails to heal. Visualized with sGMCA, scale bar, 25 μm. Time in minutes, where T = 0 is maximum height of wound.

Targeted tissue in either rpkRNAi (n = 6/6) or trpA1RNAi (n = 4/6) expressing embryos fail to complete wound healing. When these smaller incisions are made in

RNAi expressing amnioserosa we observe some behaviors associated with wound

healing, including actin projections oriented towards the opening (data not shown). We often observe that the wound gets smaller, but it is unclear if this is due to active wound healing or the decreasing area of the dorsal opening (for example, Figure 18B). Together these observations suggest that regulation of force in dorsal closure epithelia in response to wounding is in part mediated by MGCs.

3.1.3 GAL4 and RNAi expression patterns.

In order to determine if widespread knockdown of these two candidate MGCs could lead to more severe defects, we ubiquitously expressed GAL4 under control of the spaghetti squash (sqh) enhancer promoter cassette to drive the RNAi. Sqh encodes myosin regulatory light chain (Karess et al., 1991). Surprisingly, we found decreased levels of embryonic lethality using this driver compared to the amnioserosa driver

MJ33a (Table 7).

These results led us to investigate the effect of several GAL4s on the embryonic lethality and dorsal closure phenotypes associated with RNAi expression. We used

MJ33a as the primary GAL4 driver in the RNAi screen based on a previous report that this GAL4 expresses in a subset of amnioserosa cells when crossed to UAS‑mCD8‑GFP

(Hrdlicka et al., 2002). Two additional GAL4 drivers, c289b11 and c381, also express in the amnioserosa; c381 is the strongest and most uniform of the three amnioserosa

GAL4s. A lethality test for each of these three amnioserosa drivers with either rpk or trpA1RNAi led to similar rates of embryonic lethality (Table 7). In order to visualize the

expression pattern of these four GAL4 drivers, they were crossed to UAS‑lacZ and stained for β‑gal. Consistent with previous reports, c289b11 and MJ33a express in single and small clusters of amnioserosa cells (Figure 19A, B) whereas c381 expresses uniformly across the tissue (Figure 19C), and sqhGAL4 expresses lacZ ubiquitously

(Figure 19D).

Figure 19. Expression pattern of GAL4 drivers used in this study. Amnioserosa drivers (A) c289b11, (B) MJ33a, (C) c381 express at low, low, and high levels respectively in the amnioserosa as determined by lacZ staining, green. Ubiquitous driver (D) squashGAL4 drives throughout the embryo. Actin is shown in red. Scale bar, 25 μm.

To further explore the expression of GAL4 in these drivers, we crossed amnioserosa drivers to UAS‑grim to evaluate their ability to induce apoptosis in the tissue. MJ33a and c381 led to comparatively lower and higher rates of embryonic lethality, respectively, consistent with their lacZ expression levels (Table 7). We interpret the finding that weaker Gal4 drivers in combination with UAS‑RNAi responders leads 90

to a more severe phenotype than with ubiquitous drivers to mean that the distribution of MGC function across the dorsal tissue may be just as an important a factor in disrupting closure as is the knockdown of MGC subunit expression (see discussion).

3.1.4 The effect of mutations in MGC candidates on dorsal closure.

To further investigate roles for trpA1 and ripped pocket during embryogenesis, we analyzed embryos homozygous for severe alleles of dtrpA1, dtrpA1ins (Rosenzweig et al.,

2005) and dtrpA11 (Kwon et al., 2008) and a hypomorphic allele of rpk, rpk53 (Horner and

Wolfner, 2008).

3.1.4.1 dtrpA1 null embryos do not exhibit dorsal closure defects.

Both null alleles dtrpA11 and dtrpA1ins were generated by ends‑in targeted homologous recombination (Hamada et al., 2008; Kwon et al., 2008). Given the effects of

RNAi on dorsal closure, we were surprised to find that neither trpA1 mutant exhibited significant embryonic lethality, and homozygous null flies are viable and fertile (Table

8). We examined dorsal closure in homozygous dtrpA1ins embryos expressing sGMCA and did not observe defects in dorsal closure (Figure 20A), change in dh/dt (dh/dt = 5.52 ±

1.0 nm/s, n = 8), or σAS (vrecoil = 1535 nm/s, n = 3) compared to controls (dh/dt = 5.9 ± 0.9 nm/s, n = 7; vrecoil = 1290 nm/s ± 300 nm/s, n = 7)

To explore why trpA1RNAi should effectively disrupt dorsal closure unlike the dtrpA1null genotypes, we determined the embryonic expression pattern of dtrpA1. We used a trpA1‑GAL4 to drive expression of a UAS‑GFP actin reporter and observed GFP

expression in the amnioserosa and lateral epidermis of dorsal closure staged embryos

(Figure 20C). Auto‑fluorescence of the yolk (labeled) partially obscures the GFP signal from amnioserosa cells (arrows). We identified GFP positive cells in the lateral epidermis as well (data not shown). Together these data suggest that dTRPA1 is expressed in the dorsal epithelium.

Table 8. Embryonic lethality associated with mutations in dtrpA1 or rpk.

Embryonic Embryonic Genotype N p Lethality w1118 7.8 ± 3.8% 400 ‑ MJ33aGAL4 > UAS TRPA1 42.0 ± 24% 514 0.03* RNAi trpA11 12.0 ± 6.5% 361 0.31 trpA1ins 26.6 ± 19% 525 0.1 trpA1ins, MJ33aGAL4/ 8.7 ± 7.8% 208 0.83 trpA1ins, UAS TRPA1 RNAi MJ33aGAL4 > UAS RPK 69. 5 ± 15.0% 462 0.0002* RNAi Rpk53 (M+/Z‑) 20.2 ± 14% 402 0.1 Rpk53 (M‑/Z+) 33.7 ± 6.4% 210 0.004* Rpk53 (M‑/Z‑) 34.0 ± 9.4% 159 0.002* Rpk53/Df(3R)ED5092 (M‑/Z‑) 43.4 ± 6.2% 407 < 0.0001* RpkEY12268 29.5 ± 5.4% 317 0.0006*

One hypothesis for the difference between dtrpA1null and trpA1 RNAi knockdown phenotype and embryonic lethality could be if there is an off‑target effect of trpA1 RNAi. To investigate this possibility we expressed trpA1 RNAi via amnioserosa‑

GAL4 in the dtrpA1ins background. We find that in contrast to driving RNAi in a wild‑ type TRPA1 background, lethality due to trpA1 RNAi is dependent on the presence of

dtrpA1 target (Table 8). This indicates that the RNAi appropriately targets trpA1 mRNA in wild type embryos. It remains unclear why tissue‑specific knockdown generates increased lethality and morphogenetic defects compared to mutant alleles of dtrpA1 (see discussion and Chapter 4).

Figure 20. Effect of rpk and dtrpA1 mutants on dorsal closure. (A) The characterized null allele dtrpA1ins does not exhibit closure defects. (B) trpA1GAL4>UAS‑trpA1 RNAi embryos do not exhibit closure defects. (C) trpA1GAL4>UAS‑actin GFP expresses in amnioserosa cells (arrows). (D) rpk/Df embryo that has failed to finish germband retraction (arrowheads). (D’) The leading edge cells of rpk/Df embryos fail to elongate or accumulate F‑actin in the purse string. (E) rpk/Df embryos can complete GBR, but still exhibit severe dorsal closure defects. Scale bars, (A, B, D, E) 25 μm; (C, D’) 5 μm.

3.1.5 The effect of rpk mutations on dorsal closure.

3.1.5.1 Developmental defects in rpk53 embryos.

Zygotic RPK (rpk53Z) mutants exhibit a low level of embryonic lethality and morphogenetic defects that phenocopy rpkRNAi defects. We performed a Southern blot and sequence analysis of the rpk53 allele and determined that rpk53 represents an incomplete P‑element excision event that does not alter the protein coding region of ripped pocket (Appendix A). To further reduce embryonic RPK levels, we crossed homozygous female escapers (rpk53/rpk53) to males carrying the deficiency Df(3R)ED5092 and analyzed rpk53M/Df(3R)ED5092 (Rpk/Df) heterozygous embryos. This led to an increase in both embryonic lethality and penetrance of morphogenetic defects (Table 8).

Of unhatched rpk/Df embryos (n = 103, 34.3% of total), 78% fail to develop recognizable cuticle or morphogenetic structures, 11% develop cuticle with clear morphogenetic defects (see below) and the remaining 11% are indistinguishable from wild type but fail to hatch. These observations are consistent with a maternal load of rpk gene product

(Adams et al., 1998). Finally, we performed an immunoblotting assay to determine the level of RPK expression in rpk53 homozygous escaper males compared to wild type and found that RPK was present at reduced levels (Figure 25B).

The 11% rpk/Df embryos that make cuticle but fail to undergo normal morphogenesis fail to complete germband retraction, head involution or dorsal closure, thus phenocopying RNAi knockdown and early GsMTx‑4 treated embryos (Figure 20D‑ 94

E). Live imaging analysis of rpk/Df embryos shows that rpk/Df embryos often fail to fully retract the germband before initiating dorsal closure (Figure 20D). Lateral epidermal cells fail to elongate in the dorsoventral axis or produce an actomyosin purse string

(Figure 20D’). rpk/Df amnioserosa cells maintain actin cytoskeleton at the cell‑cell boundaries and in apical networks. Overall our data support a role for RPK in morphogenesis, but indicate that lowered expression of RPK alone is not sufficient to block all force producing structures and behaviors.

3.1.5.2 Embryonic RPK expression and localization.

To investigate the distribution of RPK protein in Drosophila embryos, we used an affinity purified peptide antibody against RPK and show that RPK is expressed at low levels throughout the embryo (Figure 21A). RPK is localizes to puncta that in most cases form patches on the apical membrane of amnioserosa cells (Figure 21B). RPK appears to be excluded from junctions, as seen by failure of RPK staining to overlap with anti‑ phosphotyrosine labeling. Embryos expressing rpkRNAi in the amnioserosa and rpk/Df embryos have both fewer and less intense RPK puncta and lack apical patch distribution in the amnioserosa (Figure 21C‑D). Finally, embryos homozygous for the deficiency

Df(3R)ED5092 do not exhibit apical RPK puncta in the amnioserosa (Figure 21E)

Figure 21. RPK localization in dorsal closure tissues. (A, B) An affinity purified anti‑RPK antibody stains apical patches in the amnioserosa of a w1118 embryo. (C) MJ33a>UAS‑rpk RNAi or (D) Rpk/Df embryos lack apical RPK patches in the amnioserosa. Yellow line delineates amnioserosa. (E) Homozygous deficiency embryo (sGMCA; Df(3R)ED5092) stained with anti‑GFP and anti‑RPK antibodies lack apical RPK patches in the amnioserosa. Scale bars, 25 μm.

To further explore the dynamic localization of RPK in dorsal closure epithelia, we constructed a UAS‑inducible, full‑length, GFP‑tagged RPK transgenic line. We co‑ expressed this construct along with UAS‑RFP moesin (to label the F‑actin cytoskeleton) using the amnioserosa driver c381‑GAL4. We find that this construct localizes strongly to the nuclear membrane (not shown), but also to puncta on the apical cell surface

(Figure 22). Due to the differences in localization between GFP‑RPK and RPK staining, we performed immunofluorescence using anti‑RPK and anti‑GFP antibodies and observed some co‑localization (arrows in Figure 22B). The discrepancy between anti‑

RPK staining and UAS‑GFP RPK may be due to artifacts introduced by fixation methods or RPK overexpression.

Figure 22. GFP‑RPK localization. (A) Still image from the amnioserosa of a live‑imaged embryo expressing GFP‑RPK (red) in the amnioserosa (c381‑GAL4; ubi‑ RFP‑moesin > UAS‑GFP‑RPK). (B) Amnioserosa of a fixed embryo expressing GFP‑

RPK under the sqh‑GAL4 driver. Arrows point to patches where anti‑GFP (green) and anti‑RPK (red) staining appear to co‑localize. Scale bar, 10 μm.

We expressed the UAS‑FL GFP RPK transgene under either the c381‑GAL4 driver or the more ubiquitously expressing sqh‑GAL4 in the rpk53 background to test for functional rescue of lethality associated with rpk53. With either GAL4 we observe rescue of rpk53 lethality (as scored by the loss of the TM3Sb balancer in adult flies).

3.1.5.3 Ripped pocket P‑element imprecise excision screen.

Since immunofluorescence and immunoblotting experiments demonstrate that the rpk53 genotype results in the reduced expression of RPK, we undertook a P‑element imprecise excision screen in order to generate a null allele of ripped pocket. rpkEY12268 is an allele with a P‑element (EPgy2) insertion 32 bp 5’ of the rpk coding region, and this insertion line has expression levels of RPK comparable to wild type (data not shown). In order to mobilize the P‑element, we used a Δ2‑3 transposase stock (Δ2‑3, Sb/TM6B) in order to generate white‑eyed stocks representing individual excision events (see methods). Using this method, in our third attempted screen we generated 9 homozygous lethal stocks maintained over either TM3Sb or TM6B balancers, and 59 homozygous viable stocks.

We analyzed the 9 balanced stocks by Southern blot to determine the nature of the P‑element excision. We found that none of these lines carried deletions in the rpk coding region, but instead represented either precise excisions or incomplete excisions.

We analyzed the remaining 59 homozygous rpk excision lines by PCR, Southern blotting

and deficiency mapping. While 8 of the 59 recovered lines were identified as incomplete excisions (similar to rpk53), none of these excisions exhibited lesions to the rpk coding region by PCR. We expect that some of the incomplete excision lines may continue to be hypomorphic for Rpk expression, but to date have been unable to recover a true null allele. A true null for rpk may require targeted knock out (Gong and Golic, 2003).

3.1.6 The role of nompC (TRPN) in dorsal closure.

3.1.6.1 Dorsal closure defects in nompC mutant embryos.

NompC is a candidate MGC that was previously identified to have weak dorsal closure defects (A. Stewart, unpublished). NompC is a TRP channel that has been associated with defects in hearing, gravity sensing, and touch in Drosophila (Walker et al., 2000; Kamikouchi et al., 2009) and implicated in hearing in vertebrates (Sidi et al.,

2003; Shin et al., 2005). To identify a role for this channel subunit in dorsal closure, we analyzed the morphology and rate of closure in two nompC mutants. nompC3 is a null mutant encoding a premature stop codon and nompC4 is a point mutation, C1400Y, in a putative extracellular loop region between transmembrane domains 3 and 4 (Walker et al., 2000). As a residual mechanoreceptor signal is observed in the bristles of animals with this point mutation, and the nompC4 mutation does not appear to alter gene expression, it is possible that nompC4 encodes a subunit with altered function (e.g., dominant negative). Both mutations abolish or decrease mechanoreceptor currents in response to bristle touch in adult flies (Walker et al., 2000).

Figure 23. Characterization of nompC expression in dorsal closure. (A) nompC3 embryos appear morphologically wild type (sGMCA). (B) nompC4 embryos exhibit an irregular purse string (sGMCA). (C) The purse string is irregular in nompC4 (red triangles) embryos compared to sGMCA (black triangles) embryos as measured by deviation from a curve fitted to the leading edge. (D) Traces of dh/dt in 5 nompC4 embryos displaying non‑linear dh/dt. (E) Anti‑GFP staining of MJ33a‑GAL4>UAS‑ nompC‑L‑GFP localized to apical puncta (note that phosphotyrosine staining is out of focus). (F) nompC‑GAL4 drives expression of UAS‑mCD8‑GFP in the amnioserosa. (G) Δ29ANK‑GFP‑nompC expression in the amnioserosa (c381‑GAL4) leads to defects

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in closure. (H) Embryo expressing Δ12ANK‑GFP‑nompC in the amnioserosa (c381‑ GAL4). Scale bar (A, H) 25 μm; (E) 5 μm.

nompC4 embryos display defects in purse string formation in the leading edge

(Figure 23B). The actomyosin purse string in nompC4 embryos is less organized and fails to generate a smooth curved leading edge compared to wild type (compare to Figure

15A). We analyzed the deviation of nompC4 (n = 6) or wild type (n = 6) purse strings from curves fit to the overall shape of the dorsal opening. We fit the snaking data to a curve and then measured the average distance between snaked points and the curve (in microns). Wild type embryos maintain a smoothly curved shape with few deviations until the end of closure, whereas nompC4 embryos have a ragged purse string that frequently deviates from the fit curve throughout closure (Figure 23C). F‑Actin is distributed between junctional belts and apical actomyosin networks in the amnioserosa as in wild type. The frequency of apical actomyosin networks in the amnioserosa of nompC4 embryos is comparable to wild type (3.11 ± 1.29 networks/10 min, n =28; see

Figure 7).

nompC4 embryos also display a biphasic dh/dt. Closure initially proceeds at nearly wild type rates in these embryos (dh/dtinit = 5.9 nm/s ± 0.9, n = 8) but shifts to a slower rate for the remainder of closure (dh/dtfin = 2.47 nm/s ± 0.9, n =8). This transition is gradual and occurs when the height of the dorsal opening (h) is 15 – 20 μm from the midline

(Figure 23D). The tension generated in the amnioserosa is slightly lower in nompC4

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mutant embryos, as σAS is slower (1.36 μm/s ± 0.5, n =7) than wild type (1.81 μm/s ± 0.9, n

= 4).

In contrast, dorsal closure appears morphologically normal in nompC3 embryos co‑expressing sGMCA (Figure 23A). We determined that dh/dt is decreased (4.74 nm/s ±

1.4, n =5) compared to wild type (6.1 nm/s ± 0.8, n = 4). This suggests that there is some defect in force production in these embryos, although the cause of this defect is not immediately observable.

In order to understand more about the role of nompC in dorsal closure, we tested genotypes that result in varied expression levels of nompC protein in embryos. First, we tested whether acute knockdown of nompC expression could result in closure defects.

We drove expression of nompCRNAi in the amnioserosa under the control of either c381‑

GAL4 or MJ33a‑GAL4. We did not observe either increased embryonic lethality or morphological defects in dorsal closure as a result of RNAi expression (Table 6). This is in contrast with our analysis of TRPA1 and RPK, in which RNAi had a more severe effect than bona fide mutants.

Next we tested the maternal contribution of nompC by generating germline clones of nompC3. Embryonic lethality of embryos lacking maternal contribution (m‑) of nompC did not increase compared to lethality in embryos lacking only a zygotic copy (z‑;

Table 9). These negative results were unexpected, given the defects associated with the nompC4 allele. Therefore, we generated germline clones where the only expressed

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nompC transcript is nompC4. Strikingly, embryonic lethality in nompC4 germline clones is

100% penetrant (Table 9). This result could reflect two possibilities: first, redundancy in

MGC function, since the loss of channel expression via null mutants leads to a weaker phenotype than a mutant with a single amino acid substitution that otherwise encodes a wild type channel subunit. Second, that the subunit encoded by nompC4 has a dominant character generating homotetramers with altered function or that interferes with the function of endogenous wild‑type subunit interactors (i.e., in heterotetramers).

Table 9. Embryonic lethality associated with nompC mutants.

Embryonic Genotype Embryonic Lethality n w1118 7.8 ± 3.8% 400

nompC3 (m+z‑) 30.0 ± 19.2% 156 nompC3 (m‑z‑) GLC 25.4 ± 7.4% 182 nompC4 (m+z‑) 35.6 ± 13.8% 181 nompC4 (m‑z+) GLC 100% 195 nompC4 (m‑z‑) GLC 100% 125 nompC‑GAL4, UAS‑ 42.6 ± 8.59% 298 Δ12ANK‑gfp‑nompC

nompC‑GAL4, UAS‑ 57.7 ± 21.7% 300 Δ29ANK‑gfp‑nompC

3.1.6.2 Localization of nompC in dorsal closure tissues.

Since previous studies focused on NOMPC function in adult and larval mechanosensing, little is known about its expression pattern in embryonic tissues

(McQuilton et al., 2012). To that end, we used nompC‑GAL4 (Cheng et al., 2010a) to

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express UAS‑mCD8‑GFP, which drives GFP in all cells that endogenously express nompC. At dorsal closure stages, we detect a strong and specific expression of GFP in the amnioserosa (Figure 23F). This suggests that nompC is expressed in this tissue in wild type conditions.

To visualize the distribution of nompC protein in the amnioserosa, we drove expression of the UAS‑nompC‑L‑GFP transgene (Cheng et al., 2010a) under the MJ33a‑

GAL4 driver, and performed immunofluorescence staining. We observed GFP puncta on the surface of amnioserosa cells (Figure 23E), as determined by z‑plane relative to phosphotyrosine, which localizes to the sub‑apical junctional belts in the amnioserosa.

We took advantage of the mosaicism of MJ33a‑GAL4 and use neighboring cells that do not express nompC‑L‑GFP as internal controls (Figure 23E). Together these findings are consistent with our findings that mutations in nompC lead to defects in dorsal closure.

3.1.6.3 Expression of nompC truncation transgenes disrupts dorsal closure.

Full‑length nompC is comprised of six transmembrane domains and an N‑ terminus that includes 29 ankyrin repeats (Walker et al., 2000). These ankyrin repeats have been shown to be involved in protein localization and signal transduction and hypothesized to function in channel gating (Howard and Bechstedt, 2004; Cheng et al.,

2010a). Given our previously mentioned result that the single amino acid substitution mutant nompC4 has a more severe phenotype than the null nompC3, we hypothesized that expression of nompC transgenes encoding subunits with truncated ankyrin repeats

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could function in a dominant negative fashion. Cheng et al previously published two nompC truncation constructs missing part (Δ12) or all (Δ29) of the ankyrin repeats. We expressed these constructs under the nompC‑GAL4 driver and analyzed the effect on dorsal closure. We observed that the expression of each of these constructs leads to increased embryonic lethality (Table 9). Embryos expressing the Δ29 construct in the amnioserosa by either c381‑ or nompC‑GAL4 consistently display tissue failure in the amnioserosa, failure to organize an actomyosin purse string, scalloped leading edge, and failure to elongate leading edge cells (Figure 23G). These phenotypes were also observed in embryos expressing the Δ12 construct in the amnioserosa by the same GAL4 drivers.

However, most Δ12 expressing embryos appear wild type (Figure 23H). These results are consistent with a role for nompC in closure, and further identify the ankyrin repeat domain as important for nompC function in the amnioserosa.

3.1.7 An integrated approach to MGC knockdown in closure.

3.1.7.1 Inhibition of dTRPA1 and rpk simultaneously: pharmacological approach.

As mentioned above, knockdown of either dTRPA1 or RPK expression in embryos by RNAi leads to a more penetrant, and in some cases more severe, developmental defect than in null embryos. To address this, first we co‑expressed both trpA1‑ and rpkRNAi in the amnioserosa. We discovered that this led to an increase in embryonic lethality (Table 7), but did not alter the dorsal closure phenotype from that observed in embryos expressing either RNAi alone (data not shown).

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As an alternate approach, we combined pharmacological and genetic tools to perturb channel function in closure. HC‑030031 is a TRPA1 specific inhibitor (Eid et al.,

2008). To determine the effects of TRPA1 pharmacological inhibition on closure, we microinjected HC‑030031 into sGMCA expressing embryos. HC‑030031 affects dorsal closure in a dose‑dependent manner (Table 10). Although dorsal closure begins as wild type in treated embryos, at the highest tip concentration tested (20 mM) the movement in the dorsal epithelium progressively slows, the actomyosin purse string disassembles, cell shapes in the amnioserosa become irregular and holes often develop in the amnioserosa (data not shown). This phenocopies treatment with the pan‑MGC inhibitors, GsMTx4 or GdCl3. At lower doses of HC‑030031 (≤ 10 mM), closure completes as in wild type (Figure 24A), but dh/dt is decreased (3.54 ± 1.01 nm/s, n=5, p = 0.002) compared to control injected embryos (5.9 ± 0.9 nm/s, n = 7).

Table 10. HC‑030031 disrupts dorsal closure in a dose‑dependent manner.

HC‑030031 tip Embryonic Genotype n Phenotype concentration (mM) Lethality 20 29 85.6% S, Z, PS, H sGMCA 10 16 29.4% S, Z, H 5 6 33.3% S weak: S, H MJ33a‑ 10 51 59.0% severe: H GAL4>UAS‑ weak: H rpk RNAi 5 5 80.0% severe: H dtrpA1ins 5 5 0% Key: (S) slow dh/dt; (Z) zipping defects; (PS) purse string defects; (H) holes in the epithelium and dorsal closure failure.

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We next microinjected HC‑030031 into embryos expressing rpkRNAi in the amnioserosa. We observe severe defects in dorsal closure when this inhibitor is injected into RNAi expressing embryos, at concentrations that do not block dorsal closure in wild type embryos. All rpkRNAi embryos injected, both those displaying a severe closure phenotype discussed above (Figure 15) and those that appeared wild type displayed these defects (Figure 24B, C). In general, dorsal closure initially slows, then the epithelium loses integrity and closure ultimately fails. Holes typically begin in the amnioserosa, particularly at the periphery where the amnioserosa meets the leading edge of the lateral epidermis. Other defects are also observed in the organization of the purse string, but with less penetrance. Microinjection of HC‑030031 into embryos null for TRPA1 (dtrpA1ins) did not delay or otherwise observably affect the progression of closure (data not shown).

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Figure 24. Microinjection of the TRPA1 inhibitor, HC‑030031, during dorsal closure. Time points (0, 60, 120 minutes, and final time point) of embryos treated with indicated mM (tip concentrations) HC‑030031 during dorsal closure. (A) sGMCA embryos are delayed but complete closure. (B) sGMCA; MJ33a‑GAL4>UAS‑rpk RNAi embryo that displays no closure phenotype develops holes in the amnioserosa (arrows) and fails to close when treated with HC‑030031. (C) sGMCA; MJ33a‑ GAL4>UAS‑rpk RNAi embryo that displays a severe closure phenotype stalls and develops holes in the amnioserosa (arrows). T = 0 min is ~3 minutes after microinjection. Scale bar, 25 μm.

In order to explore the relationship between dTRPA1 and RPK, we analyzed the expression of RPK in flies homozygous null for dtrpA1 via immunoblotting. We found that the amount of RPK present in either dtrpA11 or dtrpA1ins flies was decreased from controls (Figure 25A). In some blots, we observed weak RPK staining in dtrpA1 null flies 108

(data not shown), but expression levels were comparable to levels detected in rpk53 homozygous escapers (Figure 25B). This finding demonstrates that rpk expression is down regulated in flies that lack dTRPA1 protein. This could mean that the expression of these two genes is co‑regulated, but further work is needed to determine the nature of this down regulation.

Figure 25. Immunoblot analysis of RPK expression. (A, B) Lysate from single adult male flies of the listed genotypes and probed for (A) RPK expression (~ 65 kD; upper panel) and actin for loading control (lower panel). (B) RPK expression (lower panel), and zip/myoII for loading control (upper panel).

3.1.7.2 Analysis of double and triple mutant embryos.

We next constructed fly stocks with various combinations of mutant alleles of the three MGC candidates (Table 11). We determined embryonic lethality for most combinations of double mutant and triple mutant for the MGC candidates. Embryos of

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the correct genotype were sorted against fluorescent balancers when needed (see

Methods). Not all genotypes were stable over the fluorescent balancers (e.g., nompC3; dtrpA1ins). We next examined available double and triple mutant embryos for dorsal closure phenotypes by immunofluorescence and phalloidin staining.

Table 11. Embryonic lethality associated with combinations of nompC, rpk and dtrpA1 alleles.

nompC3 nompC4 dtrpA1ins rpk53 30.0 ± 19.2% nompC3 (156) 36.1 ± 16.6% 35.6 ± 13.8% nompC4 (239) (181) 53.0 ± 13.8% 12.0 ± 6.5% dtrpA1ins 72% (n = 200)** (142) (525) 65.5 ± 11.9% 79.6 ±4.5% 17.3 ± 11.6% 20.2 ± 14.0% rpk53 (200) (136) (185) (402) 85.0 ±4.8% 72.6 ± 11.5% dtrpA1ins, rpk53 (172) (166) ** nompC3; dtrpA1ins double mutants are not stable over fluorescent balancers. The embryonic lethality observed here is total lethality for the double mutant stock (including lethality associated with CyO and TM6Tb balancers).

Combinations of rpk53 hypomorphic and dtrpA1 null mutations did not increase embryonic lethality over levels observed with either allele alone, consistent with our immunoblotting results. The dorsal closure phenotypes observed in dtrpA1, rpk double mutant embryos were identical to those observed in rpk53 embryos (data not shown).

However, the combination of these two alleles is homozygous lethal and only balanced adults eclose, indicating that these alleles may interact at a later stage in development.

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In contrast, embryonic lethality was increased in all combinations including either nompC allele. In all cases lethality was comparable in embryos carrying the nompC4 allele versus the nompC3 allele (Table 11). However, we immunostained embryos phosphotyrosine and found that the double and triple mutant embryos we analyzed display some unique morphogenetic defects (Figure 26). Double mutants displayed a wider range of phenotypes. Weak phenotypes (Figure 26 A, A’) include irregular cell shapes in the amnioserosa and failure to dorsoventrally elongate lateral epidermal cells.

Strong phenotypes in double mutants include failure to complete germband retraction

(Figure 26C), as well as failure to complete dorsal closure (Figure 26B)

In triple mutant embryos, the amnioserosa is distinguishable from the lateral epidermis in these embryos (Figure 26D, E). The amnioserosa tissue is reduced in area, but since no seam formation is observable at either canthus, it is not clear that tissue area reduction is due to contraction or the presence of fewer cells. Amnioserosa cells have irregular shapes in this tissue (Figure 26D). The lateral epidermis is bunched along the anteroposterior axis, consistent with segment furrow formation and indicating that dorsal closure is delayed. Other epidermal tissue (e.g., head tissue) is often disrupted, suggesting that morphogenetic events besides dorsal closure are also affected. We interpret our observations of double and triple mutant embryos as an indication that the function of these three genes converges during embryogenesis. Specifically they function in dorsal closure towards the generation of wild‑type cell shapes and tissue movements.

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Figure 26. Embryos mutant for combinations of nompC, rpk, and dtrpA1 display dorsal closure defects. Embryonic genotypes are listed, and all embryos are stained with anti‑phosphotyrosine antibody indicating cell‑cell boundaries. (A) Arrow indicates a hole in the amnioserosa. (A’) Higher magnification image of the embryo in (A). (B, E) Dashed yellow line delineates the amnioserosa. (C) Arrow points to the posterior end of the embryo, which has failed to complete germband retraction. Scale bar (A) 25 μm; (A’) 10 μm.

3.2 Discussion

Here, we have demonstrated a role for three ion channel subunits encoded by ripped pocket, dtrpA1, and nompC, in regulating the process of dorsal closure. These three subunits are demonstrated to function in mechanosensing in Drosophila and other model 112

organisms. We show here that knockdown of expression of any of these three genes by

RNAi leads to dorsal closure defects. Furthermore, all three of these subunits are expressed in dorsal closure epithelia. We show that these subunits function redundantly, either with each other or with additional mechanosensitive interactors. Finally, we show that loss of function of these MGC subunits leads to defects in actomyosin localization and dynamics as well as cell shape changes during closure.

The mechanism by which pharmacological reagents like GsMTx4 and GdCl3 inhibit MGCs is not known (Bowman et al., 2007; Ermakov et al., 2010; Kamaraju et al.,

2010). Therefore, we sought to confirm and extend our pharmacological findings with a genetic approach. Our study identifies three genes encoding candidate channel subunits, ripped pocket, dtrpA1, and nompC, that are involved in Drosophila morphogenesis. Dorsal closure is disrupted by either the genetic or pharmacological perturbation of MGCs; similarities include failure of key actomyosin structures and cell shape changes.

Discrepancies in the phenotypes (RNAi vs. pharmacology) may be due to the difference between developmental timing and pattern of MGC knockdown or inhibition. Whereas our RNAi experiments heterogeneously decrease expression of a channel subunit in one tissue (amnioserosa), the pharmacological inhibitors are delivered at the onset of closure and encompass all tissues that express those genes. Interestingly, as shown in Chapter 2, when the MGC inhibitor GsMTx4 is microinjected at the onset of germband retraction we see a much similar phenotype to that of MGC knockdown embryos.

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As mentioned, a major effect of MGC knockdown is defects in actomyosin organization and contractility. RNAi expressing and mutant embryos lack a robust purse string and fail to generate the correct cell shapes in the lateral epidermis. We observe a reduction in the area of the amnioserosa in severely affected embryos although the purse string is disrupted such that the canthi never form and the localization of F‑actin to the leading edge is reduced compared to wild type. Since amnioserosa tissues exhibit recoil after targeting by laser microsurgery, it is clear that this tissue is under tension. Together these findings indicate that mechanical crosstalk between the lateral epidermis and amnioserosa exists in order to trigger leading edge cell elongation and purse string formation. RNAi experiments targeting either tissue independently demonstrate that the purse string and dorsoventral elongation defect only occurs when MGCs are knocked down in the amnioserosa. How the status of amnioserosa cell mechanosensitivity confers cell shape change to the lateral epidermis – whether it is a direct interaction between the peripheral amnioserosa cells and the leading edge or requires the whole amnioserosa – is unclear.

The distribution of cell shapes in the amnioserosa appears to be unaffected in embryos where a single channel subunit has been knocked down. Previously, it was suggested that this cell shape distribution might be due to a mechanical feedback either within the amnioserosa tissue or between the amnioserosa and lateral epidermis

(Gorfinkiel et al., 2009; Solon et al., 2009). If our hypothesis is correct that these genes

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encode functional MGCs for closure, this suggests that apical cell area distribution does not require mechanosensing input from this source. Since the integrin‑ and cadherin‑ based cell junctions that are required for dorsal closure remain intact in these embryos, we cannot rule out mechanosensing from these essential complexes in the dorsal epithelia. Indeed there is evidence that integrin‑based cell‑ECM adhesions are required for these cell shape distributions in the amnioserosa (Narasimha and Brown, 2004) as well as other morphogenetic gradients (e.g., DPP signaling, Fernandez et al., 2007).

Additional research will be required to determine the gating mechanisms of

RPK, dTRPA1 and NOMPC in embryonic epithelia, whether they are directly gated by forces or indirectly gated by additional mechanosensitive regulators. Still, there are several lines of evidence that lead us to believe that these subunits function in MGCs during closure. RPK is expressed in the dorsal epithelia at the appropriate stages of development. rpk mRNA is maternally loaded; RPK is GdCl3‑sensitive and is associated with egg activation, which is a mechanosensitive event during early Drosophila embryogenesis (Adams et al., 1998; Horner and Wolfner, 2008). NOMPC has been shown to be mechanically sensitive in vivo in Drosophila bristles (Walker et al., 2000).

Application of the patch‑clamp technique on homologs of dTRPA1 and RPK in heterologously expressing Xenopus oocytes or cell culture (TRPA1 and MEC‑4) supports the hypothesis that these channels are mechanically gated (OʹHagan et al., 2005; Kindt et al., 2007). However, our findings that these subunits do contribute to closure add

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another layer of regulation to what we already understand about this morphogenetic process. How MGCs fit into the landscape of genes and signaling mechanisms that drive closure is an exciting question. We can see already that MGC function is closely tied to regulation of the actomyosin cytoskeleton and that JNK signaling occurs without apparent perturbation. Furthermore, when MGC activity is perturbed, we see changes in the persistence of E‑cadherin‑GFP at the cell‑cell boundaries in amnioserosa cells. The role of actin cytoskeleton, JNK signaling and cadherin‑based junctions are well established in closure, and some of the signaling pathways are known (Harden, 2002); what remains to be seen is the mechanism by which MGC function impinges on them.

Our genetic experiments also raise the question of why dTRPA1 knockdown results in a more severe phenotype than knockout. We speculate that the embryo, which can compensate for the congenital loss of a channel subunit via transcriptional or translational regulation of other channels with some redundant function, cannot cope with a more acute loss of a channel subunit due to RNAi knockdown. ModENCODE temporal expression data for all three subunits indicates that they are highly expressed at various points during embryogenesis (Daines et al., 2011; Washington et al., 2011).

Significant RPK expression is observed at the earliest stages of embryogenesis, whereas

NOMPC and dTRPA1 expression is low early and increases by stage 14 of embryonic development (McQuilton et al., 2012). The amount of transcript required to express functional MGCs in embryonic tissues is not known.

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Our analyses of double and triple mutants, as well as other co‑knockdown or inhibition experiments suggest that these channel subunits may interact at some level.

TRP channels are known to be heteromeric – although NOMPC and TRPA1 have not been shown to interact, it has been hypothesized that the subunit constitution of an

MGC complex with TRP subunits can affect its function in a tissue (Cheng et al., 2010b).

To date, only one study has indicated that TRP channels and DEG/ENaCs have overlapping functions in larval environmental sensing (Johnson and Carder, 2012). We speculate that RPK, dTRPA1, and NOMPC form homomeric and/or heteromeric MGCs in the plasma membrane of dorsal epidermal cells (both amnioserosa and lateral epidermis) that respond to the forces generated by cell shape change and movements during dorsal closure. In response to force, channels are opened, leading to ion flux. In the case of dTRPA1and NOMPC, which are primarily permeable to Ca2+, ion flux can lead to local increases in Ca2+‑sensitive actomyosin contractility (via the activity of calmodulin and myosin light chain kinase). The activity of RPK channels, which are primarily permeable to Na+, could influence Ca2+‑sensitive actomyosin contractility through the additional activity of sodium‑calcium exchangers (NCX). Functioning the

‘reverse’ mode, these exchangers can transport three Na+ ions out of the cell while transporting one Ca2+ into the cell (Ruknudin et al., 1997; Schwarz and Benzer, 1997).

One of the three Drosophila NCXs (encoded by calX) appears to be maternally loaded

(McQuilton et al., 2012). This work adds to an understanding that DEG/ENaCs and TRP

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channels can function in parallel or perhaps in complex to affect the same output, which here is the regulation of actomyosin organization and contractility.

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4. Conclusions

The forces that drive dorsal closure to completion are robust, resilient and precisely coordinated across the dorsal surface of the Drosophila embryo. We hypothesized that mechanosensory circuits consisting of one or more mechanosensitive proteins regulated force production during dorsal closure. We present evidence that

MGCs have a role in regulating force production for closure. Treatment of embryos with the specific inhibitor GsMTx4 or the less specific inhibitor GdCl3 both disrupt dorsal closure and MGC inhibition causes epithelial breakdown. We find that epithelial failure may also be due to weakened cell‑cell junctions due to MGC inhibition. Force production is decreased in the amnioserosa when MGCs are inhibited; consistent with this we observe defects in actomyosin organization in both the amnioserosa and the lateral epidermis. These pharmacological findings suggest that ion fluxes may play an important role in closure. We find that chelation of extra‑ and intracellular Ca2+ decreases the rate of closure, while UV‑induced uncaging of Ca2+ causes rapid cell contraction. We discuss the challenges in and present data towards establishing the presence of Ca2+ flux in oscillating amnioserosa cells, and the relationship between flux and wild‑type apical cell contraction. We identify three genes that encode channel subunits whose knockdown or mutation leads to dorsal closure defects: ripped pocket, nompC and dtrpA1. We examine the defects associated with knockdown or mutation of these gene products alone and in combination. We observe that these subunits

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contribute to overall tissue morphology, actomyosin organization, and wound healing.

We provide evidence of the expression pattern of these channel subunits during dorsal closure. Together this research supports a novel role for MGC subunits in a cell sheet morphogenetic event.

4.1 Specific conclusions and future directions.

4.1.1 Identification of MGCs for dorsal closure: electrophysiological analysis.

We identified a role for MGCs in dorsal closure using a specific peptide inhibitor of MGCs, GsMTx4. This is the first time that this inhibitor has been used on a developmental system; our findings with this inhibitor support a role for MGCs in force production via actomyosin regulation. However, a key experiment to verify these findings is the application of patch clamp recordings. Applying patch clamp can not only establish the presence of MGCs but also provide information for their conductance, selectivity, and activation pressure (Hamill, 2006).

Ideally, we would apply the probe directly to either amnioserosa or lateral epidermal cells to record channel activity. However, the Drosophila embryo is completely encased by a vitelline membrane, which limits direct access to embryonic tissues. If the embryo could be removed from the vitelline intact, or even if the embryo was dissociated (where the tissue of interest was labeled), then we should be able to determine the presence of MGCs in these tissues. Furthermore, patch clamping would allow us to perform more precise pharmacological experiments. In the GsMTx4 and

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GdCl3 experiments presented in Chapter 2, we reported all concentrations in microneedle tip concentrations, rather than actual concentration in the embryo. As mentioned, we do not know what the actual concentration of inhibitor is at any given point in the embryo because we do not know how much of the space inside the vitelline is accessible to inhibitor, or what amount of inhibitor is bound off‑target. Providing either MGC inhibitor in the probe buffer of the patch clamp will allow us 1) to have a better understanding of the amount of inhibitor that blocks MGC activity in embryonic epithelia; and 2) to perform wash out experiments. MGC inhibition by GsMTx4 and

GdCl3 is known to be reversible (Hamill and McBride, 1996; Bowman et al., 2007), but the presence of the vitelline membrane occludes drug wash out experiments. Observing a mechanosensitive current under patch clamp conditions would give us substantial evidence that MGCs are present in dorsal closure epithelia.

4.1.2 Identification of MGCs for dorsal closure: heterologous expression systems.

An alternate (or complementary) approach to applying the patch clamp directly to dorsal closure tissues is to heterologously express the MGC candidates in Xenopus oocytes. Positive results from these experiments would demonstrate that the MGC subunits can form homomultimer channel complexes that are directly mechanosensitive.

Given our results that dorsal closure defects are more pronounced when more than one channel subunit is knocked down or inhibited, another experiment would be to co‑ express subunits. The expression of one subunit may not be sufficient to confer 121

mechanosensitivity (as in the case for MEC4 and MEC10 in C elegans; OʹHagan et al.,

2005), and co‑expressing subunits may give us further insight into whether these channels act in complex with each other. Indeed there is evidence that Drosophila TRP channels can form heteromeric channels in vivo (Gong et al., 2004).

4.1.3 Site directed mutation of MGC candidates.

Our experiments addressing the genetic contribution of MGC candidates to dorsal closure resulted in an unexpected result. We observed that alleles proposed to be molecular nulls (dtrpA1ins, dtrpA11, and nompC3) have weak to no dorsal closure defects.

In the case of rpk, we were unable to recover a null allele, so have no similar conclusion for this gene. One reason why this was unexpected is because expression of RNAi leads to dorsal closure defects. We concluded that embryos can compensate for the congenital loss of a channel subunit through redundant mechanisms, but cannot cope with the acute loss of a subunit via RNAi. In addition, the dorsal closure defects observed using

MJ33a (a weak and patchy amnioserosa driver) were not observed in embryos using ubiquitous drivers (i.e., sqh‑GAL4). We hypothesize that a mechanosensing mechanism exists across the amnioserosa such that an appropriately balanced expression pattern of

MGCs is critical for tissue morphogenesis. This hypothesis assumes that an individual cell has several means of sensing force, including MGCs (consider this wild‑type, tension sensing cell as cell type ‘A’). Loss of an individual MGC subunit by RNAi or null mutation leads to loss of one or more of these force‑sensing mechanisms (cell type ‘B’).

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Dorsal closure is unperturbed in tissue that is comprised of all A cells (i.e., dorsal closure in wild type embryos) or all B cells (i.e., null embryos, dtrpA1ins for example) because there is sufficient force sensing mechanisms in place that can be coordinated across the tissue. When MGC subunits are knocked down by MJ33a‑GAL4 expression of RNAi, the result is an amnioserosa tissue that is a mixed population of A and B cells. We hypothesize that a mixed population of cells, each with slightly different capacity for sensing force, are defective in coordinating forces across the tissue.

Another result that conflicted with our analysis of MGC null alleles is our observation of dorsal closure defects in embryos expressing nompC4 or nompC lacking ankyrin repeats (Chapter 3). We propose that these constructs and allele may have dominant characteristics (i.e., interacting with and altering endogenous channel function) or have the ability to form homotetrameric channels with altered function. One way to dissect MGC function during dorsal closure is to generate new alleles via site‑ directed mutagenesis. This may lead to new dorsal closure phenotypes that provide insight into wild‑type channel function. A general strategy for the mutants described below would be to generate transgenic flies carrying UAS‑inducible versions of subunit mutants, and express them in corresponding null or wild‑type backgrounds in order to observe their effects on dorsal closure.

4.1.3.1 Mutating MGC candidates: dTRPA1.

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Research on the chemical nociceptive function of dTRPA1 provided a phylogenetic analysis of TRPAs indicating that vertebrate and invertebrate TRPA1s share functional characteristics (Kang et al., 2010). Human dTRPA1 function has been shown to be perturbed by specific mutations that affect the gating and pore selectivity

(Wang et al., 2008; Benedikt et al., 2009). Of interest are the mutations N945A, which results in a constitutively active phenotype, and D918A, which decrease the permeability of TRPA1 to Ca2+. The pore region of Drosophila TRPA1 substitutes a glutamic acid residue at the site corresponding with D918; interestingly, a D918E substitution increases the Ca2+ permeability of human TRPA1 (Wang et al., 2008).

Recently, a molecular structure for mouse TRPA1 was proposed based on EM

(Cvetkov et al., 2011). These structures indicate an organization of TRPA1 in the membrane such that the cytoplasmic, N‑terminal, ankyrin repeats may interact with each other. This suggests that mutant TRPA1 constructs with ankyrin repeat deletions

(such as those generated for nompC) could have defective function when expressed in embryos. These specific mutations may give us more insight into the function of dTRPA1 during dorsal closure.

4.1.3.2 Mutating MGC candidates: RPK.

Previous research on DEG/ENaCs indicate what sites would be appropriate for initial targeting by site‑directed mutagenesis. The DEG mutation has been reported to lead to a dominant neurodegenerative phenotype in C elegans and leads to dominant

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active behavior of DEG/ENaCs expressed in Xenopus oocytes (Huang and Chalfie, 1994;

Waldmann et al., 1996; Adams et al., 1998). For RPK, this would constitute a single amino acid substitution at residue 524 to Valine (A524V; Adams et al., 1998). In heterologously expressing systems, this mutation causes a current 20 ‑ 50 times larger than wild‑type RPK currents.

An additional challenge with rpk will be to generate a null allele, which will allow us to both determine the requirement for RPK transcript in dorsal closure and provide a background to express RPKA524V for further studies. We were unable to generate a null by imprecise P‑element excision. However, methods taking advantage of homologous recombination (targeted ends‑in or ends‑out mutagenesis; Huang et al.,

2008) should allow us to generate a true rpk null allele. We already have in place a number of tools to verify the nature of new alleles (molecular biological, immunoblotting, and rescue constructs). The null alleles in dtrpA1 used in this study were generated by this method (Rosenzweig et al., 2005; Kwon et al., 2008).

4.1.4 RNAi screen for MGC candidates: dmPeizo.

We performed an RNAi screen for candidate MGCs in dorsal closure. Our screen targeted the knockdown of candidate genes that were 1) known to be involved with mechanosensory processes in Drosophila, or 2) homologs of genes encoding MGCs analyzed in other model systems. We tested 15 genes for dorsal closure phenotypes.

Since most characterized MGCs are TRPs and DEG/ENaCs, our screen would fail to

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identify any MGC not belonging to one of these families. Given that 140 genes in the

Drosophila genome encode known or putative channel subunits, making the assumption that we would identify candidate TRPs or DEG/ENaCs in order to narrow the focus of the screen was reasonable.

Since the RNAi screen in this work was completed, a new Drosophila MGC was discovered: Dmpiezo (Coste et al., 2010; Bae et al., 2011; Coste et al., 2012; Kim et al.,

2012). Dmpiezo encodes a large channel subunit (39 predicted transmembrane domains) that is conserved between vertebrates and invertebrates. The mouse Piezo1 homolog is inhibited by GsMTx4 treatment (Bae et al., 2011). In larvae and adults, Dmpiezo is expressed in sensory neurons and non‑neuronal tissues; Dmpiezo null mutants fail to respond to noxious mechanical stimuli (Kim et al., 2012). Finally, patch clamp recordings of DmPiezo expressed in HEK293T cells indicate that channel subunits homomultimerize and are directly mechanosensitive (Coste et al., 2012). Temporal expression profiles associated with this locus (Dmpiezo is also known as CG8486) indicate that this subunit has high expression levels during the first 12 hours of embryogenesis (McQuilton et al., 2012). We could request and use existing Dmpiezo reagents (Dmpiezonull, ‑RNAi, ‑GAL4, ‑GFP; Kim et al., 2012) in similar experiments as those outlined in Chapter 3 for rpk, nompC, and dtrpA1 to determine if this novel MGC subunit contributes to dorsal closure.

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4.1.5 Calcium signaling during Drosophila embryogenesis.

Given our results that calcium chelators and donors modify the behavior of dorsal closure epithelia (Chapter 2), we hypothesized that calcium signaling plays a role in regulating epithelial cell behavior. Namely, we aimed to show that native cell shape changes in the amnioserosa are associated with Ca2+ flux. Little is known about Ca2+ signaling during Drosophila embryogenesis (Creton et al., 2000; Webb and Miller, 2003;

He et al., 2010). Global waves of Ca2+ are observed as the embryo undergoes cellularization and is thought to be involved in establishing dorsoventral embryonic polarity (Creton et al., 2000). Ca2+ was also found to be necessary for follicular epithelium behavior in Drosophila oogenesis (He et al., 2010). In contrast, Ca2+ signaling has been observed to regulate morphogenetic movements such cell elongation during Xenopus and zebrafish convergent extension (Wallingford et al., 2001; Webb and Miller, 2003;

Zhang et al., 2011). Many other developmental processes exhibit dynamic Ca2+ transients that correspond with cell and tissue movements (Markova and Lenne, 2012). The downstream mechanisms affected by these calcium transients are unknown.

4.1.5.1 Membrane bound genetically encoded Ca2+ indicators.

As is, our findings contribute significantly to the understanding of how Ca2+ is required during dorsal closure and Drosophila embryogenesis, but fall short of providing a downstream mechanism for MGC function. Our results with the C2‑GFP Ca2+ indicator showed a weak correlation between Ca2+ flux and amnioserosa cell apical contraction.

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The first step towards trying to improve this data is to address our method of data acquisition (see methods, Chapter 2). For the majority of these experiments, we imaged

Ca2+ indicators by confocal microscopy (Zeiss Axiovert 200M, Kiehart lab or Leica SP5,

Duke LMCF). Given the high signal‑to‑noise ratio associated with visualizing

Ca2+indicators in the amnioserosa – in part due to auto‑fluorescence in the yolk – we may benefit from exploring other illumination and image acquisition options (e.g., TIRF microscopy).

In addition to this, we can make modifications to one (or more) of the Ca2+ indicators to alter their cellular localization. Since the Ca2+ signal that we are interested in (Ca2+ flux due to MGC activity) is located close to the cell membrane, we may find that a membrane targeted Ca2+ indicator provides a more sensitive reporter. Membrane targeted GCaMP3 has been reported and used in neurobiological studies (Shigetomi et al., 2010). Here, GCaMP3 is fused to the N‑terminal membrane targeting sequence of Lck

(a Src tyrosine kinase). The basis of our studies of endogenous Ca2+ signaling in the amnioserosa relied on being able to distinguish GFP fluorescence at the cell boundaries from cytoplasmic GFP. Using this reporter we were unable to observe Ca2+ dynamics at the apical membrane, which poses a problem for understanding the role of Ca2+ in the dynamics of apical actomyosin networks. Lck‑GCaMP3 could give us better signal‑to‑ noise since its activation (and subsequent GFP fluorescence) would be localized to areas of high Ca2+ along the membrane. This may be a sufficient reduction in cytoplasmic GFP

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signal for visualizing Ca2+ at the apical membrane. We can apply this tool to similar experiments to those described in Chapter 2, towards determining if Ca2+ flux is associated with apical cell oscillations in the amnioserosa. If the method provides robust and reproducible data, the next step would be to observe changes in Ca2+ dynamics upon treatment with MGC inhibitors or in genetic background lacking MGC subunits or expressing defective MGC subunits.

4.1.5.2 Alternative hypothesis for MGC knockdown: Ca2+ signaling and gap junctions.

We outlined one hypothesis for how pattern of MGC knockdown might be an important factor in the observed RNAi phenotypes (Chapter 3). This hypothesis was based on the coordination of mechanotransduction in single cells; an alternative hypothesis is that MGC knockdown affects the coordination of mechanotransduction across multiple cells through Ca2+ signaling and gap junctions. In this model, the heterogeneous tissue (a mix of A and B cells) behaves as described above. The loss of

MGC subunit leads to defective or altered Ca2+ flux in a B cell, which can subsequently affect neighboring cells (of the A or B subtype) by the transmission of Ca2+ through gap junctions. We expect that the spatiotemporal pattern of Ca2+ flux across the amnioserosa is not predictable from embryo to embryo, in the same way that apical area oscillations appear to stochastic between individual embryos (Solon et al., 2009; Sokolow et al.,

2012). However, it has been noted that there is a tendency for groups of amnioserosa cells to synchronize their oscillations in phase (Blanchard et al., 2010). It is not known if

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this is a random synchronization or coordination due to an unknown signaling mechanism. Since gap junctions are present in dorsal closure tissues and Ca2+ is required for closure to complete (Tepass and Hartenstein, 1994; Chapter 2), it is possible that these cells contract in phase due to the movement of Ca2+ through gap junctions. The disruption of robust Ca2+ signaling across the amnioserosa may be sufficient to disrupt the coordination of cell behavior in this tissue and the progression of dorsal closure.

Two major experiments could begin to address these hypotheses. First, determining the genetic requirement for genes that encode gap junction genes during dorsal closure (e.g., innexins, which function in place of vertebrate connexins in invertebrates; Bauer et al., 2005). Zygotic loss‑of‑function mutations in the gene kropf, which encodes Innexin‑2, leads to severe epidermal defects during late stages of embryogenesis (Bauer et al., 2004). Thus mutants and tools (RNAi, GFP‑tagged proteins) exist that can be useful for identifying a function for gap junctions during closure.

Second, experiments correlating spatiotemporal dynamics of Ca2+ across the whole amnioserosa as dorsal closure progresses with the patterns of cell shape change in this tissue. This may provide significant insight into whether Ca2+ signaling is coordinated across multicellular regions. Do we see coordinated Ca2+signaling in cell regions that contract in phase? These experiments can also contribute to understanding

Ca2+ signaling in patterns of contraction that we know are primarily regulated by cell signaling, for example, Ca2+ signaling in the peripheral amnioserosa compared to central

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amnioserosa cells. The critical challenge for this set of experiments is improved visualization of Ca2+ dynamics in the amnioserosa through expression of genetically encoded Ca2+ indicators and microscopy techniques.

4.1.5.3 Mechanisms of calcium signaling to the cytoskeleton.

The contraction of amnioserosa and lateral epidermal cells is mediated by actin and zip/myoII (Kiehart et al., 2000; Franke et al., 2005). Certainly, our experiment uncaging Ca2+ in amnioserosa and lateral epidermal cells demonstrates that Ca2+‑ responsive proteins and contractile elements can drive cell contraction (Chapter 2). In other non‑muscle cells, myosin activity is regulated by phosphorylation of the myosin regulatory light chain by myosin light chain kinase (MLCK). MLCK activity can be regulated by Ca2+/Calmodulin (CaM) complexes (Gallagher et al., 1997). The role of

MLCK and CaM during dorsal closure is not known, but genetic tools to target knockdown of their expression during development may provide a first step towards understanding their role.

4.2 Implications of this work.

Dorsal closure provides a model system for epithelial cell sheet morphogenesis and the study of force production during embryogenesis (Gorfinkiel et al., 2010).

Epithelial cell sheet morphogenesis is a conserved mechanism of development and wound healing. Force production is a ubiquitous characteristic of cells, and is increasingly found to be a critical component of cell and tissue morphogenesis. We

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provide the first evidence that MGCs contribute to regulating force production during dorsal closure. Consistent with channel function during closure, we show that manipulation of extra‑ and intracellular Ca2+ can affect cell behaviors and tissue organization during closure. These Ca2+ studies contribute to our current understanding of the role of Ca2+ in Drosophila morphogenesis. We identify three loci that encode channel subunits whose knockdown leads to dorsal closure defects. These MGC candidates were not previously known to participate in Drosophila embryogenesis, but we show that their knockdown, individually or in combination, leads to defects in cell shape changes and tissue movements. Finally, our results support a role for subunits from both TRP and DEG/ENaC families that converges on regulation of actomyosin‑ based shape changes during closure. The observations and analyses presented in this work give rise to a novel model for the coordination of force producing cell behaviors across the embryo during dorsal closure.

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5. Methods

5.1 Drosophila strains.

Unless otherwise specified, ‘wild type’ refers to fly stocks expressing sGMCA or ubi‑cadherin‑GFP, as described (Oda et al., 1998; Kiehart et al., 2000). The following

GAL4 drivers were used: MJ33a‑GAL4 or c381‑GAL4 (amnioserosa); sqh‑GAL4 or β‑ tubulin‑GAL4 (ubiquitous); e22c‑GAL4 or en‑GAL4 (epithelial) (Lawrence et al., 1995;

Manseau et al., 1997; Hrdlicka et al., 2002). Genotypes of embryos were established using fluorescent balancers: twi‑GAL4, UAS‑GFP, CyO; twi‑GAL4, UAS‑GFP, TM3Sb;

CyO, GMR, Dfd‑YFP; TM3Sb, Tb, GMR, Dfd‑YFP; or ubiquitin‑driven RFP‑moesin

(Singh and Kiehart, unpublished). For rpk‑ or trpA1RNAi experiments, VDRC stocks

#8549 and #37249 were used, respectively. Embryos were collected from fly crosses or stocks maintained at 25℃, or aged approximately 12 hours at 16℃.

5.2 Pharmacology and microinjection of Drosophila embryos.

Embryos collected on agar plates were dechorionated, sorted for stage and genotype, and mounted on glass coverslips. For microinjections, embryos were desiccated, covered with halocarbon oil, and injected with a Pico Injector (Harvard

Apparatus). GdCl3 and GsMTx4 were prepared to final concentrations between 1 – 100 mM in a 1X standard solution for extracellular injection into embryos (180 mM NaCl, 10 mM HEPES, 5 mM KCl, 1 mM MgCl2; Bowman et al., 2007) supplemented with 1mg/mL bovine serum albumin (BSA, Sigma). NP‑EGTA AM (Sigma) was prepared in 1X

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standard solution to final concentrations of 0.5 mM for lateral epidermis uncaging and 1 mM for amnioserosa uncaging. BAPTA (Invitrogen) was prepared in 1X standard solution and injected at concentrations as described. HC‑030031 (Tocris) was prepared to final concentrations between 1 – 50 mM in DMSO, and diluted to a 1X standard solution for extracellular injection into embryos. Control microinjection solution contained 1X standard solution with BSA alone.

5.3 Molecular Biology

5.3.1 D4cpv transgenics.

We generated transgenic stocks of D4cpv (D4.1), a FRET‑based genetically encoded Ca2+ indicator (Palmer and Tsien, 2006). D4.1 was cloned into the BglII site of the pUAST‑AttB vector (E. Spana), following digestion and blunting reaction of both the pUAST‑AttB vector and the D4cpv‑pBAD vector. Specifically, pUAST‑AttB was digested with Xho1 and blunted with the Quick Blunting Kit (NEB). Following heat inactivation of the blunting enzyme, the linearized vector was digested with BglII. Similarly, the

D4cpv‑pBAD vector was digested with EcoRI and blunted. Following heat inactivation of the blunting enzyme, the linearized vector was digested with BamH1 and the appropriate fragments (with compatible cohesive ends) were ligated and transformed into competent E. coli. Purified pUAST‑D4cpv (Promega SV Wizard miniprep) was microinjected into w1118 embryos, w+ lines were recovered, and inserts mapped to chromosomes.

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5.3.2 C2‑GFP transgenics.

We cloned the C2‑GFP construct into the pUASt (w+) vector NotI and XbaI sites using the following primers:

(F) 5ʹ‑ ATAATAATGCGGCCGCCAAAACATGGTGAGCAAGGGC ‑3’

(R) 5ʹ‑ CTGCGTCTAGATCACACAGGAACGTTAAAGTATTCTCCCTCCTCC ‑3ʹ

W1118 (w‑) embryos were injected with the plasmid, w+ lines were recovered, and inserts mapped to chromosomes.

5.3.2 GFP‑tagged, full‑length RPK transgenics.

Full length ripped pocket (cDNA DGRC #LD07574) was amplified by PCR and TA cloned into the pCR8/GW/TOPO TA vector (Invitrogen). Primers used:

RPK‑F = 5ʹ‑ ACC ATG ACC ATA TCG GAT TCG GAA CTC GAC AGC ‑3ʹ

RPK‑R =5ʹ‑ TCC TTT AAC CAG GCG CTT CAG ATT GGT AAA GAG C ‑3’

Full length RPK in the pCR8/GW/TOPO TA vector was recombined into the PTWG destination vector (Drosophila Gateway Vector Collection). Purified PTWG‑RPK

(Promega SV Wizard miniprep) was microinjected into w1118 embryos. W+ lines were recovered, and inserts mapped to chromosomes.

5.3.3 Rpk excision screen: crosses.

P0 cross: w;;rpkEY12268 adult male flies (w+) were crossed to w;;Δ2‑3, Sb/TM6 (w+) virgin females.

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F1 generation: Mosaic eyed, Sb‑ male progeny (w;; rpkEY12268/Δ2‑3, Sb) were isolated.

Single males were crossed to virgins from balancer stocks (w;Xa/Cyo;TM3Sb, w;;TM6/TM3Sb, or w;;Ly/TM3Sb).

F2 generation: (w‑, Tb‑) or (w‑, CyO‑, Sb+) single males were crossed to virgin balancer stocks to establish independent excision lines.

F3 generation and beyond: In the first two attempts at this screen, all homozygous lethal w‑ stocks were kept. In the third iteration of this screen, all w‑ stocks, both homozygous lethal and homozygous viable, were kept for analysis.

5.3.4 Rpk excision screen: PCR.

We performed a high‑throughput PCR screen for the 67 fly stocks isolated in the third screen attempt. Genomic DNA samples were taken from single male adults of each stock. A fragment spanning both the P‑element insertion site and the 5’ end of RPK was amplified for each sample, as well as a control fragment on chromosome 2. Samples that did not yield PCR product were set aside for further analysis. Primers for the initial screen: rpkscreen_F01: 5’‑ GGG ATA ACG TGT TAA CAA TG ‑3’ rpkscreen_R01: 5’‑ GCC AGA TGG ACA GGA TCT TG ‑3’

Additional primer pairs for analyzing disruptions in the rpk coding region:

RPK_excision5: 5’‑ GCA AGT TCT CTC CAA GAT CC ‑3’

RPK_excision6a: 5’‑ CGA ATC CGA TAT GGT CAT GG ‑3’

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5.3.5 Analysis of the rpk53 allele.

Genomic DNA was isolated from homozygous male escapers of the genotype rpk53/rpk53

(Qiagen DNeasy). Genomic DNA from rpkEY12268 and w1118 stocks were used as controls.

The following primers were used to amplify the site of excision:

F: 5’‑ CGT TTT CGT GCC AGA TTC TC ‑3’

R: 5’‑ CGA ATC CGA TAT GGT CAT GG ‑3’

Sanger sequencing (Applied Biosytems 3730 xl) using the following primers to determine rpk53 lesion:

RPK53‑seq1: 5’‑ CGC ACA AAC ACA GCT TCA AG ‑3’

RPK53‑seq2: 5’‑ CGC TAA ATT GTC ACC ATC CG ‑3’

5.3.6 Southern blot analysis.

Genomic DNA was prepared from adult flies (DNeasy kit, Qiagen). Digests were performed using BstBI. We used a protocol and reagents from Roche: PCR DIG probe synthesis kit, DIG Easy Hyb, Wash and Block buffer set, positively‑charged nylon membranes, anti‑DIG AP Fab fragments (1:5000), and CDP Star (1:100). The following primers were used to generate the DIG‑labeled probe:

DIG‑F = 5ʹ‑ CGT TTT CGT GCC AGA TTC TC ‑3ʹ

DIG‑R =5ʹ‑ CGA ATC CGA TAT GGT CAT GG ‑3’

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5.4 Confocal microscopy and laser microsurgery.

Embryos were prepared as above, without desiccation. Coverslips were pressed against a transparent teflon membrane for imaging (Kiehart et al., 1994). Fluorescent images were obtained using either a Zeiss Axiovert 200M or Zeiss LSM510 confocal, with Metamorph or LSM v4.0 acquisition software, respectively. Objectives used: 25x/0.8

NA multi‑immersion, 63x/1.4 NA and 100x/1.45 NA oil immersion. For general closure analysis, a stack of 5 – 11 Z‑sections taken 1 ‑ 2 μm apart were acquired and Z‑ projections were analyzed. For analysis of apical cell oscillations, Z‑sections of 0.5 μm slice spacing were acquired but not projected.

Mechanical jump experiments (Hutson et al., 2003) were performed on a Zeiss

Axio Imager.M2m, with a 40X/1.2 NA water objective, using Simple PCI or

Micromanager acquisition software. An Nd:YAG UV laser (Continuum, Santa Clara,

CA) was interfaced with the confocal microscope to generate steered laser incisions. The same system was used to uncage NP‑EGTA AM. Laser power ranging from 1 ‑ 2 μJ was used for incisions, and power ranging from 0.7 ‑ 1 μJ was used for uncaging.

5.4.1 Image analysis.

Images were analyzed using an active contour algorithm (snakes, Kass et al.,

1988) in ImageJ (NIH). Images were snaked and custom algorithms were applied to determine recoil velocity or to quantify seam formation (Mathematica, Wolfram

Research Inc., Champaign, IL; Peralta et al., 2007). Apical oscillations and actomyosin

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networks were quantified in 5 mM GsMTx4 injected (5 embryos, total of 44 amnioserosa cells) and control injected (4 embryos, total of 32 amnioserosa cells) sGMCA embryos.

Determination of the total number of oscillating cells (control vs. 5 mM GsMTx4 injected) was based on a field of view using the 63x/1.4 NA objective, approximately 30 cells. A student’s t‑test was used to determine significance.

5.5 Immunoblotting and Immunofluorescence

A polyclonal antibody was raised against RPK using the peptide

YDRAERELLVREFKRV and affinity purified (Abgent, California). For western blotting, equal numbers of dechorionated embryos or adult male flies were ground in SDS‑PAGE buffer. Antibodies and dilutions for blotting were anti‑RPK antibody (1:2000, from 0.36 mg/mL stock), anti‑dTRPA1 antibody (1:2000, AbCam), and anti‑β actin (1:5000, Sigma), with HRP‑conjugated anti‑rabbit secondary antibodies (1:3000, BioRad) and an ECL detection kit (Pierce). Immunofluorescence was performed as described (Sullivan et al,

2000). Briefly, embryos were fixed in 1:1 heptane and 4% paraformaldehyde in 1X PBS.

For antibodies, we used a methanol‑fixation method. We used anti‑RPK (1:1000), anti‑

GFP (1:3000, BD bioscience), anti‑phosphotyrosine (1:2000, Upstate), and anti‑β gal

(1:2000) primaries. Alexafluor 488 goat anti‑mouse and 568 goat anti‑rabbit secondaries

(1:3000) were used. For phalloidin staining, embryos were fixed as above and treated with Oregon green phalloidin 488 (1:1000, Invitrogen). For phalloidin staining we used an ethanol‑fixation method.

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5.6 Reagents

We wish to acknowledge the following people and institutions for providing fly stocks and constructs: Vivek Jayaraman (UAS‑GCaMP3), Amy Palmer (D2,3,4 cpv), William

Bement (C2‑GFP, C2‑RFP), Fen‑Bao Gao (UAS‑ppk RNAi), Mariana Wolfner (rpk53 allele), Paul Garrity (dtrpA1ins allele), and Yuh Nung Jan (UAS‑nompC constructs and antibody). RNAi stocks were obtained from the Vienna Drosophila RNAi Center, NIG

Fly Stock Center (Japan), and the TRiP Facility at Harvard Medical School. The following people and labs provided technical support for the work presented here:

Janice Crawford (cloning full‑length GFP RPK, C2‑GFP, and D4cpv), Julian Genkins

(RNAi image analysis), Emily Bates (Rpk excision screen), the A. Alspaugh lab

(Southern blot protocols), and the M. Noor lab (PCR screening). Funded by GM33860 to

D.P.K and HD040372‑09 Training Program in Developmental Biology to G.L.H.

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Appendix A

A.1 rpk53 allele sequence analysis.

5’‑ end of the P‑element insertion: ripped pocket ORF (5’) indicated in lower case, genomic

DNA in uppercase italics, and P‑element in uppercase. Sequence is (‑) strand as the orientation of the P‑element is 5’ to 3’ (‑). Rpk ORF is 5’ to 3’, (+). Approximately 8.9 kB of the P‑element was excised (of the initial 10.9 kB full length P‑element), with approximately 2 kB of P‑element remaining. This was verified by southern blotting.

5’‑ agacaaatggcaaaacgagaattttttcccgtttcttggcagacctgccagattgacaggatcttggagagaacttgcaaaaaaa cgttcggatCATGATGAAATAACATAAGGTGGTCCCGTCGATAGCCGAAGCTTACCG

AAGTATACACTTAAATTCAGTGCACGTTTGCTTGTTGAGAGGAAAGGTTGTGTGC

GGACGAATTTTTTTTTGAAAACATTAACCCTTACGTGGAATAAAAAAAAATGAA

ATATTGCAAATTTTGCTGCAAAGCTGTGACTGGAGTAAAATTAATTCACGTGCCG

AAGTGTGCTATTAAGAGAAAATTGTGGGAGCAGAGCCTTGGGTGCAGCCTTGGT

GAAAACTCCCAAATTTGTGATACCCACTTTAATGATTCGCAGTGGAAGGCTGCA

CCTGCAAAAGGTCAGACATTTAAAAGGAGGCGACTCACGCAGATGCCGTACCT

AGTAAAGTGATAGAGCCTGAACCAGAAAAGATAAAAGAAGGCTATACCAGTGG

GAGTACACAAACAGAGTAAGTTTGAATAGTAAAAAAAATCATTTATGTAAACA

ATAACGTGACTGTGCGTTAGGTCCTGTTCATTGTTTAATGAAAATAAGAGCTTGA

GGGAAAAAATTCGTACTTTGGAGTACGAAATGCGTCGTTTAGAGCAGCAGCCGA

141

ATTAATTCTAGTTCCAGTGAAATCCAAGCATTTTCTAAATTAAATGTATTCTTATT

ATTATAGTTGTTATTTTTGATATATATAAACAACACTATTATGCCCACCATTTTTTT

GAGATGCATCTACACAAGGAACAAACACTGGATGTCACTTTCAGTTCAAATTGT

AACGCTAATCACTCCGAACAGGTCACAAAAAATTACCTTAAAAAGTCATAATAT

TAAATTAGAATAAATATAGCTGTGAGGGAAATATATACAAATATATTGGAGCAA

ATAAATTGTACATACAAATATTTATTACTAATTTCTATTGAGACGAAATGAACC

ACTCGGAACCATTTGAGCGAACCGAATCGCG‑3’

3’‑end of P‑element insertion. Genomic DNA in uppercase bold, and P‑element in uppercase. Gap indicating partial P‑element excision denoted (‑‑‑‑‑).

5’ ‑

TGCGAGCACCCGGAAGCTCACGATGAGAATGGCCAGACCCACGTAGTCCAGCG

GCAGATCGGCGGCGGAGAAGTTAAGCGTCTCCAGGATGACCTTGCCCGAACTG

GGGCACGTGGTGTTCGACGATGTGCAGCTAATTTCGCCCGGCTCCACGTCCGCCC

ATTGGTTAATCAGCAGACCCTCGTTGGCGTAACGGAACCATGAGAGGTACGACA

ACCATTTGAGGTATACTGGCACCGAGCCCGAGTTCAAGAAGAAGCCGCCAAAG

AGCAGGAATGGTATGATAACCGGCGGACCCACAGACAGCGCCATCGAGGTCGA

GGAGCTGGCGCAGGATATTAGATATCCGAAGGACGTTGACACATTGGCCACCA

GAGTGACCAGCGCCAGGCAGTTGAAGAAGTGCAGCACTCCGGCCCGCAGTCCG

ATCATCGGATAGGCAATCGCCGTGAAGACCAGTGGCACT‑‑‑‑‑CAGACTCAATAC

142

GACACTCAGAATACTATTCCTTTCACTCGCACTTATTGCAAGCATACGTTAAGTG

GATGTCTCTTGCCGACGGGACCACCTTATGTTATTTCATCATGgttcggatGGTGACA

ATTTAGCGATATATTTTACTTCCGAATTAAATTAAATTTTGTTGAAATTAACAAAAAC

GAAAAACAGGTACATATTTTGCTGCAACTTTATTAATTTGTCATTGTTAACACGTTAT

CCCAATTCATTATTTGCACATTCGCCTGTGTTGTTAAA ‑3’

The underlined gDNA sequence is a duplication of 8 bp 5’ upstream of rpk ORF.

Bases in red are those that are inconsistent between BLAST sequence and our sequence

(insertions or misreads). Remaining bases in this second sequence (italicized) is identified as gDNA between dip2 and rpk. This deletion does not disrupt the sequence of rpk or the adjacent gene dip2.

Figure 27. Southern blot analysis of the rpk53 allele. Genomic DNA was extracted from W1118, rpkEY12268 (P‑element insertion line), or rpk53 (homozygous escapers) adult males.

143

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Biography

Ginger Hunter (b. May 25, 1982 in Fairfax, Virginia) received a Bachelor of

Science degree in Biology from the University of Virginia (Charlottesville, VA) in 2004.

She participated in the Post‑baccalaureate Intramural Research Training Award program at the National Institutes of Health, and performed research at NIDDK from

2004 to 2006. Since matriculating at Duke, she has been awarded a Sigma Xi grant‑in‑aid of research, a Department of Biology grant‑in‑aid of research, and a conference travel award by the National Institutes of Health. She completed the certificate in

Developmental and Stem Cell Biology (DSCB program) and was a fellow in the

Preparing Future Faculty program.

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