# Spatiotemporal dissection of collective cell migration and tissue morphogenesis during development by optogenetics

Sijia Zhou, Bing Liu, Jiaying Liu, Bin Yi, Xiaobo Wang

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Sijia Zhou, Bing Liu, Jiaying Liu, Bin Yi, Xiaobo Wang. Spatiotemporal dissection of collective cell migration and tissue morphogenesis during development by optogenetics. Seminars in Cell and Developmental Biology, 2025, 166, pp.36-51. 10.1016/j.semcdb.2024.12.004. hal-05534650

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Submitted on 3 Mar 2026

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Review

# Spatiotemporal dissection of collective cell migration and tissue morphogenesis during development by optogenetics

Sijia Zhou<sup>a,b,*†</sup>, Bing Liu<sup>b,**†</sup>, Jiaying Liu<sup>b</sup>, Bin Yi<sup>a,**</sup>, Xiaobo Wang<sup>b,**‡</sup>

<figure><img src="image_3.png" /></figure>

ARTICLE INFO

Keywords:

Collective cell migration

Tissue morphogenesis

Development

Genetic dilemma

Optogenetics

Biochemical and biomechanical signals

ABSTRACT

Collective cell migration and tissue morphogenesis play a variety of important roles in the development of many species. Tissue morphogenesis often generates mechanical forces that alter cell shapes and arrangements, resembling collective cell migration-like behaviors. Genetic methods have been widely used to study collective cell migration and its like behavior, advancing our understanding of these processes during development. However, a growing body of research shows that collective cell migration during development is not a simple behavior but is often combined with other cellular and tissue processes. In addition, different surrounding environments can also influence migrating cells, further complicating collective cell migration during development. Due to the complexity of developmental processes and tissues, traditional genetic approaches often encounter challenges and limitations. Thus, some methods with spatiotemporal control become urgent in dissecting collective cell migration and tissue morphogenesis during development. Optogenetics is a method that combines optics and genetics, providing a perfect strategy for spatiotemporally controlling corresponding protein activity in subcellular, cellular or tissue levels. In this review, we introduce the basic mechanisms underlying different optogenetic tools. Then, we demonstrate how optogenetic methods have been applied *in vivo* to dissect collective cell migration and tissue morphogenesis during development. Additionally, we describe some promising optogenetic approaches for advancing this field. Together, this review will guide and facilitate future studies of collective cell migration *in vivo* and tissue morphogenesis by optogenetics.

1. Introduction

Cell migration is critically important in physiological processes including embryo development and homeostasis, and pathological processes such as inflammation and tumour metastasis [1,2]. Aberration of cell migration can cause a variety of problems including birth defects, immune problems, and cancer metastasis [1,2]. In general, cell migration experiences three major stages: at the beginning, migratory cells are selected by appropriate environmental signals or intrinsic signals [3]; then they undergo a dramatic reorganization of cell adhesion and cytoskeleton network to gain their migration ability; and finally these cells move under a guidance signal which is either chemotactic (attractant or repellent: chemotaxis [4]) or mechanic (durotaxis [5]).

Cell migration is generally classified into two principal groups, single and collective cell migration [6]. The well-known cells that migrate

individually are primordial germ cells, leukocytes, and hematopoietic stem cells. Most cells derived from epithelium usually obtain their migratory ability by the epithelial-to-mesenchymal transition (EMT), a process during which epithelial cells acquire mesenchymal phenotypes and behavior following the downregulation of epithelial features [7]. During an EMT process, epithelial cells typically reduce or lose epithelial cell-cell junctions and apical-basal polarity, while enhancing interactions with basement membrane [7]. This well-studied transition explains many biological processes such as embryo gastrulation, neural crest development, metastasis, and tissue fibrosis [8]. However, in many other biological processes, including most embryo morphogenesis, wound healing, angiogenesis, and invasion of certain types of carcinoma, migrating cells maintain their cell-cell junctions and apical-basal polarity, and move as groups, sheets or strands [6,9,10]. This type of cell movement is termed collective cell migration. Compared to single cell

<footnote><sup>a</sup> Department of Anesthesiology, Southwest Hospital, Third Military Medical University, Chongqing, China</footnote>

<footnote><sup>b</sup> Molecular, Cellular and Developmental Biology Department (MCD), Centre de Biologie Integrative (CBI), University of Toulouse, CNRS, UPS, Toulouse, France</footnote>

<footnote>* Corresponding author at: Department of Anesthesiology, Southwest Hospital, Third Military Medical University, Chongqing, China.</footnote>

<footnote>** Corresponding authors.</footnote>

<footnote>E-mail addresses: sijia.zhou@univ-tlse3.fr (S. Zhou), bing.liu@univ-tlse3.fr (B. Liu), yibin1974@163.com (B. Yi), xiaobo.wang@univ-tlse3.fr (X. Wang).</footnote>

<footnote n="1">¹ These authors contributed equally to this work</footnote>
migration, the study of which is greatly benefited from the *in vitro* culture of migratory cells such as leukocytes and fibroblasts, collective cell migration is much less understood and thus draws increasing attention in development and cancer research.

Collective cell migration shares many fundamental requirements with individual one, such as cell extension regulated by actomyosin cytoskeleton, cell adhesion formed dynamically, and cell polarization controlled by either secreted guidance signal or spontaneous break of the symmetry [4,8]. However, there are some major differences between cells that migrate collectively and cells that move as an individual. The first hallmark of collective cell migration is that migrating cells usually maintain apical-basolateral polarity and intercellular adhesion to form a multicellular polarity or super-cellular organization of the cytoskeleton, which makes their whole group mechanically engaged [11–13].

The second specificity of collective cell migration is that cells need to regulate their behavior cooperatively in response to guidance signals, either chemical or mechanical signals. This raises an interesting question unique to collective cell movement: the specification of the “leader” cells vs. the “followers” cells, whose positions are normally in the front or at the back, respectively, of migratory groups [14–17]. Because each cell senses the guidance signal locally, a cell-cell contact-dependent communication mechanism is required to achieve such coordination: the contact inhibition of locomotion and contact following of locomotion are both involved in this coordination [18–20]; JNK, Cadherin adhesion components, and the ERM family protein (moesin and merlin) have been reported to play a role in intercellular communication [21–23].

In addition, the type of extracellular environment is also important in the control of collective cell migration. Most studies have been focused on matrix environments. Many *in vitro* cultured cell models have been used in either 2D or 3D matrix environments to study different types of collective cell migration, such as MDCK wound healing, astrocyte migration, mammary gland branching development and cancer-associated fibroblasts-induced collective cancer cell invasion [24–27]. For the cell-on-matrix collective migration *in vitro*, the front-rear polarization of collective cells is spontaneously formed when migrating cells interact with matrix environment, via the distinct sensing and interaction of leader vs. follower cells with chemical and mechanical cues from matrix [14]. Normally, chemotactic factors (attractant or repellent) provide the directional chemical guidance for collective chemotaxis, while matrix physical properties (stiffness, geometry and topography) also influence directional guidance, group polarization and migratory modes of a cell group [28–31]. Compared with collective cell migration *in vitro*, collective cell migration during development and pathological processes often occurs in the cell-rich tissues that are composed of not only matrix but also other types of cells (called substrate cells), thus providing much more complex chemical and mechanical cues and more combinations.

## 2. *In vivo* research models of collective cell migration

As mentioned above, collective cell migration during different developmental processes shows several major characteristics: mechanical coupling within a whole migratory group, migration direction determined by guidance signals, migration behavior influenced by surrounding environment. However, there are a lot of variations and specificities for these three characteristics. To understand all these characteristics, variation and specificities, some traditional *in vivo* models have been used to study collective cell migration during development (Fig. 1). Here, we will first highlight several traditional models demonstrating that collective cells actively migrate within tissues during development. Then, we will describe alternative models suggesting that collective cell migration-like behaviors are not a mode of active migration, but rather a change in cell shapes induced by mechanical forces of tissue morphogenesis.

### 2.1. Models for active collective cell migration

Here, we will describe several models of active collective cell migration based on species from lower to higher order in the evolutionary tree.

#### 2.1.1. *Drosophila* tracheal branching morphogenesis

The insect tracheal system is a network of epithelial tubules that act as respiratory organs, providing oxygen to numerous target organs. In the absence of cell division, tracheal branching occurs in *Drosophila* through collective movement and cell intercalation. Clusters of tracheal cells separate from epidermal cells and invaginate to form a sac-like tracheal structure (Fig. 1A') [32]. They respond to the ligand branchless (Bnl) secreted in adjacent tissues through the receptor breathless (Btl), migrate towards the source of fibroblast growth factor (FGF) and remain attached to their tracheal adjacencies. The tip cells at the end of the branches consistently have the highest activity of Btl signaling. They produce large protrusions and lead the group to migrate, while the rest are seen as followers making only small, faint protrusions (Fig. 1A, A') [33]. With the movement of tip cells, stalk cells form passive intercalation, thus contributing to the elongation of the branches [34].

#### 2.1.2. *Drosophila* border cell migration

*Drosophila* border cell migration is particularly adapted to study the collective migration within cell-rich tissue, thanks to the feasibility of genetic modifications (mutants, conditional transgene expression, mosaic analysis), and of *ex vivo* live imaging during the whole migratory period (3–6 hours in total) [35]. The basic unit of *Drosophila* ovary is egg chamber, each of which is composed of one oocyte and fifteen nurse cells surrounded by a monolayer of about 650 epithelial cells. When egg chamber develops to early stage 9, a pair of specialized epithelial cells called polar cells recruit 4–8 nearby epithelial cells to form a migratory cluster [36]. Then, they detach from their neighboring cells and basement membrane, migrate about 150 microns through the nurse cell junctions, and reach the oocyte at stage 10 (Fig. 1B, B') [36]. Finally, border cells form the micropyle, a structure for the entry of sperm. Border cell migration is a type of collective chemotaxis, driven by different growth factors (EGF and PVF1) produced and secreted from the oocyte [37,38].

#### 2.1.3. Zebrafish posterior lateral line primordium (pLLP) migration

The lateral line in fish and amphibians is a mechanosensory system. It can detect changes in water flow and is necessary for feeding and swimming [39]. Due to its proximity to the skin surface, this is a readily tractable system to explore the process of collective cell migration *in vivo*. The posterior lateral line primordium (pLLP) in zebrafish consists of approximately 100 cells that travel down the flanks of the embryo throughout development (Fig. 1C'). At about 22 hours after fertilization (hpf), the distant part of the pLLP begins to move along one side of the body, while the part including the sensory neurons stays behind (Fig. 1C, C'). Migration along the body is directed to the tail and will reach 48 hpf. During the pLLP migration, some clusters of around 20–30 cells at the tail edge will differentiate into mechanosensory neuromasts (NMs). pLLP migration is complete when the terminal cluster (i.e., a group of two or three NMs in a faraway area) is put down [40,41].

#### 2.1.4. Embryo neural crest migration

The neural crest is a multipotent embryonic stem cell population that emerges during early vertebrate development. During neurulation, cells at the border of the neural plate are induced to stratify and undergo an EMT, losing their apical-basal polarity and undergoing a switch in the expression of adhesion proteins, thereby acquiring migratory properties [42,43]. After delaminating from the neuroepithelium, neural crest cells can migrate long distances along the anterior-posterior (A-P) axis of the embryo and, upon completing their migration, differentiate in a region-specific manner to form various organs (Fig. 1D, D'). The neural
<figure><img src="image_4.png" /><figcaption>**Fig. 1.** *In vivo* research models of collective cell migration. A) and A') Schematic representation of *Drosophila* tracheal branching morphogenesis. In A, the overall branching pattern of the tracheal system in a developing embryo is shown, with red dashed circles indicating regions of active cell migration and green arrows highlighting the direction of cell movement during branching morphogenesis. In A', a close-up view of individual tracheal cells is provided, showing the arrangement of these cells within the tubular structure. B) and B') Schematic representation of *Drosophila* border cell migration. In B, the migration of border cells is illustrated within an egg chamber. The green arrow indicates the direction of their migration through the nurse cell junctions. In B', a close-up view of the border cell cluster is provided, showing the arrangement and coordinated migration of these cells as they migrate. C) and C') Schematic representation of Zebrafish posterior lateral line primordium migration. In C, an overview of the migrating posterior lateral line primordium is shown in a developing embryo. The green arrow indicates the direction of its migration along the body axis. In C', a close-up view of the posterior lateral line primordium is provided, showing the organization of individual cells as they migrate. D) and D') Schematic representation of neural crest migration. In D, the neural crest cells are shown migrating from the dorsal region of the embryo. The green arrow indicates the direction of their migration during neurulation. In D', a close-up view of the neural crest cells is provided, showing their initial organization and subsequent transition into migratory cells as they delaminate and move to their destinations during embryogenesis. E) and E') Schematic representation of *Drosophila* embryo epithelial folding. In E, two stages of epithelial invagination are depicted. The green arrow indicates the direction of inward movement of the ventral epithelial cells (highlighted in red), demonstrating the folding process that leads to tissue invagination. In E', a close-up view of the ventral epithelial cells is provided, showing their arrangement and the apical constriction that drives the invagination process. F) and F') Schematic representation of *Drosophila* egg chamber elongation. In F, model of tissue elongation along the anterior-posterior (A-P) axis. The green arrow indicates the direction of forces. In F', a close-up view of the follicular epithelial cells is provided, showing F-actin stress fibers at the follicular cell basal domain in red.</figcaption></figure>
crest, which significantly contributes to the diversity of biological tissues and organs, serves as a classic model of collective cell migration *in vivo*, with most related studies utilizing animal models such as *Xenopus laevis*, zebrafish, and chick embryos [44–46].

## 2.2. Collective cell migration-like behaviors

During tissue morphogenesis, some cells appear to undergo collective cell migration behaviors. However, unlike the above-mentioned models, these cells do not exhibit active migration on their surrounding matrix or substrate cells, whereas these seemingly migratory behaviors are due to some changes in cell shape caused by mechanical forces of tissue morphogenesis. Here, we will describe two examples of collective cell migration-like behaviors during tissue morphogenesis.

### 2.2.1. *Drosophila embryo epithelial folding*

Epithelial folding is a critical morphogenetic process during early embryonic development, involving large-scale reorganization, migration, and arrangement of cell populations [47]. During early *Drosophila* development, several key epithelial folding events occur, including the formation of the ventral furrow, cephalic furrow, and dorsal furrow [48]. The most extensively researched one is the formation of ventral furrow. The ventral furrow formation is the initial step in *Drosophila* gastrulation. In this process, approximately 1000 ventral epithelial cells undergo apical constriction and invagination, completing within about 15 minutes (Fig. 1E. E') [49]. This rapid process is essential for the internalization of the mesoderm. Following ventral furrow internalization, mesodermal precursor cells undergo EMT and migrate dorsally [50, 51].

### 2.2.2. Molecular corset for *Drosophila* egg chamber elongation

During *Drosophila* oogenesis, the egg chamber gradually changes its shape from round to elongated by extending along the anterior-posterior (A-P) axis from stages 5–14 (Fig. 1F), driven by a 'molecular corset' [52, 53]. This molecular corset is composed of contractile F-actin stress fibers at the follicular cell basal domain and of a fibrillary extracellular matrix. Both networks are organized into arrays parallel to the dorsal-ventral (D-V) axis. The mechanical function of the molecular corset is to constrain the D-V growth of the underlying germ cells, leading to epithelial extension along the A-P axis [52,53]. The molecular corset is mainly governed by oscillating basal actomyosin networks in the follicular cells surrounding the oocyte during stages 9–10 (Fig. 1F').

# 3. Genetic dilemma for studying collective cell migration *in vivo*

Collective cell migration *in vitro* can be considered as a simple migratory process of collective cells on the extracellular matrix (ECM). However, collective cell migration *in vivo* typically occurs during specific developmental periods and within specific tissues of the animal body. For example, *Drosophila* border cell migration occurs within the egg chamber during stage 9 of oogenesis [36], and *Xenopus* neural crest migration begins with the later stages of gastrulation, starting at approximately stage 15 of embryogenesis [54]. Moreover, collectively migratory border cells and neural crest cells are situated within a cell-rich tissue or on the tissue surface, and thus they are often affected by and communicate with their surrounding cells that typically exhibit other developmental processes. Currently, genetic tools have been routinely used to study collective cell migration and they uncovered important aspects of collective cell migration [34,36,40,44,50]: the transcriptional factors determining the migratory cell fate, chemo-attractant and their receptors guiding the migratory direction, cell-cell adhesions controlling the migratory integrity and coordination, cell-matrix or cell-cell adhesions mediating the transient interaction with surrounding environments. But due to the temporal and spatial characteristics, collective cell migration during development typically encounters two complexities compared to *in vitro* migration. Firstly,

collective cell migration *in vivo* is often integrated with and interacts with other cellular and tissue processes. Secondly, collective cell migration *in vivo* often occurs in the cell-rich tissues including both matrix and substrate cells.

Regarding the first complexity, several classical models of collective cell migration during development show different types of combinations of collective cell migration with other cellular and tissue processes. Many collectively migrating cells *in vivo* demonstrate the combination of EMT and collective cell migration, such as during migratory process of *Drosophila* border cells and embryo neural crests [36,42]. These migratory cells first undergo EMT, delaminating from their original location, and then begin collective cell migration within or on the tissue [36,42]. Although many genes have been identified as critical for collective cell migration, some key factors that are also important for EMT cannot be easily and well-studied for their roles in subsequent collective cell migration. Here, genetic tools have strong limitations in precise temporal control of key factors during EMT and subsequent collective cell migration. A well-known example is Rac1, whose loss-of-function mutants and expression of both active and dominant negative forms can block the detachment of border cells from epithelial monolayer, so how Rac1 controls border cell migration between nurse cells within the tissue has long been unclear [23,55]. Another example is the transcriptional factor Slbo that determines the migratory fate of border cells [56]. Strong genetic mutants of Slbo prevent the formation of border cells, so the precise role of Slbo during border cell migration process has not been clearly understood [23,56]. Recently, Li et al. unraveled a novel epithelial cell expansion behavior that combines ruffling phase and subsequent collective movement phase of follicle epithelial cells during *Drosophila* late oogenesis [57]. Li et al. found that the cell ruffling process can initiate collective cell movement on the ECM. Thus, genetic inhibition of Rac1, a key factor in cell ruffle formation, also significantly blocked our study of collective cell movement since epithelial cells cannot enter ruffling phase [57]. Furthermore, collective cell migration processes are also coupled with the intercalation of follower cells (during *Drosophila* tracheal branching morphogenesis [34]), cell differentiation (during Zebrafish pLLP migration [58]), and mechanics forces from other cell types (during *Drosophila* embryo epithelial folding [59]). Genetic studies of these *in vivo* collective cell migration models also hint at the complicated but important crosstalk between collective cell migration and these different cellular and tissue processes. This complexity thus leads to spatial limitations of genetic tools. Considering *Drosophila* embryo epithelial folding, many genetic perturbations in early embryos affect all cells within a tissue, but not certain specific cell pools, thereby complicating the interpretation of both intrinsic and extrinsic forces driving cell shape changes [60–64]. Some genetic methods have been applied to achieve temporal control (such as temperature sensitive Gal4 system in *Drosophila* [57], and hormone dependent gene expression system in *Xenopus* [65]) and spatial control (mosaic system [66] used to randomly express gene in many animal models), but these tools lack precision in temporal and spatial control of gene expression.

These genetic limitations are also reflected in the second complexity, that is, collective cell migration often occurs in the cell-rich tissues which are composed of matrix and other types of cells. Current studies of collective cell migration during development mostly lack information from these cell-rich tissues. Normally these other types of cells on which migrating cells move are named stroma cells (also called substrate cells) [67]. The characteristics and role of multicellular environment (substrate cell environment) in collective cell migration *in vivo* are poorly understood, compared with the ones of matrix environment. Border cell migration is a perfect model for studying collective migration within cell-rich tissue [36]. During the past 30 years, many important signaling pathways and factors (such as JAK-STAT, Ecdysone, chemo-attractants...) have been identified by traditional genetic studies, almost completely from border cells, to differently control border cell migration: migratory cell fate, time window, direction etc [36]. Although the
nurse cell environment is also thought to play an important role in border cell migration, little is known about the important factors of nurse cells that control border cell migration. This lack of information is due to the genetic dilemma (that is, the lack of precise spatial and temporal control), potentially affecting other *in vivo* models of cell-on-cell collective migration, which prevents the studies of multi-cellular environments. Since the genetic inhibition of important factors in the nurse cell environment (such as the Rho1-myosin signaling pathway that controls cortical tension in nurse cells, as well as Lamin B that controls nurse cell division process, both from our unpublished data) often blocks the ovarian germline development and progression to the stages when border cells form and migrate, it is impossible for genetic methods to study the role of nurse cells in the control of border cell migration. In addition to cell-on-cell migration, other types of cells in contact with migrating collective cells seem to also influence collective cell migration through their effects on the ECM. For example, mesoderm stiffening, which comes from convergent extension of the mesoderm and cell density increase, is necessary and sufficient to trigger neural crest migration [68]. How these neighboring cells influence the substrate environment to control collective cell migration during development is currently poorly explored, as these studies may encounter a similar genetic dilemma as border cell migration.

In addition to these two complexities, genetic tools are also limited in their ability to control the subcellular activities of factors and signals of interest. Indeed, during collective cell migration, many biochemical signals and biomechanical forces occur at specific subcellular regions. An example is small GTPase Rac1 or RhoA, the key signal mainly controlling lamellipodia protrusions or actomyosin contractility, respectively. It is known that either GTPase shows different subcellular regions in different models of collective cell migration *in vivo*, such as migrating border cells and neural crests [23,69,70], implicating that the same factors and signals might exert different biological roles. Thus, developing a technique with more precise spatiotemporal control is urgent to overcome these genetic and subcellular limitations. Optogenetics has recently emerged as a useful strategy for spatiotemporally dissecting collective cell migration both *in vitro* and *in vivo*.

## 4. Optogenetics and its working mechanisms

As an emerging biological technology, optogenetics is less than 50 years, and it consists of genetic and optical technologies that induce or inhibit the activity of proteins in live tissues [71]. Light is utilized to spatiotemporally manipulate protein activity of interest using optogenetics [71]. The rise of optogenetics with great spatiotemporal resolution started in 2005, when a single component optogenetic instrument was first used in neuroscience [72]. Following its success in neuroscience, optogenetics is now being applied to a wide range of other biological concerns including biochemical signals and biomechanics.

Well-defined light-sensitive domains for optogenetic regulation of biochemical signals include cryptochrome proteins, phytochrome proteins, light-oxygen-voltage domains (LOV) and fluorescent protein Dronpa. 1) Cryptochrome 2 (CRY2) protein isolated from *Arabidopsis thaliana* can be activated by blue light (excitation wavelength around 405–488 nm). When this occurs, CRY2 can homo-oligomerize and recruit its binding partner, Cryptochrome interacting basic helix-loop-helix 1 (CIB1) [73]. Within minutes, CRY2 can transition back to its original condition during the dark phase. Endogenous flavin ubiquitously produced across species serves as the chromophore for CRY2 in this system. 2) Phytochrome B (PHYB) can be triggered by red light (maximal excitation wavelength at 650 nm) and deactivated by far-red light (700 nm) (maximal excitation wavelength at 750 nm). Initially, the chromophore-free apo-PHYB protein is not light sensitive, but upon binding to the chromophore Phycocyanobilin (PCB), it becomes light sensitive. PCB exits in photosynthetic organisms, and through autocalysis, it can maintain the function of PHYB [74,75]. The conformation of PHYB-PCB molecule can be altered by red light excitation, and the

compound then attaches to the Phytochrome interacting factor (PIF) protein [76]. In response to far-red stimulation, the association process is reversible. 3) The LOV domains found in a variety of species are sensitive to blue light with the excitation wavelength range of 440–473 nm, and they also utilize flavin as a chromophore. It is known that LOV domains undergo the light-induced conformational change that abolishes auto-inhibition and allows activation [77]. In the absence of blue light irradiation, the LOV domain conformation rapidly resets to its original state. 4) Dronpa is a monomeric fluorescent protein that is sensitive to light with a maximal excitation wavelength at 390 nm. This monomer can be dimerized by light treatment. In this way, Dronpa can limit the activity of the target protein through its own C-terminal aggregation. On the other hand, Dronpa can be converted back to a monomer by light with maximal excitation wavelength at 490 nm [78].

Using a variety of light-sensitive proteins and controllers, optogenetics allows for the manipulation of intracellular signals in five distinct ways (Fig. 2) [79]. 1) Controlling the target by protein association (Fig. 2A). Light can stimulate the interaction of the two proteins and subsequently modulate intracellular signaling, since the light-sensitive protein and the effector heterodimerize upon exposure to light. This heterodimerization leads to the recruitment or repulsion of proteins, either activating or inhibiting intracellular signals, respectively. These light-inducible proteins are mainly composed of the CRY2–CIBN, PHYB–PIF, and LOV domains. 2) Clustering-based activation (Fig. 2B). Activation of a signaling cascade begins with the oligomerization of signaling proteins in response to light. This form of activation usually requires significant quantities of signaling proteins in the local area. Cryptochrome-based regulation of β-catenin and Rho in charge of transcription and cytoskeletal rearrangements, respectively, is an illustration of this activation [80]. 3) Conformational changes within a processed protein (Fig. 2C). This method has been applied to the LOV domains to control Rac1, Cdc42, and formin in the regulation of cell movements [81], and used with Dronpa to control Cdc42 and proteinase K. It shares the principle of converting an autoinhibited signaling protein into an active state by light treatment, and it is highly efficient and reversible [78]. However, since it relies so much on the conformational match between the light-sensitive domain and the target protein, fewer tools have successfully been developed with this technique than with other techniques. 4) Inhibition based on sequestration (Fig. 2D). A cluster trap is employed to remove proteins from their operational location [82]. For instance, the light activated reversible inhibition by assembled trap (LARIAT) system relies on the heterodimerization of CIB1 and CRY2. It consists of a multimeric protein (MP) that is attached to CIB1 and CRY2. The MP domain is a component of calmodulin-dependent protein kinase II that self-assembles into an oligomer composed of 12 identical subunits [83]. Upon blue light treatment, CRY2 proteins oligomerize and attach to CIB1, and then the cross-linked MPs form into clusters. This method has been used to stop multiple regulators of the actin cytoskeleton and as a general way to control GFP-tagged proteins with a GFP-nanobody [84]. 5) Regulating gene expression through heterodimerization (Fig. 2E). Light-mediated heterodimerization of PHYB–PIF[85] or LOV-domains [86,87] has been used to control the target protein. By targeting a transcriptional activator to a gene’s promoter, this activation mechanism can control gene transcription.

## 5. Optogenetic studies of collective cell migration *in vivo*

Collective cell migration is a complex process that plays a critical role during development and is orchestrated by multiple cellular and molecular players. Small GTPases, such as Rac1, Cdc42, and RhoA, are key regulators of cytoskeletal dynamics, driving cell polarization, protrusion, and contractility through downstream effectors. Cell adhesion molecules, including cadherins and integrins, mediate intercellular and cell-matrix interactions, ensuring coordinated movement of the cell group. Nuclear factors, such as transcription regulators and nuclear
<figure><img src="image_5.png" /><figcaption>B Clustering-based activation</figcaption></figure>

<figure><img src="image_6.png" /><figcaption>C Conformational changes within a processed protein</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>D Inhibition based on sequestration</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>E Regulating gene expression through heterodimerization</figcaption></figure>

<figure><img src="image_9.png" /></figure>

**Fig. 2.** Five distinct ways for optogenetic manipulation. A) Upon light activation, the light-sensitive protein (red) binds to its binding partner (blue). This heterodimerization leads to the recruitment of signaling molecule (green), resulting in the recruitment of a signaling domain to the substrate located on the membrane. B) Upon light activation, CRY2 (red) naturally clusters, bringing together attached signaling domains (green). This clustering increases the local density of the signaling domains, which can trigger downstream signaling pathways that are dependent on domain density. C) Upon light activation, the photosensitive protein (red) undergoes a conformational change, exposing a previously hidden signaling domain or relieving allosteric autoinhibition. This conformational shift allows interaction with signaling domains (green). D) Upon light activation, signaling is inhibited by sequestering the photosensitive protein (red) and signaling domains (green) in a cluster (blue), separating the signaling domains from downstream effectors or upstream activators. E) Upon light activation, the photosensitive protein (red) undergoes homodimerization, bringing together attached transcriptional activators or other DNA-modifying proteins (green) and binding to the DNA (blue/green strands), thereby initiating the expression of the gene of interest. The curved blue arrow indicates light-induced changes, while the straight black arrow represents dark-reversed changes.
structural proteins, modulate gene expression to facilitate cellular responses to environmental cues. Optogenetics, since its introduction in 2010, has provided a powerful tool to manipulate these components with light, enabling precise spatiotemporal control over cytoskeletal regulators, cell interactions, cytosolic proteins, and nuclear factors [23]. Over the past 14 years, this technology has advanced our understanding of how these key players regulate collective cell migration during development (Table 1). In this section, we will review current applications of the optogenetic tools in vivo, in order from the outside to the inside of the cell.

## 5.1. Optogenetic manipulation at the cortex

Among all the methods, the use of optogenetics to control Rho-family small GTPases is the most widely applied and well-established approach for studying collective cell migration and tissue morphogenesis in vivo. The Rho-family GTPases are key regulators of actin organization, dynamics, and myosin activity at the cell cortex. The core Rho-GTPases, including Rac1, Cdc42, and RhoA, serve as central hubs in signaling circuits, integrating extracellular and intracellular signals while regulating the activity of various effector proteins. Under the enzymatic activity of guanine nucleotide exchange factors (GEFs) and GTPase-activating proteins (GAPs), Rho-GTPases continuously switch between an inactive GDP-bound state and an active GTP-bound state [88]. In their active state, Rho-GTPases play distinct roles in cell behavior: Rac1 and Cdc42 mainly control the extension of lamellipodia and filopodia, respectively, in migrating cells, while RhoA mainly governs the formation of actomyosin stress fibers and the retraction of the rear of motile cells [89]. Additionally, optogenetic tools targeting actin regulators, such as phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) and Arpin, have also been developed to modulate cortical actin dynamics, complementing the study of optogenetic manipulation at the cortex.

<table><caption>**Table 1**
Already Applied Tools.</caption><thead><tr><th>Optogenetic Tool</th><th>Light-Sensitive Domain</th><th>Target or Effector Protein</th><th>In Vivo Collective Migration Model</th><th>Ref.</th></tr></thead><tbody><tr><td>PA-Rac1</td><td>LOV2</td><td>Rac1</td><td>Border cell migration, Follicle cell expansion (Drosophila)</td><td>[21,23,57,70]</td></tr><tr><td>PA-Cdc42</td><td>LOV2</td><td>Cdc42</td><td>Border cell migration (Drosophila)</td><td>[70]</td></tr><tr><td>OptoGEF-RhoA</td><td>CRY2</td><td>ARHGEF11 (DHPH)</td><td>Neural crest migration (Xenopus)</td><td>[16,95]</td></tr><tr><td>Opto-RhoGEF2</td><td>CRY2</td><td>RhoGEF2 (DHPH)</td><td>Embryo epithelial folding, Border cell migration, Follicle cell expansion (Drosophila)</td><td>[57,70,97,98,104,105]</td></tr><tr><td>Opto-Rho1DN</td><td>CRY2</td><td>Rho1DN</td><td>Embryo epithelial folding, Border cell migration, Follicle cell expansion (Drosophila)</td><td>[57,70,101–105]</td></tr><tr><td>LARIAT</td><td>CRY2</td><td>E-cadherin, Integrin, ROCK, Flapwing, aPKC, Myosin II, α-tubulin</td><td>Ovarian epithelial cells, Border cell migration, Pupal notum extrusion (Drosophila)</td><td>[109–112]</td></tr><tr><td>LEXY/iLEXY</td><td>AsLOV2</td><td>Twist</td><td>Embryo development (Drosophila)</td><td>[119]</td></tr></tbody></table>

### 5.1.1. PA-Rac1 and PA-Cdc42

The first optogenetic tool developed and employed to control collective cell migration in vivo is the photoactivatable analog of Rac (PA-Rac1), which is based on the LOV2 domain. In this system, Rac1 active or dominant negative mutant is fused to the LOV domain from phototropin, which sterically blocks the interaction of Rac1 with its downstream effectors. Upon light irradiation, the helix connecting LOV to Rac1 unwinds, allowing Rac1 to become active [77]. This tool was first created in 2009 for controlling cell migration in vitro, and just a year later, Wang et al. expanded its application to border cell migration to study the spatiotemporal role of Rac1 in controlling collective cell migration during Drosophila oogenesis [23].

Wang et al. first generated UAS-PA-Rac1 transgenic flies and then used the GAL4/UAS system to specifically express PA-Rac1 in migrating border cells [23]. When expressed in border cells, PA-Rac1 is distributed throughout the cell, including the cytoplasm, nucleus, and cell surface. Pulsed laser exposure activates Rac (PA-RacQ61L), inducing lamellipodia formation and redirecting cell movement, even the opposite direction to the intended path [23]. Conversely, activation of dominant-negative Rac (Rac-T17N) strongly inhibits border cell movement [23]. Furthermore, activating Rac1 in the leading edge promotes group polarization by inhibiting protrusions at the rear of border cell group [23]. Conversely, inhibiting Rac1 in the leading cell causes other cells to protrude in all directions, as if the guidance receptor activity is lost [23]. Combining these results with Rac FRET experiments, Wang et al. concluded that asymmetric Rac1 activity is crucial for directional sensing of collective cells in vivo. Additionally, inhibition of guidance receptor activity by expressing dominant negative forms of PDGF/VEGF receptor (PVR) and epidermal growth factor receptor (EGFR) abrogates the Rac1 asymmetry in border cell group, which can be rescued by the photo-activation of PA-Rac1 in one border cell [23]. But in this study, how a border cell group integrates directionality, with coordination remained unclear. In 2014, Montell's team further confirmed the asymmetric distribution of Rac1 activity in border cells, with a front-to-back gradient regulated by cell adhesions [21]. However, both studies had the same limitation: original design of the Rac FRET probe missed the CAAX motif, which is a membrane-targeting signal at the C terminus of most GTPases. Thus, distribution detection of Rac1 activity in border cells is inaccurate [21,23].

Lately, this issue was addressed by using another Rac probe, PAK3RBD, which can exclude intracellular distribution of Rac1 but focus on Rac1 activity directly controlling the cytoskeleton at the cell periphery [90]. Zhou et al. showed no significant gradient in Rac1 activity from front to rear of border cell group, while implicating Rac1 activity at different subcellular regions (supracellular cables vs. protrusions) [70]. Combining more precise optogenetic control of PA-Rac1 with important cytoskeleton reporters of F-actin and non-muscle myosin II (myosin II), Zhou et al. surprisingly revealed that Rac1 exists in two interconvertible pools: the protrusive Rac1 pool that controls cell protrusions, and the tensile Rac1 pool that regulates cell cables [70]. Optogenetic activation of Rac1 in either pool increases Rac1 activity in that pool while decreasing Rac1 activity in the other (optogenetic inhibition of Rac1 in either pool produces the opposite effects), thereby maintaining an overall balance of Rac1 activity in one border cell [70]. Notably, activating Rac1 in the protrusive pool leads to forward-directed cell protrusions, promoting directional migration of border cells, while activating Rac1 in the tensile pool strengthens supracellular cables, enhancing cell coordination [70]. Therefore, maintaining a dynamic balance between these two Rac1 pools is essential for effective collective cell migration, as it successfully integrates cell directionality with coordination. Optogenetic control of PA-Rac1 was also used to confirm that the guidance receptors PVR and EGFR can control the protrusion and tensile Rac1 activities, respectively, thereby guiding border cell migration efficiency [70]. This study perfectly demonstrates that modulation of protein activity distribution with subcellular precision during collective cell migration in vivo can only be achieved through
optogenetics [70].

To explore how these two pools exchange signals, another opto-
genetic tool, PA-Cdc42, was applied to photo-activate or photo-inhibit
Cdc42 activity in border cells. Similar to PA-Rac1, this tool involves
fusing a Cdc42 active or dominant negative mutant to the LOV domain
[91]. Wang team first introduced this optogenetic tool in another
*Drosophila* *in vivo* system to study how intracellular filopodia serve as
bridges linking basal stress fibers between two adjacent follicle epithe-
lial cells [91]. In migrating border cells, using this tool, Zhou et al. found
that photo-activated Cdc42 significantly enhances retrograde flows,
which represent communication between the two Rac1 pools, and in-
creases border cell migration speed by 1.6-fold [70]. In contrast,
photo-inhibition of Cdc42 blocks retrograde flows and halts border cell
movement [70]. Furthermore, combining the two optogenetic tools,
PA-Rac1 and PA-Cdc42, further confirmed these findings. When Cdc42
activity is inhibited, activating or inhibiting Rac in one pool has no effect
on the other pool [70]. Conversely, when Cdc42 activity is activated, the
effects of Rac activation or inhibition are amplified [70]. These results
indicate that Cdc42 is a key factor in controlling signal communication
and exchange between the protrusive and tensile Rac1 pools, and that
the cooperation between Cdc42 and Rac1 achieves higher migration
efficiency [70]. This discovery challenged the previous theory that
suggests a purely competitive relationship between Cdc42 and Rac1
[92]. Whether these two small GTPases compete or cooperate may
depend on their spatial distribution, which cannot be detected by
traditional genetic and pharmacological approaches.

Our ongoing studies utilized the PA-Rac1 tool to continue exploring
how the spatial distribution of Rac1 within multicellular tissues in-
fluences border cell migration. We lately found that photo-activating
Rac1 in surrounding nurse cells along the migration path significantly
increases the tension of these nurse cells, effectively opening the sur-
rounding environment and facilitating border cell migration (from our
unpublished data). Conversely, photo-inhibition of Rac1 activity in the
same region leads to reduced tension and thus closure of the environ-
ment, completely blocking border cell movement (from our unpublished
data). Interestingly, we discovered a threshold below which environ-
mental cellular tension positively affects cell migration. When tension
exceeds this threshold, the formation of micro-bubble structures will be
induced, which can also impair border cell movement (from our un-
published data). These unpublished findings suggest that tension in
surrounding environmental cells plays a crucial role in regulating the
efficiency of collective migration within multicellular tissues.

In addition to being widely used in studying border cell migration, PA-Rac1 has recently been employed in research investigating collective migration of follicle cells that contributes to tissue elongation *in vivo*. In another study of Wang team, Li et al. sought to elucidate the mechanisms that control tissue extension along the A-P axis during *Drosophila* late oogenesis [57]. Using traditional genetic approaches, this study achieved temporal inhibition of Rac1 activity, resulting in an acute suppression of cell ruffling and a blockade of collective cell spreading process [57]. By employing optogenetic manipulation of PA-Rac1, follicle cell ruffling process was precisely modulated in both activating and inhibiting manner [57]. This approach further confirmed the critical role of Rac1 in regulating cell ruffle formation, which subsequently triggers collective migration and tissue elongation [57].

Currently, the direct application of PA-Rac1 and PA-Cdc42 in other
classic collective migration models *in vivo* has not yet been reported.
However, a recent study utilized optogenetics to precisely control the
growth direction of motor neuron axons in zebrafish, highlighting the
great potential of PA-Rac1 and PA-Cdc42 in other systems [93].

5.1.2. *Opto-RhoGEF2 and Opto-Rho1DN*

Different from Cdc42 and Rac1, RhoA/Rho1 regulates Rock and thus
myosin II to control cellular contractility; additionally, RhoA/Rho1 can
activate mDia protein, which contributes to actin filament nucleation
and elongation. Together, these two pathways facilitate cytoskeletal

reorganization. RhoA/Rho1, like Cdc42 and Rac1, is post-translationally
modified by the addition of a lipid moiety to its C-terminal CAAX motif,
enabling it to signal from the plasma membrane [94]. This is the design
basis for RhoA/Rho1 optogenetic regulation.

In 2017, Valon et al. developed OptoGEF-RhoA, an optogenetic tool based on the CRY2/CIBN light-induced dimerization system [95]. They selected the DHPH domain of ARHGEF11 (one of the RhoGEFs) [96] and fused it with CRY2-mCherry to create ARHGEF11 (DHPH)-CRY2-mCherry. They also constructed CIBN-GFP-CAAX to anchor the CIBN chaperone protein to the plasma membrane [95]. The interaction between CBIN and CRY2 under blue light illumination can activate endogenous RhoA by directly recruiting the RhoA catalytic domain of GEF to the plasma membrane. The advantage of over-expressing a RhoA activator, rather than RhoA itself, lies in the fact that changes in contractility are driven by endogenous RhoA levels, allowing the system to maintain its physiological tension homeostasis. Using pulsed blue light, they were able to increase traction forces and membrane tension by 50 % while promoting the formation of actin stress fibers [95]. Conversely, they designed an optogenetic tool that inhibits RhoA activity by recruiting RhoA to the mitochondrial membrane (mito-CIBN-GFP) under light exposure [95]. Optogenetic inhibition of RhoA results in a 4-fold reduction in cellular traction forces and tension, with the disappearance of basal stress fibers [95]. Both the effects of RhoA photo-activation and photo-inhibition are fully reversible. More importantly, they achieved optogenetic induction of epithelial cell deformation; by activating RhoA in the central region of MDCK cell monolayer, they successfully directed the cells to move toward the illuminated area [95]. This success demonstrates the potential for future applications of optogenetic control of RhoA activity in collective cell migration *in vivo*.

Indeed, two 2018 studies from the De Renzis lab introduced this
optogenetic strategy to investigate ventral furrow formation in
*Drosophila* embryos, a good model for studying collective cell migration
during development [97,98]. The De Renzis lab employed
Opto-RhoGEF2 to activate Rho1 signaling on the apical surface of
developing *Drosophila* embryos [97]. This approach allows the
light-mediated recruitment of the DHPH domain of cytoplasmic Rho-
GEF2 to the plasma membrane within 30 sec, leading to sustained and
specific accumulation of myosin II at the apical surface without inducing
apoptosis [97]. By tracking changes in apical cell surface area over time,
they confirmed that recruitment of RhoGEF2 to the membrane is suffi-
cient to trigger apical constriction [97]. Furthermore, by utilizing pre-
cise spatiotemporal regulation enabled by optogenetics, they induced
Rho1 activation at different stages of embryonic development. This
consistently leads to embryo invagination, forming a U-shaped furrow,
demonstrating that Rho1 activation is sufficient to induce tissue invag-
ination at specific times [97]. Spatially, by altering the shape of the light
exposure, they successfully reversed the normal anisotropic contraction
along the D-V axis [99], confirming that the anisotropy of ventral furrow
cells is related to tissue geometry and mechanical constraints rather than
the overall shape of the embryo. In addition, they utilized pulsed light to
study the periodicity of apical surface contraction and relaxation; this
study successfully replicated the endogenous mechanochemical oscil-
lation system [100] and revealed the correlation between pulsatile
contraction and tissue invagination [97].

Although apical constriction can induce invagination, it is not suf-
ficient to facilitate a folding of internalized tissue into a tubular struc-
ture. In a subsequent study, the De Renzis lab quantitatively controlled
myosin II levels at the basal surface of invaginating cells through
optogenetic stimulation of Rho1 signaling, achieving a 1.6-fold increase
in myosin II within 5 minutes and maintaining this increase at pre-
invagination levels [98]. By activating Rho1 signaling at the basal sur-
face during different stages of epithelial folding, they found that
increasing basal myosin II levels before invagination inhibits cell elon-
gation and preserves columnar shape, interfering with apical constric-
tion and tissue invagination, whereas increasing basal myosin II levels
after cell elongation and initial tissue invagination inhibits cell short-
ening and the folding of the ventral furrow into a tubular structure [98].
These results indicate that a specific ratio of apical to basal myosin II is
crucial for inducing the correct changes in cell shape, which may drive
the complete invagination process [98]. Thus, these two optogenetic
studies support that both apical constriction and basal relaxation are
necessary for successful tissue folding.

Two years later, the He group developed the Opto-Rho1DN tool for
optogenetic inhibition of Rho1 activity [101,102]. This tool is also based
on the CRY2-CIBN system. The OptoRho1-DN tool directly fuses the
dominant-negative form of Rho1 (Rho1DN) with CRY2 to create a fusion
protein [102]. Upon blue light illumination, Rho1DN is recruited to the
plasma membrane, where it binds to and sequesters the Rho1 GEF,
thereby preventing endogenous Rho1 activation and subsequent myosin
II activation [103]. Using this tool, Eritano et al. investigated the effects
of perturbing myosin II contractility before cephalic furrow formation
during *Drosophila* embryogenesis [101]. Their results demonstrates that
optogenetic inhibition of myosin II contractility prior to cell internali-
zation prevents fissure formation, confirming their hypothesis that ce-
phalic furrow initiation depends on myosin II contractility [101].
Subsequently, by utilizing spatial Rho1 photo-inhibition in the center of
the cephalic furrow, they observed that a reduction in contractility
within the illuminated cell region causes the contraction band to split
into two [101]. This mechanical decoupling of the two lateral furrows
leads to decreased spatial alignment and impedes further tissue folding.
These findings indicate that tissue-scale mechanical coupling is critical
to ensure the linearity and spatial precision of fold morphology [101]. A
few years later, Guo et al. also used this tool to photo-inhibit Rho1 ac-
tivity before or during apical surface contraction in *Drosophila* embryos,
and successfully prevented and reversed apical contraction [102].
However, they observed that photo-inhibition of Rho1 activity during
late-stage of contraction does not affect ventral furrow invagination,
suggesting that myosin II contractility is required for early but not
late-stage of furrow formation [102]. They further examined the impact
of early and contraction-phase apical inhibition of Rho1 activity on
ectodermal movement and found that ectoderm continues to move
ventrally even without ventral furrow formation [102]. This decoupling
of tissue movement implies that ectodermal shortening may act as an
independent mechanical input, acting in concert with other mechanical
factors to promote mesoderm invagination [102].

The Rauzi lab simultaneously applied two optogenetic techniques, Opto-RhoGEF2 and Opto-Rho1DN, in their studies of epithelial folding in *Drosophila* embryos [104,105]. In their studies of ventral furrow formation, they observed planar polarization and cell intercalation behaviors during apical constriction of mesoderm cells [104]. To explore the molecular mechanisms behind these behaviors, they first employed light-activated apical Rho1 and outer myosin II activity to induce cell apical constriction, and their results ruled out the possibility that mesodermal furrow itself drives cell intercalation [104]. However, by activating medial Rho1 activity and thus the second layer of myosin II, they successfully mediated junctional contraction, indicating that the second layer of myosin II controls planar polarized cell intercalation [104]. This result has been confirmed by the presence of lateral tension within the mesodermal cells during folding, revealed by their laser ablation experiment [104]. Furthermore, they utilized Opto-Rho1DN to downregulate lateral myosin II contractility, thereby inhibiting mesodermal tissue internalization; and conversely, they used Opto-RhoGEF2 to upregulate lateral myosin II contractility, thereby enhancing both tension and localized tissue bending [104]. These findings demonstrate that myosin II-driven lateral tension facilitates mesodermal tissue bending. This study revealed for the first time the synergistic role of the A-P and D-V gene patterns in controlling embryonic development [104]. The precision in temporal, spatial, and protein-specific control afforded by the OptoRho optogenetic tools is crucial to the success of this research. On this basis, they then used OptoRho technology to conduct in-depth research on the formation of cephalic furrow in early *Drosophila*

embryos and deeply analyzed the role of Rho signaling in cephalic
furrow formation [105]. This study uncovered the driving forces, me-
chanical mechanisms, and developmental roles involved in cephalic
furrow formation [105].

In addition to embryonic epithelial folding, OptoRho technology has
also been applied to study embryo neural crest migration. Aiming to
understand the role of rear cells in collective cell migration, the Mayor
lab first utilized laser ablation to demonstrate that ablation of actomy-
osin rings in rear cells impairs chemotaxis, whereas equivalent ablation
in front cells does not, suggesting that actomyosin cables in the rear cells
are crucial for chemotaxis [16]. To specifically test the contractility of
these supracellular actomyosin rings in defined regions, they employed
two OptoGEF-RhoA based optogenetic systems [95], OptoRho-contract
and OptoRho-relax. They found that that photo-inhibition of rear cell
contractility using OptoRho-relax disrupts collective migration regard-
less of the presence or absence of the chemotactic factor SDF1 [16,69],
whereas photo-inhibition of front cell contractility has no effect. In
contrast, photo-activation of front cell contractility using
OptoRho-contract inhibits collective cell chemotaxis, while
photo-inhibition of front cell contractility induces directional migration
even in the absence of SDF1 [16]. These findings suggest that rear cell
contractility generated by a supracellular actomyosin cable can drive
collective cell chemotaxis *in vivo* [16].

More recently, OptoRho technology was utilized in our investigation of supracellular actomyosin cable within border cells. By photo-activating or photo-inhibiting Rho1 activity in border cell cable, Zhou et al. observed a corresponding increase or decrease in myosin II accumulation, which is also accompanied by respective changes in contractile Rac1 activity [70]. Chemical inhibition of Rock activity [106] significantly reduces Rac1 activity in border cell cables, and localized Rho1 activation fails to upregulate Rac1 activity under this ROCK inhibition condition, suggesting that Rho1, through the Rock/myosin II signaling pathway, cooperates with Rac1 to maintain mechanical coupling between border cells for intercellular communication [70]. Furthermore, OptoRho technology was also applied to investigate the local and global effects of Rho1 signaling on follicle cell expansion behaviors. By photo-activating or photo-inhibiting Rho1 activity at basal domains of multiple follicle cells, Li et al. found that global activation or inhibition of Rho1 signaling significantly enhances or inhibits follicle cell collective spreading over ECM [57]. However, by photo-activating Rho1 activity at one follicle cell basal domain, this local photo-activation has no prominent effect on collective spreading [57]. These results thus confirm the spatiotemporal role of Rho1, but also highlight that basal pulsatile actomyosin networks present in multiple follicle cells are essential for collective spreading process [57].

In summary, as an acute tool for manipulating myosin II contrac-
tility, the OptoRho technology enables researchers to distinguish be-
tween “active” forces and “passive mechanical properties” in
morphogenesis and collective cell migration without altering cell fate,
exploring the autonomy and contributions of local regions within the
whole tissue, making it a powerful tool for studying cell mechanical
properties during development. This spatiotemporal dissection of
cellular and tissue mechanical roles cannot be achieved by traditional
genetic manipulation of migrating cells and their surrounding
environments.

5.1.3. *Optogenetics targeting actin regulation*

To study the apical constriction for *Drosophila* ventral furrow for-
mation, the De Renzis group utilized the CRY2/CIB1 optogenetic
dimerization system to control the subcellular localization of the cata-
lytic domain of inositol polyphosphate 5-phosphatase OCRL, creating
the CRY2-OCRL tool [107]. By depleting plasma membrane PI (4,5) P2,
a critical regulator of cortical actin dynamics, they successfully modu-
lated cell contractility during tissue morphogenesis [107]. When
expressed in *Drosophila* embryos, this tool effectively disrupted ventral
furrow formation by inhibiting apical cell contraction. They also applied
light at varying scales and regions within the ventral furrow, demonstrating that as long as a sufficient number of cells remain free to contract, these cells can locally coordinate and generate enough tension to drive invagination [107]. And recently, the Gonzalez-Gaitan group developed another optogenetic tool by co-expressing Lifeact-CRY2FL-mCherry and CIB1-GFP-Arpin in sensory organ precursor cells [108]. This system targets Arpin to cortical actin, inhibiting F-actin nucleation and weakening the actin network, providing a potential approach to studying collective cell migration *in vivo*.

## 5.2. Optogenetics targeting GFP tagged factors by LARIAT

Compared with optogenetics approaches targeting special factors, LARIAT (Light-activated reversible inhibition by assembled trap) offers broader applicability as it can directly target GFP-tagged proteins and perform cluster traps within the cytoplasm. LARIAT is an optogenetic tool developed by Lee et al. in 2014 [84]. LARIAT is a versatile inactivation method, in which light-mediated CRY2 heterodimerization with CIB1 mediated fused to MP robustly sequesters and inhibits target proteins [84]. To achieve broader applications, Lee et al. introduced an anti-GFP nanobody with CRY2, which can allow LARIAT to photo-inhibit any GFP-tagged proteins in cultured cells [84]. LARIAT creates a light-inducible GFP trap and thus disturbs the localization and function of GFP-tagged protein [84]. GFP-LARIAT succeeds in the photo-inhibition of small GTPase (Rac1, RhoG, Cdc42), RhoGEF (Vav2), and RacGEF (Tiam1) [84] in cultured cancer cells. Photo-inhibition of Rac1 and Vav2 blocks the ability of cells to migrate, whereas photo-inhibition of tubulin tagged with YFP strongly represses the cell division process [84].

A few years later, to understand the mechanism of basal myosin oscillation in *Drosophila* egg chamber elongation (molecular corset), Wang team introduced GFP-LARIAT in *Drosophila* *in vivo* system and applied GFP-LARIAT to control various GFP-tagged proteins in *Drosophila* ovarian epithelial cells [109,110]. Two studies of Qin et al. demonstrate the precise spatiotemporal roles of myosin II and cell-matrix/cell-cell adhesions (E-cadherin, integrin), ROCK and phosphatase (Flapwing), in controlling the basal pulsatile actomyosin contractility for *Drosophila* egg chamber elongation [109,110]. Clustering-trap of these GFP-tagged proteins rapidly decreases basal myosin II oscillation in follicle cells within around 30 min [109,110]; these optogenetic effects resemble the phenotypes from the genetic limitation of these targets, suggesting that basal myosin II oscillation can be controlled by short-term manipulation of its upstream controlling factor [109,110]. Moreover, Osswald et al. applied GFP-LARIAT to spatiotemporally dissect the precise role of apical polarity factor (aPKC) in *Drosophila* ovarian epithelial cells for tissue homeostasis [111]. They found that in *Drosophila* proliferative follicular epithelium, fast optogenetic inactivation of aPKC results in increased apical constriction rather than loss of apical-basal polarity, generating larger epithelial gaps [111]. This study shows that GFP-LARIAT can also be applied to dissect short-term inhibitory effects of apical-basolateral polarity proteins, which are previously inaccessible using genetic tools.

In addition to the studies of biomechanics and tissue morphogenesis, GFP-LARIAT has also been applied to study collective cell migration *in vivo*. In our unpublished work, we applied GFP-LARIAT to photo-inhibit myosin II-GFP in migrating border cells or surrounding environmental nurse cells, to test the spatiotemporal role of actomyosin contractility in border cell migration. We found that photo-inhibition of myosin II-GFP by LARIAT in either migrating or environmental cells can strongly block border cell migration (from our unpublished data), indicating that actomyosin contractility is critical for each cell type for collective cell migration within multicellular environments. By localized photo-inhibition, we also dissected the precise role of actomyosin contractility at different localizations of migrating or environmental cells, and these studies suggest that actomyosin contractility at front cells and rear cells (of both migrating and environmental cells) can play different roles

in controlling border cell migration (from our unpublished data). Our success in photo-inhibition of E-cadherin-GFP and integrin-GFP [110] seems to support future applications of this tool to dissect the spatiotemporal role of either adhesion in collective cell migration *in vivo*. In addition to our study, the Levayer lab employed GFP-LARIAT to induce the aggregation of GFP-tagged α-tubulin under blue light during collective cell extrusion *in vivo* [112]. Microtubes are well known for their stabilizing function of cell membrane protrusions during cell migration [113]. Their study demonstrates that microtubules can independently stabilize the cell apical region, regulating its contractility during early stage of cell extrusion [112].

Regarding the current availability of GFP-knock-in proteins, GFP-LARIAT will undoubtedly provide a broad application of optogenetic control on various signals and factors in different types of collective cell migration models *in vivo*.

## 5.3. Optogenetics targeting nuclear factors by LEXY

Similar to the cytosolic signaling cascades, nuclear signals are very dynamic, and their spatiotemporal controls play essential functions in physiological and pathological processes including embryo development, tumorigenesis, and cancer progression [114]. Nuclear-cytoplasmic shuttling is an essential mechanism controlling the activity of transcriptional factors [115], thereby providing the basis of optogenetic tools for nuclear factors [116]. Among them, LEXY (light-inducible nuclear export system) has been applied to control embryo development involving collective cell migration *in vivo*.

LEXY is a blue light-induced, fast, and fully reversible system that enables precise spatiotemporal control of nuclear protein export [117]. LEXY consists of an engineered LOV2 domain from *Avena sativa* phototropin-1 (AsLOV2), in which the C-terminal Jα helix has been converted into an artificial NES [117]. In the dark, the NES is tightly packed against the AsLOV2 core and thus inactive. On blue light absorption, the modified Jα helix unfolds, thereby exposing the NES to nuclear CRM1 (exportin-1) receptors, which mediates active nuclear export through the nuclear pore [117]. LEXY enables straightforward control of nuclear export of various nuclear factors, including transcription factors (p53, Sox2 and Nanog), a phosphatase (Acp1), a protein export-related factor (Nxt1), a cyclin-dependent kinase inhibitor (p21), a copper chaperone (Cox17), as well as a bacterial repressor (LexA) and a bacteriophage-derived protein (Cre recombinase), in human non-small cell lung carcinoma cells [117]. In addition, Zhao et al. established an MRTF-A-LEXY light-induced nuclear localization system in cancer cell lines [118]. Enhanced MRTF-A activity induces membrane blebbing formation to promote invasive cancer cell motility [118], suggesting that targeting MRTF-A or its downstream signaling pathways by LEXY optogenetic approach could be a useful strategy for dissecting complex cellular behaviors in cancer cell invasion and metastasis.

To adapt to the study of mesoderm development during gastrulation (collective cell migration model), Kögler et al. generated a series of mutations to increase LEXY's efficiency and produce different recovery times of LEXY variants to accommodate different time scales of biological processes of interest [119]. An improved optogenetic tool (iLEXY) enables rapid (t1/2 < 30 s), efficient, and reversible nuclear depletion in *Drosophila* embryos [119]. LEXY has mainly been applied to photo-inhibit Twist, a master regulator directing critical mesoderm cell fate decisions. In *twist* null mutant, embryos exhibit embryonic lethality, gastrulation defects, and the loss of mesoderm-derived muscle tissues [60–64]. However, it has been very difficult to functionally dissect the role of Twist after gastrulation due to early developmental defects in twist mutant embryos. iLEXY provides a unique opportunity to address these issues by allowing embryos to gastrulate before Twist is removed from the nucleus. In this study, iLEXY-mediated mislocalization of Twist results in developmental defects that phenocopy genetic loss-of-function allele, demonstrating efficient perturbation of transcriptional factor function in living embryos [119]. Then, by manipulating the presence of
Twist in the nucleus after gastrulation using iLEXY, this study investigated the developmental window in which Twist is crucial for embryonic survival and revealed that Twist is essential during gastrulation and early post-gastrulation stages (stages 8–9) [119]. Using the iLEXY system to temporally control Twist expression, this study found that 1) before and during gastrulation (stages 5–8), Twist regulates a small set of genes crucial for mesoderm migration and early development; 2) after gastrulation (stages 9–10), Twist influences a broader set of genes, particularly those involved in somatic muscle development [119]. This iLEXY optogenetic study highlights the importance of Twist in early mesoderm development and its ongoing role in regulating gene expression post-gastrulation, particularly in somatic muscle formation. Finally, this study also demonstrates that this strategy can be applied to various nuclear proteins with diverse sizes and molecular functions, including TFs (Mef2, Tinman, Biniou, Bagpipe, TfIIB) and chromatin regulators (Sin3A), in *Drosophila* and possibly many other model systems [119].

However, iLEXY system has several limitations. Although it can be applied to many nuclear proteins, it may not work effectively for proteins that are tightly anchored in the nucleus (for example H2B [117]). The presence of the mCherry tag can sometimes interfere with protein function, so it is recommended to test both tagged and untagged iLEXY variants [119]. Additionally, the effectiveness of blue light activation of iLEXY depends on the nuclear retention of proteins, so the intensity and duration of light should be optimized [119]. Finally, iLEXY displays slight leakiness in the dark, and even in the absence of blue light, a small amount of the protein is exported from the nucleus, so the dark control should be included in experiments [119]. This is the actual situation that we encountered in border cell migration. Our recent unpublished study demonstrates that both Slbo-LEXY and Slbo-iLEXY translocate primarily into the cytoplasm even in the dark. It indicates that some nuclear proteins can be rapidly translocated into the cytoplasm via the NES/AsLOV2 motif but slowly shuttle back to the nucleus. Thus, the relative abilities of nuclear-cytoplasmic shuttling are critical for applying this optogenetic approach to study collective cell migration *in vivo*.

## 6. Potential optogenetic tools for collective cell migration *in vivo*

In addition to Rho GTPases, cell-cell interactions, receptors, and other cytosolic proteins are also critical for collective cell migration during development [36]. Recently, optogenetic tools targeting these molecular mechanisms have been successfully developed and applied *in vitro*. Integrating these tools into the study of collective cell migration *in vivo* is expected to significantly advance the fields of developmental biology and regenerative medicine. In this section, we will review current applications of these optogenetic tools *in vitro*, in order from the outside to the inside of the cell (Table 2).

### 6.1. Cell-environment interactions and cell-cell interactions

The microenvironment surrounding collectively migrating cells primarily consists of two types: ECM and intercellular space. Cell-on-ECM migration depends on a variety of focal adhesion proteins, such as integrins, talin, vinculin, and paxillin [120,121]. To elucidate the molecular mechanisms underlying focal adhesion-mediated mechanotransduction, it is crucial to control the transmission of specific forces through supramolecular linkages in both spatial and temporal dimensions. As a result, several optogenetic tools have been developed and applied to the study of focal adhesion proteins.

For example, the Weber lab utilized the red/far-red light-mediated PHYB/PIF system to investigate cell-matrix interactions [122]. By anchoring PhyB to the substrate and inserting a PIF6 variant into the extracellular domain of integrin αVβ3 [123], they achieved optogenetic control of cell-matrix interactions *in vitro* [122]. Their findings demonstrate that αVβ3 integrin-mediated cell-matrix interactions stabilize cell

<table><caption>**Table 2**
Potential Tools.</caption><thead><tr><th>Optogenetic Tool</th><th>Light-Sensitive Domain</th><th>Target or Effector Protein</th><th>Ref.</th></tr></thead><tbody><tr><td>OptoIntegrin</td><td>PHYB</td><td>αVβ3</td><td>[122]</td></tr><tr><td>iLID-nano</td><td>LOV2</td><td>Talin 1</td><td>[124]</td></tr><tr><td>Opto-focal adhesion</td><td>LOV2</td><td>CAAX, ALK3, β3-integrin, BMPRII</td><td>[126]</td></tr><tr><td>Opto-Ecadherin</td><td>LOV2</td><td>E-cadherin</td><td>[128]</td></tr><tr><td>Opto-Mfgr1</td><td>LOV</td><td>mFGFR1</td><td>[133]</td></tr><tr><td>OptoEGFR</td><td>CRY2</td><td>EGFR</td><td>[135,136]</td></tr><tr><td>2paRAF</td><td>CRY2</td><td>Raf1</td><td>[144]</td></tr><tr><td>OptoRas</td><td>CRY2</td><td>Hras, KRas4B, RasGRP4, RASAL3</td><td>[145]</td></tr><tr><td>OptoKANK</td><td>LOV2</td><td>KANK1</td><td>[149]</td></tr><tr><td>LINuS</td><td>AsLOV2</td><td>Fyn kinase, p21, Mxi1, dCas9,</td><td>[153–157]</td></tr><tr><td>LANS and LINX</td><td>ASLOV2</td><td>H2B, Bre1, Set2</td><td>[159,162]</td></tr><tr><td>OptoYAP</td><td>LOV2</td><td>YAP</td><td>[167,168]</td></tr><tr><td>ShineGal4/UAS</td><td>Magnet photoswitches</td><td>His2B, E-cadherin</td><td>[169]</td></tr><tr><td>NLS-LARIAT</td><td>CRY2</td><td>Lamin, Slbo, LSD1</td><td>Unpublished</td></tr></tbody></table>

adhesion, promote filopodia formation, and facilitate cell spreading on the matrix [122]. The Yan lab employed an iLID system in which two talin segments have been fused with LOV2-SsrA and SspB, respectively [124]. Photo-activation of the iLID system induces dimerization and reassembly of full-length talin 1, thereby promoting cell polarization and migration [124]. Similarly, the Albiges-Rizo lab developed a set of optogenetic tools using the iLID system, including Opto-CAAX, Opto-ALK3, Opto-BMPRII (fused with tagRFP-SspB), and Opto-β3 integrin (fused with iLID). These tools were employed to study the spatiotemporal regulation of BMPR [125] and integrin interactions. Their research revealed that the spatiotemporal segregation of ALK3 and BMPRII within integrin-based adhesions is crucial for cell adhesion and migration [126]. Altogether, these diverse studies demonstrate that optogenetic technologies can successfully elucidate key molecular mechanisms of focal adhesion proteins in cellular mechanosensing and migratory behavior. These tools for studying cell-on-ECM migration may be applicable to classic collective cell migration models, such as zebrafish pLLP migration, *Drosophila* tracheal branching morphogenesis, and embryonic neural crest migration.

Compared with cell-on-matrix migration, studies on collective cell migration within multicellular environments remain relatively underexplored. E-cadherin is a fundamental adhesion molecule that facilitates cell-cell adhesion through the interaction of its extracellular domains between neighboring cells [127]. Our ongoing research introduces a novel optogenetic tool, Opto-E-cadherin, to investigate the dynamics of cell contact and separation during border cell migration. Developed by the Wegner lab, this tool modifies the Ca²⁺ binding site of E-cadherin by inserting the LOV2 domain, allowing light-induced Ca²⁺ affinity reduction and thereby disrupting E-cadherin-mediated cell-cell adhesion [128]. They found that Opto-E-cadherin can induce cells to lose adhesion to neighboring cells, leading to increased single cell migration [128]. However, in typical collective cell migration, loss of cell adhesion often results in the disruption of intercellular communication and mechanosensing, which in turn slows migration [129]. This suggests that collective cell migration *in vivo* is governed by more complex regulatory mechanisms, requiring precise spatiotemporal control to be fully understood. Therefore, successful application of Opto-E-cadherin to study border cell migration will undoubtedly facilitate our understanding of the precise spatiotemporal control of cell-cell interactions for collective cell migration within multicellular environments. Additionally, E-cadherin has also been extensively studied in *Drosophila* epithelial folding and neural crest cell migration [130,131]. Introducing this optogenetic tool into these models could further explore the underlying mechanisms of cell-cell contact and intercellular communication within migrating cell groups with higher spatiotemporal precision.
## 6.2. Chemoattractant receptor

Cell surface receptors act as critical intermediaries between extracellular stimuli and intracellular signaling cascades, playing a pivotal role in coordinating various cellular behaviors, including cell migration, proliferation and differentiation. Among these, the receptor tyrosine kinase (RTK) family is one of the most extensively studied surface receptor families, capable of sensing growth factors and hormones to regulate chemotaxis [132]. Key members of this family include EGFR, platelet-derived growth factor receptor (PDGFR), fibroblast growth factor receptor (FGFR), and vascular endothelial growth factor receptor (VEGFR). To achieve precise regulation of RTK signaling, several research groups have developed optogenetic RTK systems based on fusion with protein domains that undergo conformational changes or oligomerization in response to light. For example, in 2014, the Janovjak lab designed Opto-mFGFR1, a chimeric receptor containing the mammalian FGF catalytic domain and the algal LOV domain, which can successfully activate the canonical MAPK/ERK pathway under blue light, inducing an EMT-like cellular morphology [133]. The Schaffer lab employed blue light-mediated oligomerization of *Arabidopsis* Cry2 protein to indirectly cluster and activate PDGFRβ by targeting the endogenous RTK with the SH2 domain of PLC-γ, demonstrating that PDGFRβ is a key mediator of fibroblast chemotaxis [134]. In addition, the Toettcher lab utilized Cry2 clustering to construct the OptoEGFR system for two optogenetic studies [135,136]. By fusing the intrinsically disordered region (IDR) of EGFR with the photolyase homology region (PHR) of Cry2, they achieved blue light-triggered clustering and autophosphorylation of the EGFR cytoplasmic domain. Their first study focused on the interaction between ECM and cells, revealing that EGFR expression and activation efficiency are regulated by ECM stiffness [135]. Their second study examined the regulation of collective cell migration *in vitro* and demonstrated that local upregulation of EGFR increases tissue fluidity and edge migration speed, thereby inducing collective migration polarity [136].

Using traditional genetic approaches, researchers have extensively investigated the role of FGF chemoattractant in pLLP migration [137], tracheal branching morphogenesis [32], and neural crest migration [138], as well as the critical regulatory role of EGFR in border cells [139]. The development and application of these novel optogenetic tools to modulate RTKs *in vitro* provide valuable insights into the *in vivo* studies of collective cell migration and offer potential optogenetic tools for studying the spatiotemporal roles of FGFR, EGFR and other RTKs *in vivo*.

## 6.3. Cytosol proteins

Cell behavior is typically the result of the interplay between external triggers and intrinsic properties. To understand how external and internal signals are integrated and coordinated during cell behavior, specific tools are needed to precisely regulate intracellular signaling pathways. Here, we will review a couple of optogenetic tools targeting some cytosol pathways and structures.

One of the most well-studied pathways is the Ras-Raf-MEK-ERK cascade, which plays a central role in cell movement, proliferation, and differentiation [140]. As a downstream pathway of RTKs, previous studies have shown that this cascade promotes single cell migration through the phosphorylation of myosin light chain (MLC) kinase and focal adhesion kinase. The introduction of optogenetics has enabled a deeper exploration of its molecular mechanisms [141,142]. In two studies, Matsuda et al. utilized the light-induced CRY2/CIBN interaction system to recruit Raf proteins to the plasma membrane, thereby activating ERK signaling [143,144]. Their first study demonstrates that ERK activation waves drive cell clusters to migrate in the opposite direction to the ERK wave, emphasizing that waves of ERK activation are key coordinators of cell migration [143]. Their second study showed that mechanical forces can mediate the transmission of ERK signals between

cells and ERK activation waves can align multicellular polarity, resulting in coordinated long-range migration [144]. These two findings, through spatiotemporal control of ERK activation waves, reveal that intercellular signal transmission relies not only on biochemical signaling but also on the cellular response to mechanical stimuli. Later, the Devreotes lab applied optogenetics to explore the impact of Ras regulation on the downstream mTORC2/Akt pathway [145]. Using the CRY2/CIBN dimerization system, they targeted Ras to the plasma membrane to activate and study mTORC2 signaling and Akt-mediated cytoskeletal effects in neutrophils [145]. They confirmed that localized activity of growth factor signaling cascades at the plasma membrane directly regulates immune cell polarity and migration [145]. Among several classic models of collective cell migration, research on ERK has mainly focused on neural crest migration [146], where it plays a crucial role in regulating the EMT behavior of these cell groups. The introduction of these aforementioned optogenetic tools will further advance our understanding of the specific mechanisms by which ERK functions during neural crest cell migration.

In many cell types, microtubules are another important cytoplasmic structure responsible for controlling cell shape polarity and directional migration behavior [147,148], which depends on their interactions with integrin-mediated cell-matrix adhesions. The Bershadsky lab developed an optogenetic construct based on the iLID system to manipulate KANK1 protein (OptoKANK) [149]. Local photo-activation of KANK1-mediated microtubule/talin connections facilitates microtubule targeting to individual focal adhesions, leading to centripetal sliding and rapid focal adhesion disassembly; and this optogenetic study elucidated the microtubule-driven focal adhesion disassembly process [149]. As a critical component regulating cell polarity, signal transduction, intercellular connections, and intracellular transport, microtubules have been widely studied in various *in vivo* models of collective cell migration [150–152]. The application of OptoKANK in these models will greatly enhance our understanding of the interactions between microtubules and focal adhesions.

## 6.4. Other NLS- or NES-mediated optogenetic tools

The major strategy of optogenetic control of nuclear factors involves the use of light to switch the nuclear entrance and exit of a specific transcriptional factor. This is how LEXY (or iLEXY) has been established to control various nuclear factors in the gastrulation of *Drosophila* embryos [119]. In addition to LEXY, here we will review some other optogenetic tools for controlling nucleocytoplasmic shuttling that have been established but not yet been applied to study collective cell migration *in vivo*.

### 6.4.1. LINus (light-inducible nuclear localization signal)

LINUS has been developed based on AsLOV2 that reversibly triggers the import of proteins of interest into the nucleus with blue light [153]. Each LINUS is a small, versatile tag fused to the protein of interest at the N or C terminus. Niopek et al. demonstrated that the efficiency of LINUS-mediated nuclear import can be tuned at different levels by introducing mutations that alter the photo-caging properties of AsLOV2 domain or by selecting NLSs of various strengths, and they applied this blue light-induced LINUS system to regulate nuclear import of several functional proteins of mitosis in mammalian cells [153]. Wehler et al. also described a detailed protocol for how to perform initial microscopy-based screening to assess which LINUS variant is best suited for the protein of interest [154]. LINUS has been applied to explore gene regulation of transcriptional factors in yeast [155], assess nuclear activity of Fyn kinase [156], and control subcellular localization of CDK-interacting protein p21 for potential application in regulating cancer cell migration and progression [157]. Conceptually, LINUS can be used to regulate the import dynamics of any protein of interest, regardless of its size or function. Different endogenous NLS and/or NES mutants are required for different nuclear import target proteins, but
fortunately, these sequences and corresponding required mutations are often known or can be predicted, making LINUS an ideal candidate for externally triggering the nuclear import of any protein of interest in collective cell migration in vivo.

### 6.4.2. LANS and LINX

The Kuhlmann lab has developed LANS and LINX, two optogenetic tools that are conceptually identical in concept to LINUS and LEXY, albeit slightly different in implementation [158,159]. LANS and LINX have been engineered by embedding NLS or NES motif into the C-terminal helix of the light-responsive AsLOV2 domain, thereby enabling target proteins to rely on light to enter and exit the nucleus of mammalian cells, yeast, and *C. elegans* [158–160]. Furthermore, Lerner et al. improved photo-switching functionality of LANS and LINX tools by generating mutations within the LOV2 hinge-loop [161]. Different from the domain structure of LEXY, LINX has a single contiguous helix connecting the two AsLOV2 domains. Fusion of LINX to the chromatin modifier Bre1 enables precise, light-controlled regulation of histone H2B monoubiquitylation in yeast [159]. Similarly, Lerner et al. established optogenetic control of Set2, the sole H3K36 methyltransferase in yeast, by fusing it to the LANS domain [162]. These optogenetic studies provided a detailed temporal view of H3K36 methylation and demethylation, indicating that H3K36me deposition and removal are transcription-dependent and transcriptional-independent machinery, respectively [162]. Given that these tools have been successfully validated in mammalian cells, yeast, and *C. elegans* models, they can be utilized to regulate nuclear factors involved in collective cell migration in vivo across a broader range of animal models.

### 6.4.3. OptoYAP

The Hippo-YAP pathway is a key signaling cascade highly relevant to development and regeneration due to its role in driving cell proliferation, migration and mechanosensing [163–165]. Central to the Hippo-YAP pathway is a transcriptional factor YAP, which is pivotal in wound healing by limiting cytoskeletal and focal adhesion maturation to promote cell motility [166]. YAP activity is determined by very dynamic nucleocytoplasmic shuttling, thereby providing the basis of optogenetic control of YAP activity [167,168]. OptoYAP is a reversible optogenetics strategy that contains a LOV2 domain to photocage NLS to the N-terminus of YAP [167]. The LOV2 domain photocages the NLS to prevent OptoYAP from importing into the nucleus in the dark; when exposed to 488 nm light, the LOV2 domain unfolds, exposing the NLS, so that OptoYAP can shuttle into the nucleus [167]. In cultured cell studies, OptoYAP photo-activation accelerates wound healing in H9c2 and MKN28 cells by enhancing collective cell migration [168]. Similarly, Opto-YAP (or OptofYap) can be used in zebrafish embryos to modulate target genes [167]. Initially, optofYap rapidly shuttles into the nuclei of both the enveloping layer and deep cells upon activation with pulsatile 488 nm light, showing a twofold increase in nuclear localization [167]. Furthermore, OptofYap can successfully induce the upregulation of target genes in vivo following prolonged activation [167]. Overall, given the importance of YAP with respect to development and regeneration, OptoYAP or OptofYAP has the potential to become a powerful tool for studying Hippo-YAP signalling in both research and clinical applications.

### 6.5. Transcriptional activity control

The Gal4/UAS system is a powerful method to modulate targeted gene expression and to induce loss or gain of function, enabling detailed studies of gene function in various biological processes. Recently, Di Pietro et al. developed a light-inducible Gal4/UAS system (ShineGal4/UAS) based on Magnet photoswitches, which can efficiently, rapidly, and robustly activate UAS-driven transgenes in different tissues and developmental stages of *Drosophila* [169]. Magnet (Mag) photoswitches comprise negatively charged nMag and positively charged pMag

proteins that heterodimerize in response to blue light. To achieve higher interaction strengths, they multimerized Magnet photoswitches [170] by fusing two tandem repeats of nMagHigh1 and pMagHigh1 to the Gal4 DNA binding domain (Gal4DBD; [171]) and the p65 transactivation domain (AD; [172]), respectively, to generate the Gal4DBD:2xnMagHigh1 and 2xpMagHigh1:AD fusion genes. nMagnet and pMagnet tandems heterodimerize upon blue light exposure and promote Gal4-mediated activation of gene expression.

Di Pietro et al. tested the ubi-Gal4MagHigh system using the UAS-His2B reporter in *Drosophila* pupal notum tissue and found that when the pupae are exposed to 491-nm blue light, His2B signals become detectable in the tissue within 2 hours and reaches a maximum expression level of 35-fold [169]. The results confirmed that ubi-Gal4MagHigh driver rapidly turns on ubiquitous gene expression throughout the living animal upon 491-nm illumination, and that the resulting protein accumulation persists for several hours after the 491-nm illumination ends [169]. In addition, the ShineGal4/UAS system can be used to locally control gene expression in a region of interest (ROI) and at a given developmental time [169]. This study also tested the versatility of the ShineGal4/UAS system to induce gene expression in various *Drosophila* tissues (including pupal wings, histoblast nests, larval wing discs, the notum, and the central nervous system) and life cycle stages, confirming its robustness and broad applicability for controlling gene expression in living *Drosophila* at different stages [169]. In addition, the ShineGal4 system has been demonstrated to induce gain- and loss-of-function phenotypes in *Drosophila* globally and locally [169].

Overall, ShineGal4 enriches the genetic toolkit of *Drosophila* by taking advantage of a large number of UAS-driven transgenes to control gene expression in vivo with high temporal and spatial resolution, enabling detailed studies of gene activity and its effects during development, including the study of collective cell migration in vivo.

### 6.6. GFP-NLS-LARIAT

Current optogenetic strategies for nuclear factors are limited to a specific transcriptional factor and thus its mediated transcriptional control. Thus, for different transcriptional factors, different optogenetic constructs will be required, while a common optogenetic tool is urgent to be developed for a broader application. More importantly, current strategies are very limited in controlling other nuclear factors and structures that are not directly related to gene transcription. Therefore, to overcome the drawback of a specific-target limitation, we recently established a common optogenetic method based on the GFP tag, called GFP-NLS-LARIAT. GFP-LARIAT is a versatile inactivation method based on the light-inducible GFP trapping strategy that interferes with the localization and function of GFP-tagged proteins [84]. However, due to the original design focus, this GFP-LARIAT tool lost NLS in the CIBN-MP partner, and it is unable to enter the nucleus to interact with nuclear Cry2-VHh(GFP) for the photo-inhibition of various nuclear GFP targets [84,110]. Thus, we recently introduced the NLS sequence into our established CIBN-MP partner, which can allow the light-mediated clustering trap of GFP-targets within the nucleus. Here, we name this novel optogenetic tool as NLS-LARIAT. In principle, the NLS-LARIAT technique could be adapted to target all endogenous GFP-tagged nuclear proteins thanks to the GFP-specific nanobody.

From our recent unpublished studies, NLS-LARIAT has robust effects on various nuclear factors, including nuclear envelope proteins (Lamin), transcription factors (Slbo) and epigenetic factors (histone demethylase LSD1), in different cell types within *Drosophila* ovary egg chambers in vivo. Especially, NLS-LARIAT mediated photo-inhibition of Slbo-GFP and LSD1-GFP phenocopied RNAi or mutant effects in border cell migration (from our unpublished data). Compared with genetics-mediated strong defect in detachment, NLS-LARIAT easily allows us to dissect the function of Slbo-GFP photo-inhibition on migratory border cells after their detachment. We also observed different nuclear transport effects after GFP clustering traps of different nuclear factors (from
our unpublished data). Now we are studying the transcriptional analysis (RNA sequencing) of border cell groups before/after clustering of Slbo-GFP or LSD1-GFP by NLS-LARIAT. NLS-LARIAT seems to provide additional and broader optogenetic choice for a variety of nuclear factors, in *Drosophila* and other model systems.

## 7. Conclusion and future perspective

Here, we summarize how different optogenetic tools have been established and applied to spatiotemporally dissect various signals, including biochemical signals and cytoskeleton-mediated biomechanics, at the plasma membrane, cytosol and nucleus to control collective cell migration during development, thereby overcoming genetic dilemmas that hinder our precise understanding of the spatiotemporal function of these signals for collective cell migration. Although many other established optogenetic tools have not yet been applied to the *in vivo* studies of collective cell migration, they have successfully modulated key proteins involved in different cellular processes and uncovered numerous molecular mechanisms that could potentially be recapitulated *in vivo*. These findings lay a solid technical and theoretical foundation to guide researchers in further applying optogenetics to study collective cell migration during development.

## Author contributions

XW: study conception and design, literature analysis, and writing. SZ: literature analysis, figure illustration, and writing. BL: literature analysis and writing. JL: figure illustration. BY: writing.

## Declaration of Competing Interest

None.

## Acknowledgments

This work is supported by Agence Nationale de la Recherche (ANR PRC AAPG2022 NEW-corset) and Scientifiques de la Fondation ARC (PJA20191209714), to XW; National Natural Science Foundation of China (82402569) to SZ; National Natural Science Foundation of China (82470655) and Natural Science Foundation of Chongqing (CSTB2023NSCQ-ZDJ0005) to BY; BL is supported by China Scholarship Council (CSC) PhD fellowship; JL is supported by ANR PRC AAPG2022 NEW-corset. We thank Karine Belguise for the discussion of manuscript preparation.

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