# Origin and evolution of microvilli

Mylan Ansel, Kaustubh Ramachandran, Gautam Dey, Thibaut Brunet

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Mylan Ansel, Kaustubh Ramachandran, Gautam Dey, Thibaut Brunet. Origin and evolution of microvilli. Biology of the Cell, 2024, 116 (11), pp.e2400054. 10.1111/boc.202400054. hal-05083682

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RESEARCH ARTICLE

Biology
of the **cell**

# Origin and evolution of microvilli

Mylan Ansel<sup>1,2,3</sup> | Kaustubh Ramachandran<sup>2</sup>

Thibaut Brunet<sup>1</sup>

Gautam Dey<sup>2</sup>

## Correspondence

Thibaut Brunet, Institut Pasteur, Université Paris-Cité, CNRS UMR3691, Evolutionary Cell Biology and Evolution of Morphogenesis Unit, 25–28 rue du docteur Roux, Paris, 75015, France.

Email: [thibaut.brunet@pasteur.fr](mailto:thibaut.brunet@pasteur.fr)

## Funding information

European Research Council; Institut Pasteur; European Molecular Biology Laboratory; École Normale Supérieure de Lyon; Centre National de la Recherche Scientifique; Joachim Herz Stiftung

## Abstract

**Background Information:** Microvilli are finger-like, straight, and stable cellular protrusions that are filled with F-actin and present a stereotypical length. They are present in a broad range of cell types across the animal tree of life and mediate several fundamental functions, including nutrient absorption, photosensation, and mechanosensation. Therefore, understanding the origin and evolution of microvilli is key to reconstructing the evolution of animal cellular form and function. Here, we review the current state of knowledge on microvilli evolution and perform a bioinformatic survey of the conservation of genes encoding microvillar proteins in animals and their unicellular relatives.

**Results:** We first present a detailed description of mammalian microvilli based on two well-studied examples, the brush border microvilli of enterocytes and the stereocilia of hair cells. We also survey the broader diversity of microvilli and discuss similarities and differences between microvilli and filopodia. Based on our bioinformatic survey coupled with carefully reconstructed molecular phylogenies, we reconstitute the order of evolutionary appearance of microvillar proteins. We document the stepwise evolutionary assembly of the “molecular microvillar toolkit” with notable bursts of innovation at two key nodes: the last common filozoan ancestor (correlated with the evolution of microvilli distinct from filopodia) and the last common choanozoan ancestor (correlated with the emergence of inter-microvillar adhesions).

**Conclusion and Significance:** We conclude with a scenario for the evolution of microvilli from filopodia-like ancestral structures in unicellular precursors of animals.

## KEYWORDS

biological evolution, Choanoflagellata, cytoskeleton, filopodia, microvilli, molecular evolution

**Abbreviations:** ABCG, ATP-binding cassette protein G; ANKS4B, Ankyrin repeat and Sterile Alpha motif domain containing 4B; BAIAP2L2, BAR/IMD Domain Containing Adaptor Protein 2 Like 2; BB, brush border; CIB, Calcium and Integrin-Binding protein; COBL, Cordon Bleu; ENA/VASP, Enabled/Vasodilator-Stimulated Phosphoprotein; ERM, Ezrin, Radixin, Moesin; EBP50, Ezrin-radixin-moesin Binding Phosphoprotein 50; ENaC, Epithelial Sodium Channel; EPS8, Epidermal Growth Factor receptor Substrate 8; FERM, 4.1 protein, Ezrin, Radixin and Moesin; GLUT, Glucose Transporter; GRCR1 or GRXCR1, Glutaredoxin and Cystein Rich Domain Containing enzyme 1; HMM, Hidden Markov Model; IMAC, Intermicrovillar Adhesion Complex; IRTKS, Insulin Receptor Tyrosine Kinase Substrate; MET, mechanoelectrical transduction; MYO, myosin; NH3, Na+/H+ exchangers 3; NPC1L1, Niemann-Pick C1 like-1; PDZD7, PDZ-Domain Containing 7; PEPT1, Peptide Transporter 1; PIP₂, phosphatidylinositol 4,5-bisphosphate; SANS, Scaffold protein containing Ankyrin repeats and SAM domain; SGLT1, Sodium/Glucose cotransporter 1; TMC, transmembrane channel; TRPM, Transient potential ion channel; TWF2A, Twinfilin-2.

This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited.

© 2024 The Author(s). *Biology of the Cell* published by Wiley-VCH GmbH on behalf of Société Française des Microscopies and Société Biologie Cellulaire de France

<footnote n="1"><sup>1</sup>Institut Pasteur, Université Paris-Cité, CNRS UMR3691, Evolutionary Cell Biology and Evolution of Morphogenesis Unit, Paris, France</footnote>

<footnote n="2"><sup>2</sup>Cell Biology and Biophysics, European Molecular Biology Laboratory, Heidelberg, Germany</footnote>

<footnote n="3"><sup>3</sup>Master BioSciences, Département de Biologie, Ecole Normale Supérieure de Lyon, Lyon, France</footnote>
# INTRODUCTION

François Jacob pointed out that “living beings are traversed by a triple flow of matter, energy and information” (Jacob, 1996). In eukaryotic cells, the detection, uptake, and response to environmental biotic and abiotic factors often relies on cellular protrusions based on cytoskeletal filaments of actin or microtubules. Microtubule-based cellular protrusions include cilia (also known as flagella [Derderian et al., 2023]; Wan, 2018), while actin-based protrusions include lamellipodia (or pseudopods), filopodia and microvilli (Mattila & Lappalainen, 2008; Pelaseyed & Bretscher, 2018a; Theofanopoulou et al., 2017). While lamellipodia and pseudopods are broad and lamellar structures filled with branched actin (Fritz-Laylin et al., 2017; Mattila & Lappalainen, 2008), both filopodia and microvilli are finger-like protrusions scaffolded by bundles of linear actin. However, as we will detail below, filopodia and microvilli differ in their structure, molecular composition, functions, and evolutionary history (Houdusse & Titus, 2021; Mattila & Lappalainen, 2008; Sebé-Pedrós et al., 2013; Sharkova et al., 2022).

In this review, we summarize the current state of knowledge on the origin and evolution of microvilli. Microvilli have not always been precisely defined in the literature and terminological inconsistencies can thus be a complicating factor in comparative analyses. Here, we propose an explicit definition of microvilli informed by current knowledge of the morphology and molecular composition of BB microvilli and stereocilia, highlighting differences with filopodia. We thereby attempt to identify a set of features that is both universal among and specific to, microvilli. We then discuss the diversity of microvilli in animals and their unicellular holozoa relatives and perform a bioinformatic analysis of the phylogenetic distribution of microvillar proteins across Amorphea (the clade that includes animals, fungi, and amoebozoans as well as all the descendants of their last common ancestor). Finally, we discuss when, how, and why microvilli evolved, and propose an evolutionary scenario for microvillar emergence and diversification.

# A STRUCTURAL DEFINITION OF MICROVILLI

## The fundamental characteristics of mammalian microvilli

Microvilli have been extensively studied in vertebrates, where they occur in multiple cell types and perform diverse functions. The earliest and best-described mammalian microvilli are those of intestinal enterocytes which are packed in a compact hexagonal lattice named the brush border (BB) (Granger & Baker, 1950; Grega-Larson et al., 2015; Sharkova et al., 2022). These microvilli notably play a role in nutrient absorption,

considerably increasing apical cell surface area (Grega-Larson et al., 2015). BB organization is also observed in the renal proximal tubule, where BB microvilli mediate solute exchange during urine formation (Weinbaum et al., 2010). Other well-characterized microvilli known as stereocilia underlie sound detection in the hair cells of the inner ear (Pelaseyed & Bretscher, 2018a). Stereocilia show a staircase organization with three rows of gradual heights and an eccentric cilium named the kinocilium (Pelaseyed & Bretscher, 2018b). When sound waves cause stereocilia deflection, tension is propagated to neighboring stereocilia through tip links, which mechanically open ion channels within the stereocilia membrane and trigger mechanotransduction (McGrath et al., 2017).

In this review, we define microvilli as finger-like, straight, and stable actin-filled cellular protrusions with a stereotypical length (Figure 1). Microvillar length and width are highly regulated within a given cell type but vary between cell types and species: about 4 µm in length and 100 nm in width for BB microvilli, 1–6 µm in length and 100–340 nm in width for stereocilia in mice and chicken (Hadi et al., 2020; Houdusse & Titus, 2021; Krey et al., 2023; Orbach & Su, 2020; Sharkova et al., 2022; Tilney et al., 1988). Stereocilia length and width decrease from one row to the next, the first row being the tallest and widest (Hadi et al., 2020; Krey et al., 2023).

Microvillar bundles are long-lived structures, with a lifetime of 12 h for BB microvilli (Meenderink et al., 2019); stereocilia do not regenerate and are thus stable lifelong (Yang et al., 2023). The morphological stability of microvilli reflects the slow turnover of their cytoskeletal components. Both BB microvilli and stereocilia are composed of parallel actin bundles (around 20–30 in BB; Figure 1) which only undergo assembly and disassembly at their tip (where plus [barbed] ends of F-actin filaments are located; [DeRosier, 2000; Goode et al., 2023]). Actin turnover occurs at the scale of hours for BB microvilli in *Caenorhabditis elegans* (Bidaud-Meynard et al., 2021; Ramalho et al., 2020) and days for stereocilia in mice (Goode et al., 2023; Narayanan et al., 2015) (consistent with globally slower protein turnover in stereocilia (Zhang et al., 2012)). Microvilli are however dynamic during their formation: immature BB microvilli move along the plasma membrane but become static and interconnected by protocadherins in their mature form (Meenderink et al., 2019). Some epithelial cells display short (<500 nm) and scattered microvilli that never assemble into bundles, and those have a much shorter lifetime (about 12 min in A6 toad kidney epithelial cells [Gorelik et al., 2003]). Some microvilli can also be remodeled under stress: BB microvilli shorten upon starvation (Liu et al., 2019), are retracted during intestinal wound healing and regeneration (Ubelmann et al., 2013), and can be replaced if damaged (Venkatachalam et al., 1978). By contrast, mammalian stereocilia cannot be regenerated (Yang et al., 2023). Mature microvilli are anchored to the underlying F-actin network by a
<figure><img src="image_2.png" /></figure>

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

<figure><img src="image_4.png" /><figcaption>**FIGURE 1** Molecular architecture of microvilli and filopodia. (a) Schematic representations of enterocyte, hair cell, and migrating mesenchymal cell. (b) Electron micrographs of mouse BB microvilli (B1), mouse stereocilia (B2), and filopodia (B3). Scale bar = 1 µm (B1)/ 1 µm (B2)/5 µm (B3). Reproduced from (Miihkinen et al. 2021; Postema et al. 2019; Vélez-Ortega et al. 2017). (c) Molecular composition of BB microvilli, stereocilia, and filopodia (see references in text). Proteins are color-coded by functional category.</figcaption></figure>
microvillar rootlet that imparts them a fixed position at the cell surface (Sharkova et al., 2022). The microvillar rootlet is composed of F-actin and specific proteins, such as myosin 6, TRIOBP, Taperin, Pejvakin, and spectrin in stereocilia (Pacentine et al., 2020). Movements are limited to occasional deflections (e.g., by sound waves [Kozlov et al., 2011] or during spontaneous oscillations [Martin et al., 2003]).

Finally, most microvilli occupy a stereotypical position at the apical pole of epithelial cells and are frequently linked to other cellular elements. Numerous microvilli are covered and protected by a glycocalyx-type extracellular matrix (Sharkova et al., 2022). Furthermore, microvilli are often (but not always) associated with microtubule-based cilia or flagella (Sharkova et al., 2022): stereocilia are physically linked to the kinocilium (Pelaseyed & Bretscher, 2018b) and microvilli in diverse animal cell types are arranged around a flagellum in a cellular architecture called the collar complex (Brunet & King, 2017; Salvini-Plawen, 1978).

To conclude, microvilli display a highly reproducible position in the cell and morphology (including a constant diameter and length for a given cell type). As we will see below, this sets them apart from other finger-like actin-based protrusions such as filopodia. Moreover, they tend to be linked by physical connections, mediating their assembly into bundles with a characteristic morphology (brush border in enterocytes and nephrocytes, stereocilia in hair cells). The structural and functional resemblance between the different mammalian microvilli is suggestive of a common evolutionary origin and, as we will see below, is further substantiated by molecular commonalities (Danielsen & Hansen, 2003; Hart et al., 2007; Vélez-Ortega et al., 2017).

## Molecular composition of microvilli: BB microvilli and stereocilia as paradigms

Most microvillar proteins common to BB and stereocilia can be classified into four functional groups: actin crosslinkers, actin capping proteins, actin-membrane linkers, and intermicrovillar linkers. Besides these four generic groups, two categories of proteins are respectively involved in BB microvilli and stereocilia specifically: molecular uptake proteins and mechanoelectrical transduction (MET) proteins (Figure 1). We will discuss below the nature and function of these six types of proteins.

### Actin crosslinkers

In both BB microvilli and stereocilia, the main microvillar actin crosslinkers are Espin and Plastin/Fimbrin, together with Fascin (in stereocilia) or Villin (in BB microvilli) (Sauvanet et al., 2015). Villin can switch from an actin-bundling to an actin severing activity by tyrosine phosphorylation and/or when exposed to high Ca²⁺ levels (Khurana & George, 2008), allowing microvilli retraction during wound healing (Ubelmann et al., 2013). The absence of Villin in stereocilia might thus contribute to their higher stability despite them being a locus of intense Ca²⁺ signaling (Sharkova et al., 2022). In BB microvilli, another actin crosslinker protein is Cordon bleu (COBL) which is necessary for microvillar growth and seems to participate in actin nucleation (Gregar-Larson et al., 2015). In addition, Epidermal growth factor receptor Substrate 8 (EPS8) and EPS8-like proteins can display an actin-bundling activity in BB microvilli and in stereocilia (Postema et al., 2018; Zwaenepoel et al., 2012). Finally, myosin 3A/B (MYO3A/B) contributes indirectly to actin bundling by transporting Espin and is critical to maintaining stereocilia shape over time (Houdusse & Titus, 2021).

### Actin capping proteins

The length of actin bundles and thus of microvilli is finely controlled by actin-capping proteins such as Twinfilin-2 (TWF2A) in stereocilia. TWF2A limits the growth of stereocilia during their biogenesis by capping actin filaments at their barbed-end and by sequestering actin monomers (Peng et al., 2009). ESP8 family proteins can exert a dual role as actin capping proteins and actin crosslinkers: for example, the EPS8-like protein EPS8L1 caps actin tips while EPS8 is involved in actin filament bundling in pig kidney epithelial cells (Zwaenepoel et al., 2012). In addition, myosin 15A (MYO15A) regulates stereocilia length by transporting and forming complexes with EPS8 and Whirlin at stereocilia tips (Houdusse & Titus, 2021).

### Actin-membrane linkers

Attachment of F-actin to the plasma membrane requires Ezrin, Radixin, and Moesin family proteins (ERM) in all microvilli, but the type of ERM involved differs between cell types: Ezrin predominates in enterocyte microvilli but is only transiently expressed in stereocilia, which mainly rely on Radixin instead (Sauvanet et al., 2015). ERM binds the plasma membrane phospholipid phosphatidylinositol 4,5-bisphosphate (PIP₂) and toggles between inactive and active conformation regulated by phosphorylation. In BB microvilli, another essential protein is Ezrin-radixin-moesin Binding Phosphoprotein 50 (EBP50 also known as NHERF1), which binds Ezrin on its FERM domain (Vélez-Ortega et al., 2017) and stabilize ERMs at the membrane (Morales et al., 2004). Although NHERF1 has been detected in stereocilia proteome and could likely interact with the FERM domain of Radixin, this interaction remains to be demonstrated. Other actin-membrane linkers can specifically
bind curved membranes via a BAR domain. Among these, I-BAR domains preferentially bind concave membrane domains (such as the tip of microvilli), while Fes/CIP4 homology BAR (F-BAR) domains bind convex membranes (such as the base of microvilli) (Zhao et al., 2011). I-BAR proteins located at microvillar tips include Insulin Receptor Tyrosine Kinase Substrate (IRTKS) and BAR/IMD Domain Containing Adaptor Protein 2 Like 2 (BAIAP2L2) (Carlton et al., 2021; Postema et al., 2018; Zwaenepoel et al., 2012). IRTKS (also known as BAIAP2L1) is expressed in BB microvilli (Postema et al., 2018) while BAIAP2L2 is found at the tips of stereocilia (Carlton et al., 2021). In contrast, F-BAR proteins such as Pacsin-1 and Pacsin-2 (also named Syndapin-2) are respectively found at the base of stereocilia and of BB microvilli. Pacsin-2 regulates the morphology of the base of BB microvilli by influencing local endocytic activity (Postema et al., 2019). Besides ERM, I-BAR, and F-BAR proteins, the myosins MYO1A/D and MYO1C connect the actin bundles to the plasma membrane of both BB microvilli and stereocilia respectively by interacting with PIP₂, and generate a membrane tension toward the tips (Benesh et al., 2010; Houdusse & Titus, 2021). An opposite tension is generated by myosins located at the microvillar base: non-muscle myosin IIC in BB microvilli, (MYO2C, also called MYH14) and MYO6 in both microvilli (Houdusse & Titus, 2021). As a result of these two opposite forces, microvilli are under constant tension.

## Intermicrovillar linkers

Microvilli belonging to the same cell are linked to each other by a complex named the Intermicrovillar Adhesion Complex (IMAC) in BB microvilli or the Usher complex (USH1 proteins) in stereocilia. Adhesion complexes consist of cadherins mediating direct adhesion between microvilli (CDHR2/CDHR5 in the IMAC, CDH23/PCDH15 in the Usher complex), of myosin 7A/B (MYO7A/B) that binds the actin core (MYO7A in stereocilia and MYO7B in brush border), and of two adaptor proteins linking the cadherins to MYO7A/B and thus to F-actin: Harmonin and Ankyrin repeat and Sterile α motif domain containing 4B (ANKS4B) in the IMAC, or Harmonin and the ANKS4B paralog Scaffold protein containing Ankyrin repeats and SAM domain (SANS) in the Usher complex (Pelaseyed & Bretscher, 2018b). Myosin 5B (MYO5B) participates in the transport of IMAC component in BB microvilli (Dooley et al., 2022). In the absence of the IMAC or the Usher complex, microvilli organization is disturbed (Caberlotto et al., 2011; Pinette et al., 2019), altering microvilli function and sometimes leading to diseases, notably hearing loss for the Usher complex (El-Amraoui & Petit, 2005). Finally, in addition to this common complex, intermicrovillar linkers specific to individual types of microvilli exist. In BB microvilli, it

has been recently discovered that nanobristles based on CDH-8 cadherin connect adjacent microvilli along their lateral side in *C. elegans*, and similar structures have been also observed in mice (Zhu et al., 2022). In stereocilia, a different adhesion complex has been documented at the stereocilia ankle that notably depends on Usherin, Whirlin, and PDZ Domain Containing 7 (PDZD7) (Sharkova et al., 2022; Zou et al., 2014), and stereocilia are also laterally connected by Stereocilin proteins (Verpy et al., 2011).

## Transporters

BB microvilli feature multiple proteins involved in the transmembrane transport of various molecules, and notably in ion exchange and nutrient uptake. In enterocyte microvilli, these include the Na⁺/H⁺ exchangers 3 (NHE3) which mediates most of the sodium absorption, is coupled with chloride (Cl⁻) import, and indirectly impacts peptide uptake via Peptide Transporter 1 (PEPT1) (Kiela & Ghishan, 2016). Na⁺ uptake is also achieved by the Epithelial sodium channel (ENaC) in enterocyte and renal BB microvilli (Kiela & Ghishan, 2016; Okamoto, 2017). Sucrose is imported by Sodium/glucose cotransporter 1 (SGLT1) and Glucose transporter 5 (GLUT5), and cholesterol is transported through Niemann-Pick C1 like-1 (NPC1L1) and the ATB-binding cassette protein G (ABCG) (Kiela & Ghishan, 2016). Phosphate is uptaken by the Na⁺ phosphate transporter NaPi-IIIb, and magnesium through Transient potential ion channels 6 and 7 (TRPM6 and TRPM7) (Kiela & Ghishan, 2016). The regulation of these proteins is not fully understood but seems to involve the small GTPases Rab11a and Rab8a, as well as MYO5B, SNARE-related proteins, and NHERF1 (Okamoto, 2017).

## Mechanotransduction proteins

MET is the primary function of stereocilia (although BB microvilli might sometimes also be mechanosensitive [Miura et al., 2015]). MET actors mediate sound detection and exert positive feedback on stereociliary growth and maintenance (Dunbar et al., 2019; Vélez-Ortega et al., 2017) The ion channels responsible for the mechanical-to-electrical signal conversion are the Transmembrane-Channels, TMC1 and TMC2 (Kurima et al., 2015), which are regulated by Calcium and Integrin-Binding protein 2 (CIB2) and CIB3 (Liang et al., 2021; Wang et al., 2023). MET depends on the physical connection between stereocilia, and thus on the intermicrovillar linkers. For example, Glutaredoxin and Cysteine Rich Domain Containing enzyme 1 (GRXCR1 or GRCR1) modulates the interaction between Harmonin and SANS, which is critical for MET (Blanco-Sánchez
et al., 2018; Lorente-Cánovas et al., 2022). Finally, the transmembrane protein Clarin-2 is also essential for MET function but its precise molecular role remains unknown (Dunbar et al., 2019).

In spite of the classification of microvillar proteins in these six functional categories, interactions have been documented between proteins belonging to distinct categories; for example, EPS8 directly interacts with phosphorylated Ezrin and I-BAR proteins (Postema et al., 2018; Zwaenepoel et al., 2012), and Pacsin-2 might recruit COBL during the initiation of BB microvilli formation (Grega-Larson et al., 2015).

Overall, BB microvilli and stereocilia express a strongly similar core of structural proteins, suggestive of a common origin. Deciphering the evolutionary origin of this core set of proteins will thus likely be key to reconstituting microvilli evolution.

## Differences between microvilli and filopodia

Although filopodia resemble microvilli in being actin-filled finger-shaped cellular protrusions, they differ in being less stable, more flexible, and more exploratory. Indeed, filopodia display actin turnover time on the order of minutes (Mallavarapu & Mitchison, 1999), display frequent elongation and retraction, and have a shorter lifetime than microvilli (2–3 min in HeLa, N1E115, COS7, and B16F1 cells [Lim et al., 2008]; about 23 min in Xenopus neurons [Robles et al., 2003]). Additionally, they tend to be longer (up to 40 µm; [Heckman & Plummer, 2013]), faster-growing (up to 240 nm/s [Bornschlögl, 2013]), and sparser than microvilli (Kyykallio et al., 2020). In filopodia, actin bundle assembly and disassembly do not result in a dynamic equilibrium like in microvilli but occur during episodes of fast demolition causing filopodia to collapse (Goode et al., 2023). Filopodia can buckle and twist (Leijnse et al., 2022) and can also move laterally, turn around their anchor points, and exert pulling forces (Bornschlögl, 2013). Filopodial motility allows active sampling of extracellular information and/or mechanical deformation of the surrounding tissue (Laishram et al., 2009). This allows filopodia to contribute to cell migration, to dynamic cell-cell adhesion (notably during wound healing), and to serve as phagocytic tentacles in macrophages (Kress et al., 2007; Mattila & Lappalainen, 2008). Another important structural difference between filopodia and microvilli is that filopodia do not adhere to neighboring filopodia within the same cell, but instead tend to adhere to other cells (including at the level of their filopodia) and/or to the extracellular matrix (Mattila & Lappalainen, 2008).

In terms of molecular makeup, filopodia resemble microvilli in relying on Fascin as the main F-actin crosslinker (Mattila & Lappalainen, 2008) but

differ in their complement of actin capping proteins (Enabled/Vasodilator-Stimulated Phosphoprotein [ENA/VASP]), actin-membrane linkers (Talin [Mattila & Lappalainen, 2008] and myosin 10 (MYO10)—although ERM is involved in dendritic filopodia [Furutani et al., 2007]). MYO10 is critical for filopodial initiation, elongation, and function in several ways: it organizes actin filaments, acts as a cargo transporter, and mediates integrin localization at the filopodia tips (Houdusse & Titus, 2021). Finally, filopodia biogenesis requires actin nucleation (de novo or from pre-existing lamellipodia) by the Actin Related Protein 2/3 (ARP2/3) complex (Mattila & Lappalainen, 2008), while this complex appears dispensable for the formation of BB and T-cell microvilli (Bernadskaya et al., 2011; Nicholson-Dykstra & Higgs, 2008; Zhou et al., 2015). In stereocilia, the ARP2/3 complex has been detected by proteomics (Krey et al., 2023) and might thus play a role, but functional evidence is lacking (Zhai et al., 2022).

Thus, microvilli and filopodia differ in molecular composition, structure, and function. Microvilli are specialized for mostly passive uptake or detection of environmental biotic or abiotic factors, whereas filopodia actively explore their environment and generate forces. Accordingly, filopodia have higher mobility and faster turnover, do not form bundles, and lack the stereotypic organization and localization of microvilli. Below, we will use these definitions of filopodia and microvilli, based on animal cells, to classify actin-based protrusions in less well-characterized eukaryotes, and attempt to reconstitute microvilli origin and evolution.

## DIVERSITY OF MICROVILLI ACROSS METAZOANS AND THEIR UNICELLULAR RELATIVES

Microvilli have been best-characterized in vertebrates and especially in mammals, but are also found in diverse animal lineages, as well as in the closest living relatives of animals, the choanoflagellates and the filastereans (Figure 2). We give below a brief overview of microvilli diversity.

### Diversity of metazoan microvilli

Several types of microvilli are conserved within metazoans. BB microvilli involved in capture and transport of small molecules (for nutrient absorption or excretion) have been described across chordates (Hartenstein & Martinez, 2019), annelids (Rost-Roszkowska et al., 2012), arthropods (Steinmetz, 2019), and nematodes (Geisler et al., 2019) (Figure 2a) but seem absent in non-bilaterians (Hartenstein & Martinez, 2019). Examples include fruit fly renal tubules (Cohen et al., 2020)
Box 1: Are T-cell microvilli related to epithelial apical microvilli?

Both BB microvilli and stereocilia are present at the apical pole in epithelial cells, but microvilli without a polarized distribution have been described in non-epithelial cell types. For instance, the term microvilli is used to refer to small actin-based finger-like protrusions covering the surface of T-cells. T-cells are lymphocytes that play an essential role in jawed vertebrate immunity (Hirano et al., 2011) by detecting ligands on the antigen-presenting cells (APCs) through their microvilli, and triggering downstream immune responses. Like apical microvilli, T-cell microvilli are short (around 0.4 µm) and present at high density (3–4 per µm²) (Majstoravich et al., 2004), but they also share characteristics with filopodia. They display an active behavior with fast displacements at around 5 µm/min (Cai et al., 2017). This velocity is on the same order of magnitude as the lateral movements of filopodia (Oldenbourg et al., 2000) and almost 30 times higher than the velocity of immature BB microvilli (Meenderink et al., 2019). Besides proteins playing a role in immunity, almost nothing is known about the molecular composition of T-cell microvilli (Cebecauer, 2021). It has been only shown that ERM is necessary for their formation (Ghosh et al., 2020) and that WASP might contribute to it (as in filopodia), but its role remains debated (Kim & Jun, 2019). Moreover, unlike other microvilli that are assembled into bundles by specific adhesion proteins, T-cell microvilli lack intermicrovillar adhesions (Jung et al., 2016). T-cells are an evolutionary innovation of jawed vertebrates and likely evolved from lymphocyte-like macrophages (Hirano et al., 2011). Thus, because macrophages possess filopodia and not microvilli (Kress et al., 2007), it is tempting to interpret T-cell microvilli as modified filopodia unrelated to other microvilli, such as those of enterocytes and hair cells. Nevertheless, without more information on their molecular composition, it is difficult to retrace their evolutionary history. Therefore, they could either be modified filopodia or microvilli that have acquired certain filopodial features to perform new functions.

and enterocytes (Hegan et al., 2007), as well as *C. elegans* enterocytes (McGhee, 2007), where they play the same role as their mammalian counterparts. The molecular composition of BB microvilli appears conserved: for example, BB microvilli in *C. elegans* rely on homologs of Plastin, MYO1D/G, ERM, EBP50, EPS8, and protocadherins (Bidaud-Meynard et al., 2021; Sepers et al., 2022; Zhu et al., 2022) and *C. elegans* intestinal cells express common transporters with mice BB microvilli (Zhu et al., 2022).

Stereocilia *stricto sensu*, defined by their staircase organization with rows of gradual heights and an eccentric kinocilium, appear unique to vertebrates (Fritzsch & Straka, 2014). However, bundles of mechanosensitive microvilli with slightly different architectures are known in other bilaterians: microvilli associated with an eccentric kinocilium (without a staircase arrangement) have been observed in tunicates (Burighel et al., 2003). Other metazoans display putative mechanosensory microvilli associated with multiple cilia (Fritzsch & Straka, 2014) or surrounding a central kinocilium, sometimes (but not always) arranged in a single ring resembling a collar complex (Amiel et al., 2021; Bezares-Calderón et al., 2019) (Figure 2b).

In addition to sound detection, microvilli are involved in metazoan light perception with different contributions in vertebrates and invertebrates. In all bilaterians except chordates, visual phototransduction occurs preferentially in microvilli, such as the rhabdomeres in arthropods (Figure 2d) (Fain et al., 2010). Microvillar photoreceptors are also known in invertebrate deuterostomes (such as sea urchins [Ullrich-Lüter et al., 2011]), suggesting they were present in Urbilateria. Microvilli facilitate light detection by enlarging the apical surface area and concentrating photoreceptor proteins (opsins) (Fain et al., 2010). Ciliary photoreceptors are also known in protostomes, but their function seems non-visual (Arendt et al., 2004). By contrast, in vertebrates, a ring of microvilli named “calyceal processes” is present in photoreceptor cells (Sahly et al., 2012) but opsins are found exclusively in a modified cilium at the center of the ring (Fain et al., 2010). Calyceal processes are present in primates (Figure 2c), frogs, and zebrafish, but have been lost in rodents (Sahly et al., 2012). In zebrafish, calyceal processes are 3–6 µm long and 130 nm wide, as typical for microvilli (Sharkova et al., 2024). The function of calyceal processes remains unclear. Even though they are not directly involved in light detection, mutations affecting those microvilli lead to retinal degeneration (Miles et al., 2021).

Metazoan microvilli can also be involved in chemosensation, notably olfaction. Vertebrate olfaction mostly relies on cilia, but some olfactory sensory neurons in the vomeronasal system of tetrapods use microvilli (Poncelet & Shimeld, 2020) and non-neuronal olfactory microvillated cells have been reported in mice (Genovese & Tizzano, 2018). Furthermore, other
<figure><img src="image_5.png" /><figcaption>**FIGURE 2** Diversity and morphology of microvilli among Metazoa and their close relatives. (a) *Caenorhabditis elegans* brush border microvilli. (b) *Nematostella vectensis* (sea anemone) mechanosensory microvilli. (c) *Macaca fascicularis* (macaque) calyceal processes. (d) *Photuris sp.* (firefly) rhabdomeres. (e) *Hypomesus transpacificus* olfactosensory neuron (OSN) microvilli -, (f) *Cotylorhiza tuberculata* (jellyfish) microvilli arranged as a collar complex. (g) *Tethya wilhelma* choanocytes with microvilli arranged as a collar complex. (h) *Salpingoeca rosetta* microvilli arranged as a collar complex. (i) *Ministeria vibrans* microvilli with a non-polarized distribution. Scale bar = 0.5 µm (a)/1 µm (c)/250 nm (d)/1 µm (f)/3 µm (i)/5 µm (g). mv = microvilli, f = flagellum, c = cilium. Adapted from (Amiel et al. 2021; Dayel 2008; Geisler et al. 2023; Hammel & Nickel 2014; Murphy & Moiseff 2019; Ruiz-Trillo 2014; Sahly et al. 2012; Solé et al. 2016; Triana-Garcia et al. 2021).</figcaption></figure>

vertebrate chemosensory cells sport microvilli such as Cerebrospinal Fluid-contacting Neurons (CSF-cNs) (Figure 2e) (Wyart et al., 2023) and tuft cells (Iqbal et al., 2023) which detect pathogens and trigger immune responses (Iqbal et al., 2023) in the epithelia of the air-way (Du et al., 2023), the small intestine (Montoro et al., 2020), and the urethra (Kummer & Deckmann, 2017).

Interestingly, microvilli across many lineages of meta-zoans are often organized as a collar complex—that is, as a ring of microvilli surrounding a central cil-ium/flagellum (Figure 2f,g). They are found in all phyla (except in Ctenophora and in certain Ecdysozoa) as

well as in the sister-group of animals, Choanoflagel-lata (Figure 2h)—making the collar complex a plausible ancestral feature of metazoans (Brunet & King, 2017). Collar complex microvilli are often involved in feeding: they mediate capture of particles prior to phagocyto-sis in sponges choanocytes (Lavrov et al., 2022) and nutrient absorption in many invertebrate enterocytes (or gastrodermal cells) (Brunet & King, 2017), but they can also play a role in mechanosensation (e.g., in cnidarians [Solé et al., 2016]) and in excretion (Arendt, 2021).

To conclude, microvilli are observed in all main meta-zoan lineages, with broadly conserved functions includ-
ing nutrition, excretion, mechanosensation (including hearing), light detection, and chemosensation (including olfaction).

## Choanoflagellate microvilli

The closest living relatives to animals are a group of aquatic protists named the choanoflagellates (Brunet & King, 2017; Grau-Bové et al., 2017; King & Carroll, 2001; Ruiz-Trillo et al., 2008; Sebé-Pedrós, Degnan et al., 2017). Choanoflagellates display microvilli surrounding a motile flagellum in a collar complex arrangement (Figure 2h). It has been noticed for more than a century that the choanoflagellate and the sponge collar complex are structurally and functionally similar (Figure 2g,h) (Brunet & King, 2022; James-Clark, 1867; Saville-Kent, 1880). In both lineages, adjacent microvilli are connected by intermicrovillar links and are covered by a glycocalyx even if their arrangement differs (Leys & Hill, 2012; Mah et al., 2014). Furthermore, their function appears similar. In both, the flagellum produces a water flow that traverses or surrounds the collar of microvilli which captures the bacteria. In this organization, the coupling between the flow created by the flagellum and the capture by the static but dense microvilli ensures efficient feeding. In some choanoflagellate species only (the loricates), the microvilli also perform an additional function: the deposition and the patterning of an extracellular silicified covering named the lorica (Leadbeater & Carr, 2022). The molecular composition of choanoflagellate microvilli is mostly unknown, but they rely on bundles of F-actin and incorporate fascin (Sebé-Pedrós et al., 2013).

## Are there microvilli in filasterea?

Microvilli have traditionally been considered exclusive to Choanozoa, the monophyletic clade regrouping the choanoflagellates and the metazoans, but recent studies have classified the “arms” found in the filasterean *Ministeria vibrans* as microvilli (Figure 2i) (Mylnikov et al., 2019). Like microvilli, those protrusions are straight, stable in time, and uniform in length and width. However, they are distributed sparsely and homogeneously over the cell surface, thus lacking the dense and polarized organization observed in choanoflagellate, stereocilia, or BB microvilli. Relatedly, no physical connections between adjacent arms were observed (and interconnections are perhaps unlikely to exist given their sparse distribution) (Mylnikov et al., 2019). In this respect, they are closer to filopodia or T-cell microvilli. Although *M. vibrans* sports a static flagellum (through which it attaches to external surfaces), the arms are not organized around the flagellum in a collar complex unlike

in choanoflagellates and choanocytes. Nevertheless, *M. vibrans* microvilli may also mediate the capture of bacteria (Mylnikov et al., 2019). The well-studied model filasterean *Capsaspora owczarzaki* lacks microvilli (and a flagellum), but displays dynamic filopodia that it uses to crawl on substrates and to aggregate with other cells (Parra-Acero et al., 2020; 2018; Ros-Rocher et al., 2023; Sebé-Pedrós et al., 2013). Finally, other potential actin-based protrusions have been noticed in the Filasterea *Pigoraptor chileana* and *Pigoraptor vietnamica* (Tikhonenkov et al., 2020), but more precise descriptions are required to classify them as either microvilli or filopodia. Thus, the *M. vibrans* microvilli resemble choanozoan microvilli in form and possibly in function (although this remains to be tested) but differ by their less dense organization and unpolarized distribution.

## PHYLOGENETIC DISTRIBUTION OF MICROVILLAR PROTEINS

To determine if the various classes of microvilli are homologous or result from convergent evolution, it is essential to elucidate their molecular composition. Although the detailed molecular composition of microvilli has only been determined for a few species and remains time-consuming to establish, a feasible approach is to survey the phylogenetic distribution of canonical mammalian microvillar proteins across sequenced eukaryotic genomes and transcriptomes. We provide here such a survey, with a focus on choanozoans and their closest relatives. Earlier studies either focused on filopodial genes (Sebé-Pedrós et al., 2013) or identified homologs based on mutual best BLAST hits (Peña et al., 2016). However, BLAST-based analyses can miss homologs at large evolutionary distances and many additional genomes and transcriptomes have been sequenced in the past few years (notably of choanoflagellates), allowing to survey microvillar gene homologs among a broader taxonomic sample than earlier studies. Thus, we decided to re-explore the phylogenetic profiles of microvillar proteins using the Hidden Markov Model (HMM) method.

In contrast to BLAST-based approaches, HMM-based methods build probabilistic profiles (profile HMMs) which indicate the likelihood of observing a certain residue at a specific column of a multiple sequence alignment (Eddy, 2023). HMMs are thought to be better for dealing with amino acid insertions and deletions, and thus for detecting similarities that are not apparent through traditional pairwise sequence comparisons. Accordingly, HMM-based methods can detect homologs even among sequences with low sequence similarity (Park et al., 1998).

We investigated the evolutionary origin of the human microvillar proteins detailed above, grouped in 20 gene
Box 2: Oocyte microvilli and their role in sexual reproduction

Oocyte microvilli are microvilli covering the whole cell surface and displaying canonical microvilli morphology. They are about 1 µm long and 100 nm wide in mice and cnidarians (Clark & Dewel, 1974; Longo, 1987). During oocyte maturation, they are believed to maintain contact between oocytes and somatic cells through the zona pellucida and to secrete factors regulating the activity of the surrounding somatic cells (Zhang et al., 2021). During fertilization, fusion of the sperm cell with the oocyte occurs at microvillus tips in sea urchin (Wilson & Snell, 1998). After sperm cell fusion, oocyte microvilli are endocytosed, providing a store of membrane for the rapid serial rounds of cell division that follow fertilization (Courjaret et al., 2016). Interestingly, a similar membrane storage role has also been described in certain epithelial microvilli (Figard & Sokac, 2014; Lange, 2011). Almost nothing is known about the molecular composition of oocyte microvilli, but radixin has been shown to be critical for their formation in mice (Zhang et al., 2021). Oocyte microvilli appear broadly distributed within animal phylogeny. They have been described in mammals (Zhang et al., 2021), fish (Abraham et al., 1984), sea urchins (Wilson & Snell, 1998), and cnidarians (Eckelbarger & Larson, 1992). Ctenophore oocytes also display microvilli but only after fertilization and they are restricted to the surroundings of the fertilization site (Carré et al., 1991). As for T-cell microvilli, a more complete molecular profile will be required to know if oocyte microvilli are homologous, if they are related to epithelial microvilli, and to reconstruct their evolutionary history. Interestingly, during sponge sexual reproduction, sperm cells are thought to first be captured by a choanocyte that then transfers the sperm to the oocyte, thus acting as a "sperm carrier cell" (Franzen, 1988) (but see [Anakina & Drozdov, 2001]). Microvilli also mediate cell-cell adhesion prior to sexual fusion in choanoflagellates (Levin & King 2013). A role for microvilli in sexual reproduction might thus be ancestral for metazoans and potentially choanozoans.

families (Figures 1–3), and examined their distribution across 105 Amorphea species (Figures 3 and S1; Supplementary Tables S1, S2 and S3). The Amorphea clade includes animals, fungi, their respective holozoan and holomycotan unicellular relatives (such as choanoflagellates and filastereans), amoebozoans (which includes amoebae such as *Dicytostelium* and *Acanthamoeba*), and all descendants of the last common ancestor of these groups. We decided to not re-investigate the distribution of myosins and cadherins because these large protein families with complex evolutionary histories would deserve studies of their own and have been already analyzed in detail (Nichols et al., 2012; Pasha et al., 2016; Sebé-Pedrós et al., 2014). The phylogenetic distribution of members of these two families was thus synthesized from the literature (gray circles in Figure 3). Finally, although the distribution of BB microvilli transporters is not displayed here, many of them, such as glucose transporters and NHE3, are broadly conserved across eukaryotes (Brett et al., 2005; Han et al., 2022; Hirsch et al., 1998; Jia et al., 2019; Milenkovic et al., 2010; Ott et al., 2023; Xiong et al., 2015). This suggests that transporters have largely been repurposed for microvillar transport of molecules from an earlier role in uptake of molecules from the environment or in transport between organelles within the cell.

Actin cross-linkers

Consistent with the literature (Peña et al., 2016; Sebé-Pedrós et al., 2013), we detected members of the Plastin (also called Fimbrin) and Villin families broadly across Amorphea. Most Villin family members that we identified displayed the headpiece domain characteristic of Villin sensu stricto (and of its close relatives Villin-like and Advillin), which sets them apart from other Villin family members such as Gelsolin and Scinderin (Figure S2). Fascin was almost only detected in Filozoa, the monophyletic group comprising Choanozoa and Filasterea (Figure 4), but surprisingly also in one amoebozoan, *Acanthamoeba* (Figure S1). As reported in a previous study (Schultz & Terhoeven, 2013), COBL was only found in bilaterians. We did not detect the COBL homolog of *Drosophila melanogaster*, but did find one in another insect, the bee *Apis mellifera* (Figure S1). Finally, the analysis of the Espin family was complicated by the presence of multiple Ankyrin repeat domains, which are present in a broad range of proteins unrelated to Espin. We built a phylogenetic tree of all the proteins detected in our first, broad preliminary survey, and only classified as bona fide Espin homologs those that matched a stricter protein domain annotation criterion (see Methods) and that belonged to a set of clades including known Espin homologs but excluding unrelated proteins (Figure S3). Thus defined, Espin
<figure><img src="image_6.png" /><figcaption>**FIGURE 3** Phylogenetic distribution of microvillar proteins. For each lineage, the black area within the pie chart indicates the proportion of species within that lineage having a homolog of the gene of interest. Black circles indicate that all species of the clade have a homolog and white circles mean that no homolog has been detected. Myosins and cadherins, which we did not analyze ourselves, are either indicated by a gray circle if homologs have been reported in the literature or by a white circle if no homologs are known (Nichols et al. 2012; Pasha et al. 2016; Sebé-Pedrós et al. 2014). Note that NHERF1 is referred to as "NHRF1" in this figure, in supplementary figures, and raw data. Proteins are classified in the same functional categories as in Figure 1 ("AC proteins" = Actin Capping proteins). The number of species in each lineage is written in parentheses after the lineage name. Detailed phylogenetic profiles for each species are in Figure S1. "MR clade" = Microsporidia and Rozellida. The red microvilli schematics indicate lineages in which at least one species displays microvilli. The yellow boxes outlining certain clades are meant as a visual guide to help follow the alignment of pie charts indicating the presence/absence of gene families.</figcaption></figure>

was detected in Filozoa, Ichthyosporea, some Holomy-
cota, Apusomonadia, and one amoebozoan species,
*Acanthamoeba castellanii* (Figure S1).

Actin-membrane linkers

Although morphologically recognizable microvilli only
exist in Filozoa, most actin-membrane linkers (BAPIA2,
Pacsin2, MYO1, MYO2, and MYO6) had a broader
distribution in our analysis. This is consistent with actin-
membrane attachment also having a broader function
in the formation and maintenance of a generic actin
cortex underneath the plasma membrane, which main-
tains cell shape and allows amoeboid motility in diverse
eukaryotes (Brunet et al., 2021; Chugh & Paluch, 2018;
Condeelis, 1993). However, ERM and NHERF1 appear
specific to Choanozoa. Although we detected ERM-like
proteins in Filasterea (in accordance with an earlier
study [Shabardina et al., 2020]) and in Ichthyosporea,
our phylogenetic tree placed them closer to the ERM-
like protein Merlin than to ERM stricto sensu (Figure
S4), and we thus do not classify them here as bona
fide ERM proteins. Due to the presence of repeated

PDZ domains, a few unrelated proteins were detected
alongside NHERF1 in our preliminary screen. Therefore,
bona fide NHERF1 homologs were defined follow-
ing a stricter criterion for protein domain composition
(Methods, Figure S5).

Intermicrovillar linkers, actin capping
proteins, and mechanotransduction
proteins

Most intermicrovillar proteins were restricted to Filo-
zoa, except for a Harmonin family homolog detected
in one ichthyosporean (Figure S1). As for NHERF1,
Harmonin family proteins contain PDZ repeats, and
bona fide Harmonin family homologs were thus iden-
tified with stricter annotation criteria (Methods, Figure
S6). In contrast to previous analyses (Peña et al.,
2016), SANS/ANKS4B was found in Choanoflagellata,
and Usherin and AGRV1 in both Choanoflagellata
and Filasterea. Furthermore, Stereocilin was restricted
to Choanozoa. Regarding Actin Capping proteins (AC
proteins), Twinfilin-2 homologs were found broadly in
Amorphea, and ESP8-like was restricted to Choanozoa,
<figure><img src="image_7.png" /><figcaption>**FIGURE 4** A scenario for the origin and evolution of microvilli. Simplified Amorphea phylogeny mapped with evolutionary events related to microvilli evolution. A green circle indicates the gain and a red cross the loss of the corresponding trait. The position of microvilli evolutionary events are interpreted from (Brunet & King 2017; Fain, Hardie, & Laughlin 2010; Fritzsch & Straka 2014; Hartenstein & Martinez 2019; Iqbal et al. 2023; Lamb 2013; Mylnikov et al. 2019; Poncelet & Shimeld 2020; Sahly et al. 2012; Sebé-Pedrós et al. 2013), and the Amorphea phylogenetic tree has been reconstructed using (Brunet & King 2017; Galindo et al. 2021; Hamann et al. 2016; Heger et al. 2020; Liao et al. 2023; Tikhonenkov et al. 2020). Species silhouettes are from PhyloPic (https://www.phylopic.org).</figcaption></figure>

as previously reported (Peña et al., 2016; Sebé-Pedrós et al., 2013). Finally, for MET actors, TMC1/2 and GRCR1 had a scattered distribution across Amorphea, while CIB2/3 and Clarin-2 were choanozoan- and metazoan-specific, respectively. Alongside CIB2/3, members of the related family CANB2 and CIB1/4 were also detected in our HMM-based survey, and bona fide CIB2/3 homologs were identified based on a phylogenetic tree and strict protein domain annotation (Figure S7). Overall, this suggests that microvillar mechanotransduction largely evolved by co-option of

pre-existing mechanosensors, rather than relying on the invention of new ones. This is consistent with an earlier report of a pre-amorphean origin for TMC (Erives & Fritzsch, 2020).

## The evolutionary assembly of the microvillar molecular toolkit

To sum up, homologs for almost all canonical microvillar proteins were identified in species displaying microvilli
(Filozoa) but few were exclusive to those species. In line with previous studies (Peña et al., 2016; Sebé-Pedrós et al., 2013), ctenophores lost most microvillar proteins, including notably intermicrovillar linkers, consistent with the apparent absence of physical connection between their microvilli (Norekian & Moroz, 2019). Fascin, MYO3, Usherin, AGRV1, and Cadherins appear to have evolved concomitantly with the emergence of microvilli at the base of Filozoa, suggesting they might have played a critical role in the origin of microvilli. However, some of these proteins might have originally played a role in filopodia as well: indeed, several of them are present in *Capsaspora*, a filasterean amoeba that lacks microvilli but has filopodia. In contrast to filasterean microvilli, choanozoan microvilli can be physically tethered to each other, allowing their assembly in bundles. The identification of the intermicrovillar linkers SANS/ANKS4B and Stereocilin as choanozoan innovations might indicate that intermicrovillar connections are an ancestral choanozoan feature. In the future, functional experiments will be key in determining the function of these microvillar protein homologs in non-model species.

# A SCENARIO FOR THE ORIGIN AND DIVERSIFICATION OF MICROVILLI

## From filopodia to microvilli in filozoa ancestors

Microvilli have been described in Metazoa, Choanoflagellatea, and one genus within Filasterea. Based on morphological data (Figure 2) and the phylogenetic distribution of homologs of metazoa microvillar genes (Figure 3), it can be hypothesized that microvilli can be traced back to the Urfilozoan, the common ancestor of Filasterea and Choanozoa. However, the molecular composition of microvilli outside Metazoa (and even outside Bilateria) is generally unknown, and determining it will be crucial in the future to determine the evolutionary history of microvilli.

Regardless of whether microvilli evolved once or multiple times, it can be hypothesized that they originated by modification of filopodia. Although filopodia and microvilli share some aspects of morphology and molecular ultrastructure, filopodia are likely more ancient, as filopodia-like structures have been reported in multiple amorphean outgroups to filozoans, such as amoebozoans (Dumontier et al., 2000; Medalia et al., 2007), apusozoans (Cavalier-Smith & Chao, 2010), and nucleariids (Mikrjukov & Mylnikov, 2001) (as well as in some more distant, non-amorphean eukaryotes such as *Naegleria* [Velle & Fritz-Laylin, 2020]).

Microvilli use part of the filopodial molecular toolkit such as Fascin, Plastin, and IRSp53, but differ by specific proteins such as Villin and a set of specialized

myosins (MYO3, MYO6, MYO7, and MYO15) different than the filopodial MYO10. Interestingly, the filopodia-specific myosin MYO10 and the microvillar myosins MYO7/MYO15 appear to be sister-groups (within the broader MyTH4-FERM myosin clade), suggesting a gene duplication (giving rise to *myo7/15* and *myo10*) might have accompanied the organelle duplication that gave rise to filopodia and microvilli. The ancestral MyTH4-FERM myosin might have resembled the amoebozoan DdMYO7, which is involved in filopodia (Petersen et al., 2016). This suggests a gene duplication alongside organelle duplication. The first microvilli might have been present in the last filozoa common ancestor, and might have resembled those of *Ministeria*, and thus might have already been straight, stable, and of constant length, but not yet restricted to a single pole of the cell or assembled in a cohesive bundle. They might already have functioned in feeding (Sebé-Pedrós et al., 2013), as the microvilli of choanoflagellates and sponges (and potentially *Ministeria*) all perform prey capture, and the microvilli of other metazoans often mediate nutrient uptake. However, the mechanisms involved differ between lineages: phagocytosis followed by intracellular digestion (as in choanoflagellate and choanocytes) is mechanistically distinct from uptake of individual soluble molecules after extracellular digestion (as in BB microvilli) (Steinmetz, 2019). It is unknown whether choanoflagellate microvilli might also mediate the uptake of some nutrients. Interestingly, sponge choanocytes can absorb dissolved organic matter (Rix et al., 2020) and choanoflagellates have recently been found to uptake iron from ferric colloids (Leon et al., 2024), raising the possibility that microvilli of sponges and choanoflagellates might play an underrated role in nutrient uptake and not just in the capture of bacteria to be phagocytosed. Because diverse microvillar transporters (such as those in BB microvilli) are of ancient eukaryotic origin (Han et al., 2022; Hirsch et al., 1998; Jia et al., 2019; Milenkovic et al., 2010; Ott et al., 2023; Xiong et al., 2015), their cellular localization in a range of phylogenetically relevant species will need to be determined to pinpoint when in evolution microvilli first started to perform nutrient uptake.

In addition to nutrition, ancestral microvilli might already have performed some of the other functions of modern microvilli, such as stores of membrane as a protection against osmotic shock, xenobiotic defense (Lange, 2011), or mechanosensation. Indeed, microvilli are often mechanosensitive not only in animals (Duan et al. 2008; Miura et al., 2015; Vélez-Ortega et al., 2017) but also in choanoflagellates [Leadbeater, 1983; Ros-Rocher & Brunet, 2023]). Consistently, some key microvillar MET proteins predate the emergence of filozoans (Figure 3). Moreover, animal filopodia can also be mechanosensitive (Albuschies & Vogel, 2013; Tsubouchi et al., 2012) in a way that can rely on Ca²⁺
signaling (as in microvilli) (Efremov et al., 2022). Thus, mechanosensation might be an ancestral property of the ancestral filopodia-like cellular appendages that gave rise to both metazoan microvilli and metazoan filopodia.

## Evolution of apical microvillar bundles in choanozoan ancestors

Only in Metazoa and Choanoflagellata are microvilli connected into bundles by inter-microvillar adhesions and do they have a polarized distribution in the cell. Inter-microvillar adhesions (allowing assembly of microvilli into bundles) and the restriction of microvilli to the apical pole thus might date back to the Urchonoazoan (Brunet & Booth, 2023; Brunet & King, 2017; Sebé-Pedrós et al., 2013), the last common ancestor of choanoflagellates and animals (Figure 4). Interestingly, a similar transition from a non-polarized to the polarized distribution of microvilli seems recapitulated during early embryonic development when blastomeres acquire apico-basal polarity (Box 3).

What might have been the ancestral morphology of microvillar bundles in the Urmetazoan? Microvilli are arranged as a collar complex (i.e., a ring surrounding a motile or non-motile cilium) in choanoflagellates and in many animal lineages, including (but not restricted to) sponges (Brunet & King, 2017; Rieger, 1976). This morphological similarity and the phylogenetic restriction of multiple intermicrovillar linkers to choanozoans (Figure 3) suggests that microvilli in the last metazoan and choanozoan common ancestors might have already been organized into a collar complex (Figure 4). Collar complex homology is accepted by many authors (Brunet & King, 2017; Colgren & Burkhardt, 2022; Nielsen, 2008) but has sometimes been questioned due to differences in structure and function between choanocytes and choanoflagellates, such as the opening angle of the collar (Mah et al., 2014) or the lack of connection between the flagellar root and the nucleus (which is absent in choanoflagellates but often present in choanocytes) (Pozdnyakov et al., 2017). However, these features can vary within choanoflagellates and/or choanocytes themselves: for example, the opening angle of the choanoflagellate collar is species-dependent and can even be behaviorally dynamic (Brunet et al., 2019), and the flagellar root-nucleus connection has been lost in the choanocytes of multiple sponges (Pozdnyakov et al., 2017). Another observation sometimes used to argue against collar complex homology is the fact that choanoflagellate genomes encode numerous genes whose sponge homologs are not expressed in choanocytes, but in other sponge cell types instead (such as amoeboid cells or stem cells) (Sogabe et al., 2019). However, choanoflagellate genomes nonethless still encode about 1000 gene families specifically

expressed in choanocytes, and the presence of non-shared families is unsurprising given that choanoflagellates are capable of functions choanocytes do not perform, such as amoeboid mobility (Brunet et al., 2021). Overall, comparative data are thus consistent with the homology of the collar. However, evolutionary convergence remains possible, and a better reconstitution of the evolution of the collar complex will require molecular characterization.

## From collar cells to enterocytes, sensory cells, and excretory cells in animal diversification

Given their localization at the plasma membrane and the large surface they provide, microvilli appear especially amenable to repurposing for the detection, capture, or uptake of a broad range of signals and molecules. In line with subfunctionalization of cell types during animal evolution, diverse types of specialized microvilli have evolved via both gain and loss of proteins from the ancestral microvillar core components. A leitmotiv seems to have been subfunctionalization, in accordance with “division of labor” models of cell type evolution, in which different cell types retain distinct subsets of functions from ancestral plurifunctional cells (Arendt, 2008; Arendt et al., 2016). This applies to microvilli specialized in feeding or mechanosensation, for example. However, functional innovation also played a part: the staircase organization of stereocilia, for example, represents a clear vertebrate innovation (Fritzsch & Straka, 2014). Thus, the evolution of microvillated cell types represents an interesting case study in the principles of cell type evolution.

Microvilli arranged as a collar complex around a flagellum occur across the metazoan tree of life in various cell types involved in feeding (Lavrov et al., 2022), excretion (Arendt, 2021), or sensation (Fritzsch & Straka, 2014). Other types of microvillar arrangement can be plausibly interpreted as modified collar complexes: for example, BB microvilli have been observed in multiple bilaterians and likely represent an Urbilaterian feature (Hartenstein & Martinez, 2019). In contrast to the enteroocyte BB, the renal tubule BB can display a single cilium (Weinbaum et al., 2010), and its formation depends on centrosome microtubules (Tonucci et al., 2018), emphasizing the functional connection between microvilli and microtubules.

The diverse stereocilia/kinocilium arrangements of chordate hair cells might have evolved from an ancestral collar complex (Caicci et al., 2007; Fritzsch & Straka, 2014), as suggested by hair cell development. During this process, microvilli are originally organized in a circle around the kinocilium before the kinocilium migrates to an eccentric position (Fritzsch & Straka, 2014). This pro-
Box 3: Blastomere microvilli and the establishment of cell polarity

Blastomeres, the first cells formed by cleavage of the zygote, are covered by microvilli in mammalian embryos (Ducibella et al., 1977). During the first rounds of mouse cell division, blastomere microvilli have a non-polarized distribution but they become apical at the eight-cell stage (Li et al., 2011). As a result, microvilli are only found at the cellular pole that faces the external environment (Lamba & Zernicka-Goetz, 2023), like in epithelial cells. This polarization is independent of microtubules; in contrast, microtubules are organized by the apical domain once it is established (Lamba & Zernicka-Goetz, 2023). Although it is currently unknown if microvilli directly participate in microtubule polarization, this further highlights the relation between microvilli distribution, microtubule network, and apico-basal cell polarity (Brunet & Booth, 2023). The precise function of blastomere microvilli is uncertain, but the cells displaying microvilli become the first differentiated cells in the mouse embryo. They form the trophoblast lineage which later gives rise to the syncytiotrophoblast (Staun-Ram & Shalev, 2005). Syncytiotrophoblast microvilli are essential for embryo implementation (Staun-Ram & Shalev, 2005) and placental nutrient exchange (Jansson et al., 2009). Like other apical microvilli, blastomere microvilli require ERM proteins (Lamba & Zernicka-Goetz, 2023) and are associated with cilia in certain bryozoans (Nekliudova et al., 2019). Outside mammals, blastomere microvilli have not been functionally characterized in detail but have nonetheless been reported in diverse phyla. For example, the microvilli of sea urchin eggs and early embryos have been proposed to mediate adhesion of the blastula to the hyaline layer (Wolpert & Mercer, 1963). In Porifera, microvilli-like protrusions cover the blastomeres but are referred to as filopodia (Leys et al., 2006). The evolution of blastomere microvilli is difficult to reconstitute as their molecular composition is poorly known.

cess is regulated by the PCP pathway, which is unique
to animals (Wright et al. 2022). Comparative data also
suggest an affinity between hair cells and collar cells:
mechanosensory cells in other bilaterians and cnidari-
ans often sport both a cilium with microvilli, frequently
arranged in a collar-like morphology (Bezares-Calderón
et al., 2018, 2019).

Microvillar photosensation relies on the R-opsin family of photoreceptors, whose origin has been traced back to the common ancestor of the Cnidaria and Bilateria, although microvillar photoreceptors have been lost in vertebrates (Lamb, 2013). Interestingly, during rhabdomere formation, a cilium can be transiently present in certain arthropods (Lamb, 2013). Reciprocally, during the development of the ciliary photoreceptors of vertebrates, microvilli can be present around the cilium in the form of calyceal process (as described in zebrafish [Miles et al., 2021], frogs and primates [Sahly et al., 2012]). Because the molecular composition of the calyceal processes is close to that of stereocilia (Sharkova et al., 2022) and since they surround a cilium, it has been hypothesized that they evolved from a collar complex organization. More generally, the currently predominant hypothesis is that both microvillar and ciliary photoreceptors evolved from an ancestral photoreceptor cell type with an ancestral opsin in the common ancestor of the Placozoa, Cnidaria, and Bilateria (Lamb, 2013). This ancestral photoreceptor would have had both cilium and microvilli (as do the photoreceptors of some cnidarian larvae [Nordström et al., 2003]), possibly arranged in a collar complex.

Finally, microvilli specialized in chemosensation and olfaction are found in various cell types with potentially distinct evolutionary history. Olfactosensory neurons (OSNs) with microvilli are conserved across vertebrates (Poncelet & Shimeld, 2020). Interestingly, actinopterygians and birds can have OSNs with both cilia and microvilli (Eisthen, 1992; Hansen & Finger, 2000) which might again reflect an evolutionary origin from collar cells. In tunicates and cephalochordates, OSNs-like cells with both cilia and microvilli have been described but it is unclear if those cells are homologous to those of vertebrates (Poncelet & Shimeld, 2020). Finally, chemosensory tuft cells have only been described in vertebrates and their evolutionary origin remains uncertain (Iqbal et al., 2023).

Thus, many microvillated cell types can be tenta-
tively traced back to an ancestral collar cell morphology.
This is consistent with the hypothesis that a collar
complex was present in the last metazoan common
ancestor, which it might have been inherited from a
common ancestor with choanoflagellates, although it
could also have evolved independently in metazoans
and choanoflagellates (as discussed in the previous
section).

CONCLUSION AND PERSPECTIVES

The definition of microvilli has often been relatively
implicit, at the risk of bringing about ambiguities in
discussions of their origin and evolution. Here, we pro-
pose an explicit structural definition. Microvilli likely
evolved from pre-existent cellular protrusions similar
to filopodia (Sebé-Pedrós et al., 2013). This implies there likely existed (and might still exist) a continuum between filopodia and microvilli. Indeed, some actin-filled protrusions lie between these two extremes, such as T-cell microvilli or the protrusions observed in *M. vibrans* (Mylnikov et al., 2019)—making their classification somewhat subjective.

Our HMM-based phylogenetic survey allowed us to identify microvillar homolog proteins in more Amorphea branches than earlier studies (Peña et al., 2016; Sebé-Pedrós et al., 2013). Notably, we could find homologs of almost all microvillar proteins we investigated in Choanoflagellata and Filasterea (Figure 3), which strengthens the hypothesis of homology of microvilli across Filozoa. Furthermore, the concomitant evolution of many intermicrovillar linkers and of the collar complex organization at the stem of Choanozoa is consistent with the homology of the collar complex. Surprisingly, homologs for Fascin, Espin, Harmonin family, and GRCR1 were each detected in a (different) single species outside Filozoa (Figure 3, Figure S1). Although these homologs were validated by protein domain analysis and occupied a coherent position in gene phylogenies, their isolated occurrence remains puzzling. In the future, more detailed investigations might clarify the nature and history of these distant apparent homologs.

We outlined in the last part of this review a potential scenario for microvilli diversification during animal evolution (Figure 4). In this hypothesis, microvilli first acquired a polarized distribution by becoming restricted to the apical pole of the cell, became organized into a collar complex thanks to intermicrovillar adhesions, and then diversified into a range of specialized microvilli across animal cell types. Additional research will be necessary to test, complete, and amend this scenario. Most studies so far have revolved around classical model systems (vertebrates, flies, and nematodes) and it is thus possible that essential and conserved microvillar proteins across other Choanozoa are undiscovered. Moreover, homologs of many bilaterian microvillar proteins are present in non-bilaterians and unicellular relatives of animals, but their location and function are generally unknown. Structural and functional characterization of microvilli across non-bilaterian animals, choanoflagellates, and filastereans might shed light on those uncertainties.

Finally, an open question is the problem of the genetic specification of microvilli—are there specific transcription factors, or gene regulatory networks, orchestrating the microvillar biogenesis program (similar to Rfx and FoxJ1 for ciliogenesis [Coyle et al., 2023])? This question has not been extensively addressed but recent studies highlighted some transcription factors important for BB microvilli or stereocilia development, including HNF4, KLF5, ATOH1, SRF, MRTFB, POU4F3, and GFI1

(Bell et al., 2013; Chen et al., 2021; Wang et al., 2024; Zhou et al., 2023). The elucidation and comparison of the gene regulatory networks for the different types of microvilli represents an exciting prospect and will likely bring a new dimension to our understanding of microvilli origin and evolution.

## ACKNOWLEDGMENTS

We thank Christine Miege, Pawel Burkhardt, Nicolas Goudemand, Diede De Haan, and three anonymous reviewers for helpful comments on the manuscript. Work in Thibaut Brunet's lab is supported by the Institut Pasteur (G5 package), the CNRS (UMR 3691), and the ERC Starting Grant EvoMorphoCell (Grant agreement ID: 101040745). Mylan Ansel is supported by the Ecole Normale Supérieure de Lyon (allocation élève normalien). Gautam Dey and Kaustubh Ramachandran acknowledge the support of the European Molecular Biology Laboratory (EMBL). Gautam Dey is funded by the European Union (ERC, KaryodynEVO, 101078291). Kaustubh Ramachandran is part of a collaboration for a joint PhD degree between EMBL and Heidelberg University, Faculty of Biosciences, Germany. Kaustubh Ramachandran holds an Add-on Fellowship for Interdisciplinary Life Science from the Joachim Herz Stiftung.

## CONFLICT OF INTEREST STATEMENT

The authors declare no conflicts of interest.

## DATA AVAILABILITY STATEMENT

All raw data generated for this manuscript are available on FigShare https://figshare.com/articles/online_resource/Origin_and_evolution_of_microvilli/25828882/2 (Ansel et al. 2024). Code used for the analysis is available on GitHub: https://github.com/kausthubhr/microvilli phylogenetics (Ramachandran et al. 2024).

## ORCID

Mylan Ansel https://orcid.org/0009-0003-5650-5956

Kaustubh Ramachandran https://orcid.org/0000-0002-6221-8166

Thibaut Brunet https://orcid.org/0000-0002-1843-1613

## REFERENCES

Abraham, M., Hilge, V., Lison, S. & Tibika, H. (1984) The cellular envelope of oocytes in teleosts. *Cell and Tissue Research*, 235(2), 403–410. https://doi.org/10.1007/BF00217866

Albuschies, J. & Vogel, V. (2013) The role of filopodia in the recognition of nanotopographies. *Scientific Reports*, 3(1), 1658. https://doi.org/10.1038/srep01658

Amiel, A.R., Michel, V., Carvalho, J.E., Shkreli, M., Petit, C. & Röttinger, E. (2021) L'anémone de mer Nematostella vectensis. *Médecine/Sciences*, 37(2), 167–177. https://doi.org/10.1051/medsci/2020282
Anakina, R.P. & Drozdov, A.L. (2001) Gamete structure and fertilization in the Barents sea sponge Leucosolenia complicata. *Russian Journal of Marine Biology*, 27(3), 143–150. https://doi.org/10.1023/A:1016761317637

Ansel, M., Ramachandran, K., Dey, G. & Brunet, T. (2024) Origin and evolution of microvilli. fish. Online resource. https://doi.org/10.6084/M9.FIGSHARE.25828882.V2

Arendt, D. (2008) The evolution of cell types in animals: emerging principles from molecular studies. *Nature Reviews. Genetics*, 9(11), 868–882. https://doi.org/10.1038/nrg2416

Arendt, D. (2021) Animal evolution: of flame and collar cells. *Current Biology*, 31(16), R1003–R1006. https://doi.org/10.1016/j.cub.2021.07.006

Arendt, D., Musser, J.M., Baker, C.V.H., Bergman, A., Cepko, C., Erwin, D.H., Pavlicev, M., et al. (2016) The origin and evolution of cell types. *Nature Reviews Genetics*, 17(12), 744–757. https://doi.org/10.1038/nrg.2016.127

Arendt, D., Tessmar-Raible, K., Snyman, H., Dorresteijn, A.W. & Wittbrodt, J. (2004) Ciliary photoreceptors with a vertebrate-type opsin in an invertebrate brain. *Science*, 306(5697), 869–871. https://doi.org/10.1126/science.1099955

Bell, S.M., Zhang, L., Xu, Y., Besnard, V., Wert, S.E., Shroyer, N. & Whitsett, J.A. (2013) Kruppel-like factor 5 controls villus formation and initiation of cytokdifferentiation in the embryonic intestinal epithelium. *Developmental Biology*, 375(2), 128–139. https://doi.org/10.1016/j.ydbio.2012.12.010

Benesh, A.E., Nambiar, R., McConnell, R.E., Mao, S., Tabb, D.L. & Tyska, M.J. (2010) Differential localization and dynamics of class I myosins in the enterocyte microvillus. *Molecular Biology of the Cell*, 21(6), 970–978. https://doi.org/10.1091/mbc.e09-070638

Bernadskaya, Y.Y., Patel, F.B., Hsu, H.-T. & Soto, M.C. (2011) Arp2/3 promotes junction formation and maintenance in the caenorhabditis elegans intestine by regulating membrane association of apical proteins. *Molecular Biology of the Cell*, 22(16), 2886–2899. https://doi.org/10.1091/mbc.e10-10-0862

Bezares-Calderón, L.A., Berger, J., Jasek, S., Verasztó, C., Mendes, S., Gühmann, M., Almeda, R., Shahidi, R. & Jékely, G. (2018) Neural circuitry of a polycystin-mediated hydrodynamic startle response for predator avoidance. *eLife*, 7(December), e36262. https://doi.org/10.7554/eLife.36262

Bezares-Calderón, L.A., Berger, J. & Jékely, G. (2019) Diversity of cilia-based mechanosensory systems and their functions in marine animal behaviour. *Philosophical Transactions of the Royal Society B: Biological Sciences*, 375(1792), 20190376. https://doi.org/10.1098/rstb.2019.0376

Bidaud-Meynard, A., Demouchy, F., Nicolle, O., Pacquelet, A., Suman, S.K., Plancke, C.N., Robin, F.B. & Michaux, G. (2021) High-resolution dynamic mapping of the C. elegans intestinal brush border. *Development*, 148(23), dev200029. https://doi.org/10.1242/dev.200029

Blanco-Sánchez, B., Clément, A., Fierro, J., Stednitz, S., Phillips, J.B., Wegner, J., Panlilio, J.M., Peirce, J.L., Washbourne, P. & Westerfield, M. (2018) Grxcr1 promotes hair bundle development by destabilizing the physical interaction between harmonin and sans usher syndrome proteins. *Cell Reports*, 25(5), 1281–1291.e4. https://doi.org/10.1016/j.celrep.2018.10.005

Bornschlögl, T. (2013) How filopodia pull: what we know about the mechanics and dynamics of filopodia. *Cytoskeleton*, 70(10), 590–603. https://doi.org/10.1002/cm.21130

Brett, C.L., Donowitz, M. & Rao, R. (2005) Evolutionary origins of eukaryotic sodium/proton exchangers. *American Journal of Physiology-Cell Physiology*, 288(2), C223–C239. https://doi.org/10.1152/ajpcell.00360.2004

Brunet, T., Albert, M., Roman, W., Coyle, M.C., Spitzer, D.C. & King, N. (2021) A flagellate-to-amoeboid switch in the closest living relatives of animals. Edited by P J Wittkopp, I Ruiz-Trillo & P López-García. *eLife*, 10, e61037. https://doi.org/10.7554/eLife.61037

Brunet, T. & Booth, D.S. (2023) Cell polarity in the protist-to-animal transition. In: *Current topics in developmental biology*, S0070215323000455. Cambridge, MA, United States: Academic Press/Elsevier. https://doi.org/10.1016/bs.ctdb.2023.03.001

Brunet, T. & King, N. (2017) The origin of animal multicellularity and cell differentiation. *Developmental Cell*, 43(2), 124–140. https://doi.org/10.1016/j.devcel.2017.09.016

Brunet, T. & King, N. (2022) The single-celled ancestors of animals: a history of hypotheses. In: *The evolution of multicellularity*, 1st edition, Boca Raton, FL, United States: CRC Press. https://www.taylorfrancis.com/chapters/oa-edit/10.1201/9780429351907-17/single-celled-ancestors-animals-thibaut-brunet-nicole-king?context=ubx&refId=78e592ca-1fe9-42d1-a7c8-2d5aa0897f2c

Brunet, T., Larson, B.T., Linden, T.A., Vermeij, M.J.A., McDonald, K. & King, N. (2019) Light-regulated collective contractility in a multicellular choanoflagellate. *Science*, 366(6463), 326–334. https://doi.org/10.1126/science.aay2346

Burighel, P., Lane, N.J., Fabio, G., Stefano, T., Zaniolo, G., Carnevali, M.D.C. & Manni, L. (2003) Novel, secondary sensory cell organ in ascidians: in search of the ancestor of the vertebrate lateral line. *Journal of Comparative Neurology*, 461(2), 236–249. https://doi.org/10.1002/cne.10666

Caberlotto, E., Michel, V., Foucher, I., Bahloul, A., Goodyear, R.J., Pepermans, E., Michalski, N., et al. (2011) Usher type 1G protein sans is a critical component of the tip-link complex, a structure controlling actin polymerization in stereocilia. *Proceedings of the National Academy of Sciences*, 108(14), 5825–5830. https://doi.org/10.1073/pnas.1017114108

Cai, E., Marchuk, K., Beemiller, P., Bepler, C., Rubashkin, M.G., Weaver, V.M., Gérard, A., et al. (2017) Visualizing dynamic microvillar search and stabilization during ligand detection by T cells. *Science*, 356 (6338), eaal3118. https://doi.org/10.1126/science.aal3118

Caicci, F., Burighel, P. & Manni, L. (2007) Hair cells in an ascidian (tunicata) and their evolution in chordates. *Hearing Research*, 231(1), 63–72. https://doi.org/10.1016/j.heares.2007.05.007

Carlton, A.J., Halford, J., Underhill, A., Jeng, J.-Y., Avenarius, M.R., Gilbert, M.L., Ceriani, F., et al. (2021) Loss of *Baiap2l2* destabilizes the transducing stereocilia of cochlear hair cells and leads to deafness. *The Journal of Physiology*, 599(4), 1173–1198. https://doi.org/10.1113/JP280670

Carré, D., Rouvière, C. & Sardet, C. (1991) In vitro fertilization in ctenophores: sperm entry, mitosis, and the establishment of bilateral symmetry in Beroe ovata. *Developmental Biology*, 147(2), 381–391. https://doi.org/10.1016/0012-1606(91)90296-F

Cavalier-Smith, T. & Chao, E.E. (2010) Phylogeny and evolution of apusomonadida (protozoa: apusozoa): new genera and species. *Protist*, 161(4), 549–576. https://doi.org/10.1016/j.protis.2010.04.002

Cebecauer, M. (2021) Role of lipids in morphogenesis of T-cell microvilli. *Frontiers in Immunology*, 12, 613591. https://doi.org/10.3389/fimmu.2021.613591

Chen, L., Luo, S., Dupre, A., Vasoya, R.P., Parthasarathy, A., Aita, R., Malhotra, R., et al. (2021) The nuclear receptor HNF4 drives a brush border gene program conserved across murine intestine, kidney, and embryonic yolk sac. *Nature Communications*, 12(1), 2886. https://doi.org/10.1038/s41467-021-22761-5

Chugh, P. & Paluch, E.K. (2018) The actin cortex at a glance. *Journal of Cell Science*, 131(14), jcs186254. https://doi.org/10.1242/jcs.186254

Clark, W.H. & Dewel, W.C. (1974) The structure of the gonads, game-togenesis, and sperm-egg interactions in the anthozoa. *American Zoologist*, 14(2), 495–510. https://doi.org/10.1093/icb/14.2.495

Cohen, E., Sawyer, J.K., Peterson, N.G., Dow, J.A.T. & Fox, D.T. (2020) Physiology, development, and disease modeling in the drosophila excretory system. *Genetics*, 214(2), 235–264. https://doi.org/10.1534/genetics.119.302289
Colgren, J. & Burkhardt, P. (2022) The premetazoan ancestry of the synaptic toolkit and appearance of first neurons. *Essays in Biochemistry*, 66(6), 781-795. https://doi.org/10.1042/EBC20220042

Condeelis, J. (1993) Understanding the cortex of crawling cells: insights from dictyostelium. *Trends in Cell Biology*, 3(11), 371-376. https://doi.org/10.1016/0962-8924(93)90085-F

Courjaret, R., Hodeify, R., Hubrack, S., Ibrahim, A., Dib, M., Daas, S. & Machaca, K. (2016) The Ca2+-activated Cl⁻ channel Ano1 controls microvilli length and membrane surface area in the oocyte. *Journal of Cell Science*, 129(13), 2548-2558. https://doi.org/10.1242/jcs.188367

Coyle, M.C., Tajima, A.M., Leon, F., Choksi, S.P., Yang, A., Espinoza, S., Hughes, T.R., Reiter, J.F., Booth, D.S. & King, N. (2023) An RFX transcription factor regulates ciliogenesis in the closest living relatives of animals. *Current Biology: CB*, 33(17), 3747-3758.e9. https://doi.org/10.1016/j.cub.2023.07.022

Danielsen, M. & Hansen, G. (2003) Lipid rafts in epithelial brush borders: atypical membrane microdomains with specialized functions. *Biochim Biophys Acta*, 1617(1-2), 1-9. https://doi.org/10.1016/j.bbamem.2003.09.005

Dayel. (2008) Choanoflagellates and animal multicellularity. Online resource. Accessed on December 14, 2008. http://www.dayel.com/choanoflagellates/

Derderian, C., Canales, G.I. & Reiter, J.F. (2023) Seriously cilia: a tiny organelle illuminates evolution, disease, and intercellular communication. *Developmental Cell*, 58(15), 1333-1349. https://doi.org/10.1016/j.devcel.2023.06.013

DeRosier, D.J. (2000) F-actin bundles are derivatives of microvilli: what does this tell us about how bundles might form? *The Journal of Cell Biology*, 148(1), 1-6. https://doi.org/10.1083/jcb.148.1.1

Dooley, S.A., Engevik, K.A., Digrazia, J., Stubler, R., Kaji, I., Krystofiak, E. & Engevik, A.C. (2022) Myosin 5b is required for proper localization of the interm微villar adhesion complex in the intestinal brush border. *American Journal of Physiology-Gastrointestinal and Liver Physiology*, 323 (5): G501-G510. https://doi.org/10.1152/ajpgi.00212.2022

Du, Y., Gao, H., He, C., Xin, S., Wang, B., Zhang, S., Gong, F., et al. (2023) An update on the biological characteristics and functions of tuft cells in the gut. *Frontiers in Cell and Developmental Biology*, 10(January), 1102978. https://doi.org/10.3389/fcell.2022.1102978

Duan, Y., Gotoh, N., Yan, Q., Du, Z., Weinstein, A.M., Wang, T. & Weinbaum, S. (2008) Shear-induced reorganization of renal proximal tubule cell actin cytoskeleton and apical junctional complexes. *Proceedings of the National Academy of Sciences*, 105(32), 11418-11423. https://doi.org/10.1073/pnas.0804954105

Ducibella, T., Ukena, T., Karnovsky, M. & Anderson, E. (1977) Changes in cell surface and cortical cytoplasmic organization during early embryogenesis in the preimplantation mouse embryo. *The Journal of Cell Biology*, 74(1), 153-167. https://doi.org/10.1083/jcb.74.1.153

Dumontier, M., Höcht, P., Mintert, U. & Faix, J. (2000) Rac1 GTPases control filopodia formation, cell motility, endocytosis, cytokinesis and development in dictyostelium. *Journal of Cell Science*, 113(12), 2253-2265. https://doi.org/10.1242/jcs.113.12.2253

Dunbar, L.A., Patni, P., Aguilar, C., Mburu, P., Corns, L., Wells, H.R., Delmaghani, S., et al. (2019) Clarin-2 is essential for hearing by maintaining stereocilia integrity and function. *EMBO Molecular Medicine*, 11(9), e10288. https://doi.org/10.15252/emmm.201910288

Eckelbarger, K.J. & Larson, R. (1992) Ultrastructure of the ovary and oogenesis in the jellyfish *Linuche unguiculata* and *Stomolophus meleagris*, with a review of ovarian structure in the scyphozoa. *Marine Biology*, 114(4), 633-643. https://doi.org/10.1007/BF00357260

Eddy, S.R. (2023) HMMER user's guide. Online resource. http://eddylab.org/software/hmmer/Userguide.pdf

Efremov, A.K., Yao, M., Sun, Y., Han Tee, Y., Sheetz, M.P., Bershadsky, A.D., Martinac, B. & Yan, J. (2022) Application of piconewton forces to individual filopodia reveals mechanosensory role of L-type Ca2+

channels. *Biomaterials*, 284(May), 121477. https://doi.org/10.1016/j.biomaterials.2022.121477

Eisthen, H.L. (1992) Phylogeny of the vomeronasal system and of receptor cell types in the olfactory and vomeronasal epithelia of vertebrates. *Microscopy Research and Technique*, 23(1), 1-21. https://doi.org/10.1002/jemt.1070230102

El-Amraoui, A. & Petit, C. (2005) Usher I syndrome: unravelling the mechanisms that underlie the cohesion of the growing hair bundle in inner ear sensory cells. *Journal of Cell Science*, 118(20), 4593-4603. https://doi.org/10.1242/jcs.02636

Erives, A. & Fritzsch, B. (2020) A screen for gene paralogies delineating evolutionary branching order of early metazoa. *G3 Genes|Genomes|Genetics*, 10(2), 811-826. https://doi.org/10.1534/g3.119.400951

Fain, G.L., Hardie, R. & Laughlin, S.B. (2010) Phototransduction and the evolution of photoreceptors. *Current Biology*, 20(3), R114-R124. https://doi.org/10.1016/j.cub.2009.12.006

Figard, L. & Sokac, A.M. (2014) A membrane reservoir at the cell surface. *Bioarchitecture*, 4(2), 39-46. https://doi.org/10.4161/bioa.29069

Franzen, W. (1988) Oogenesis and larval development of *Scypha ciliata* (Porifera, Calcarea). *Zoomorphology*, 107(6), 349-357.

Fritz-Laylin, L.K., Riel-Mehan, M., Chen, B.-C., Lord, S.J., Goddard, T.D., Ferrin, T.E., Nicholson-Dykstra, S.M., et al. (2017) Actin-based protrusions of migrating neutrophils are intrinsically lamellar and facilitate direction changes. *eLife*, 6(September), e26990. https://doi.org/10.7554/eLife.26990

Frittsch, B. & Straka, H. (2014) Evolution of vertebrate mechanosensory hair cells and inner ears: toward identifying stimuli that select mutation driven altered morphologies. *Journal of Comparative Physiology A: Neuroethology, Sensory, Neural, and Behavioral Physiology*, 200(1), 5-18. https://doi.org/10.1007/s00359-013-0865-z

Furutani, Y., Matsuno, H., Kawasaki, M., Sasaki, T., Mori, K. & Yoshihara, Y. (2007) Interaction between telencephalin and ERM family proteins mediates dendritic filopodia formation. *The Journal of Neuroscience*, 27(33), 8866-8876. https://doi.org/10.1523/JNEUROSCI.1047-07.2007

Galindo, L.J., López-García, P., Torruella, G., Karpov, S. & Moreira, D. (2021) Phylogenomics of a new fungal phylum reveals multiple waves of reductive evolution across Holomycota. *Nature Communications*, 12(1), 4973. https://doi.org/10.1038/s41467-021-25308-w

Geisler, F., Coch, R.A., Richardson, C., Goldberg, M., Denecke, B., Bossinger, O. & Leube, R.E. (2019) The intestinal intermediate filament network responds to and protects against microbial insults and toxins. *Development*, 146(2), dev169482. https://doi.org/10.1242/dev.169482

Geisler, F., Remmelzwaal, S., Jankowski, V., Schmidt, R., Boxem, M. & Leube, R.E. (2023) Intermediate filament network perturbation in the C. elegans intestine causes systemic dysfunctions. *eLife*, 12, e82333. https://doi.org/10.7554/eLife.82333

Genovese, F. & Tizzano, M. (2018) Microvillous cells in the olfactory epithelium express elements of the solitary chemosensory cell transduction signaling cascade. *PLoS ONE*, 13(9), e0202754. https://doi.org/10.1371/journal.pone.0202754

Ghosh, S., Di Bartolo, V., Tubul, L., Shimoni, E., Kartvelishvily, E., Dadosh, T., Feigelson, S.W., Alon, R., Alcover, A. & Haran, G. (2020) ERM-dependent assembly of T cell receptor signaling and co-stimulatory molecules on microvilli prior to activation. *Cell Reports*, 30(10), 3434-3447.e6. https://doi.org/10.1016/j.celrep.2020.02.069

Goode, B.L., Eskin, J. & Shekhar, S. (2023) Mechanisms of actin disassembly and turnover. *Journal of Cell Biology*, 222(12), e202309021. https://doi.org/10.1083/jcb.202309021

Gorelik, J., Shevchuk, A.I., Frolenkov, G.I., Diakonov, I.A., Lab, M.J., Kros, C.J., Richardson, G.P., et al. (2003) Dynamic assembly of surface structures in living cells. *Proceedings of the National Academy*
of Sciences of the United States of America, 100(10), 5819–5822.
https://doi.org/10.1073/pnas.1030502100

Granger, B. & Baker, R.F. (1950) Electron microscope investigation of the striated border of intestinal epithelium. *The Anatomical Record*, 107(4), 423–441. https://doi.org/10.1002/ar.1091070409

Grau-Bové, X., Torruella, G., Donachie, S., Suga, H., Leonard, G., Richards, T.A. & Ruiz-Trillo, I. (2017) Dynamics of genomic innovation in the unicellular ancestry of animals. *eLife*, 6(July), e26036. https://doi.org/10.7554/eLife.26036

Grega-Larson, N.E., Crawley, S.W., Erwin, A.L. & Tyska, M.J. (2015) Cordon bleu promotes the assembly of brush border microvilli. *Molecular Biology of the Cell*, 26(21), 3803–3815. https://doi.org/10.1091/mbc.E15-06-0443

Hadi, S., Alexander, A.J., Catalina Vélez-Ortega, A. & Frolenkov, G.I. (2020) Myosin-XVa controls both staircase architecture and diameter gradation of stereocilia rows in the auditory hair cell bundles. *JARO: Journal of the Association for Research in Otolaryngology*, 21(2), 121–135. https://doi.org/10.1007/s10162-020-00745-4

Hamann, E., Gruber-Vodicka, H., Kleiner, M., Tegetmeyer, H.E., Riedel, D., Littmann, S., Chen, J., et al. (2016) Environmental Breviatae harbour mutualistic Arcobacter epiobionts. *Nature*, 534(7606), 254–258. https://doi.org/10.1038/nature18297

Hammel, J.U. & Nickel, M. (2014) A new flow-regulating cell type in the Demosponge Tethya Wilhelma—functional cellular anatomy of a leuconoid canal system. *PLOS ONE*, 9(11), e113153. https://doi.org/10.1371/journal.pone.0113153

Han, L., Qu, Q., Aydin, D., Panova, O., Robertson, M.J., Xu, Y., Dror, R.O., Skiniotis, G. & Feng, L. (2022) Structure and mechanism of the SGLT family of glucose transporters. *Nature*, 601(7892), 274–279. https://doi.org/10.1038/s41586-021-04211-w

Hansen, A. & Finger, T.E. (2000) Phyletic distribution of crypt-type olfactory receptor neurons in fishes. *Brain Behavior and Evolution*, 55(2), 100–110. https://doi.org/10.1159/000006645

Hart, A., Gadegaard, N., Wilkinson, C.D.W., Oreffo, R.O.C. & Dalby, M.J. (2007) Osteoprogenitor response to low-adhesion nanotopographies originally fabricated by electron beam lithography. *Journal of Materials Science: Materials in Medicine*, 18(6), 1211–1218. https://doi.org/10.1007/s10856-007-0157-7

Hartenstein, V. & Martinez, P. (2019) Phagocytosis in cellular defense and nutrition: a food-centered approach to the evolution of macrophages. *Cell and Tissue Research*, 377(3), 527–47. https://doi.org/10.1007/s00441-019-03096-6

Heckman, C.A. & Plummer, H.K. (2013) Filopodia as sensors. *Cellular Signalling*, 25(11), 2298–2311. https://doi.org/10.1016/j.cellsig.2013.07.006

Hegan, P.S., Mermall, V., Tilney, L.G. & Mooseker, M.S. (2007) Roles for Drosophila melanogaster myosin IB in maintenance of enterocyte brush-border structure and resistance to the bacterial pathogen Pseudomonas entomophila. *Molecular Biology of the Cell*, 18(11), 4625–4636. https://doi.org/10.1091/mbc.E07-02-0191

Heger, P., Zheng, W., Rottmann, A., Panfilio, K.A. & Wiehe, T. (2020) The genetic factors of bilaterian evolution. *eLife*, 9, e45530. https://doi.org/10.7554/eLife.45530

Hirano, M., Das, S., Guo, P. & Cooper, M.D. (2011) The evolution of adaptive immunity in vertebrates. *Advances in Immunology*, 109:125–157. https://doi.org/10.1016/B978-0-12-387664-5.00004-2

Hirsch, D., Stahl, A. & Lodish, H.F. (1998) A family of fatty acid transporters conserved from mycobacterium to man. *Proceedings of the National Academy of Sciences*, 95(15), 8625–8629. https://doi.org/10.1073/pnas.95.15.8625

Houdusse, A. & Titus, M.A. (2021) The many roles of myosins in filopodia, microvilli and stereocilia. *Current Biology: CB*, 31(10), R586–R602. https://doi.org/10.1016/j.cub.2021.04.005

Iqbal, S., Karim, M.R., Yang, D.-C., Mathiyalagan, R. & Kang, S.C. (2023) Tuft cells—the immunological interface and role in disease regulation. *International Immunopharmacology*, 118(May), 110018. https://doi.org/10.1016/j.intimp.2023.110018

Jacob, F. (1996) *La logique du vivant: une histoire de l'hérédité*. Collection Tel 2. Paris: Gallimard.

James-Clark, H. (1867) IV.—Conclusive proofs of the animality of the ciliate sponges, and of their affinities with the Infusoria flagellata. *Journal of Natural History*, 19(109), 13–18.

Jansson, T., Myatt, L. & Powell, T. (2009) The role of trophoblast nutrient and ion transporters in the development of pregnancy complications and adult disease. *Current Vascular Pharmacology*, 7(4), 521–533. https://doi.org/10.2174/157016109789043982

Jia, B., De Peng Yuan, W.J.L., Xuan, Y.H. & Jeon, C.O. (2019) New insight into the classification and evolution of glucose transporters in the metazoa. *The FASEB Journal*, 33(6), 7519–7528. https://doi.org/10.1096/fj.201802617R

Jung, Y., Riven, I., Feigelson, S.W., Kartvelishvily, E., Tohya, K., Miyasaka, M., Alon, R. & Haran, G. (2016) Three-dimensional localization of T-cell receptors in relation to microvilli using a combination of superresolution micros. *Proceedings of the National Academy of Sciences*, 113(40), E5916–E5924. https://doi.org/10.1073/pnas.1605399113

Khurana, S. & George, S.P. (2008) Regulation of cell structure and function by actin-binding proteins: villin's perspective. *FEBS Letters*, 582(14), 2128–2139. https://doi.org/10.1016/j.febslet.2008.02.040

Kiela, P.R. & Ghishan, F.K. (2016) Physiology of intestinal absorption and secretion. *Best Practice & Research. Clinical Gastroenterology*, 30(2), 145–159. https://doi.org/10.1016/j.bpg.2016.02.007

Kim, H.-R. & Jun, C.-D. (2019) T cell microvilli: sensors or senders? *Frontiers in Immunology*, 10(July), 1753. https://doi.org/10.3389/fimmu.2019.01753

King, N. & Carroll, S.B. (2001) A receptor tyrosine kinase from choanoflagellates: molecular insights into early animal evolution. *Proceedings of the National Academy of Sciences*, 98(26), 15032–15037. https://doi.org/10.1073/pnas.261477698

Kozlov, A.S., Baumgart, J., Risler, T., Versteegh, C.P.C. & Hudspeth, A.J. (2011) Forces between clustered stereocilia minimize friction in the ear on a subnanometre scale. *Nature*, 474(7351), 376–379. https://doi.org/10.1038/nature10073

Kress, H., Stelzer, E.H.K., Holzer, D., Buss, F., Griffiths, G. & Rohrbach, A. (2007) Filopodia act as phagocytic tentacles and pull with discrete steps and a load-dependent velocity. *Proceedings of the National Academy of Sciences*, 104(28), 11633–11638. https://doi.org/10.1073/pnas.0702449104

Krey, J.F., Chatterjee, P., Halford, J., Cunningham, C.L., Perrin, B.J. & Barr-Gillespie, P.G. (2023) Control of stereocilia length during development of hair bundles. *PLOS Biology*, 21(4): e3001964. https://doi.org/10.1371/journal.pbio.3001964

Kummer, W. & Deckmann, K. (2017) Brush cells, the newly identified gatekeepers of the urinary tract. *Current Opinion in Urology*, 27(2), 85–92. https://doi.org/10.1097/MOU.00000000000000361

Kurima, K., Ebrahim, S., Pan, B., Sedlacek, M., Sengupta, P., Millis, B.A., Cui, R., et al. (2015) TMC1 and TMC2 localize at the site of mechanotransduction in mammalian inner ear hair cell stereocilia. *Cell Reports*, 12(10), 1606–1617. https://doi.org/10.1016/j.celrep.2015.07.058

Kyykallio, H., Oikari, S., Bueno Álvez, M., Dodd, C.J.G., Capra, J. & Rilla, K. (2020) The density and length of filopodia associate with the activity of hyaluronan synthesis in tumor cells. *Cancers*, 12(7), 1908. https://doi.org/10.3390/cancers12071908

Laishram, J., Avossa, D., Shahapure, R. & Torre, V. (2009) Mechanical computation in neurons. *Developmental Neurobiology*, 69(11), 731–751. https://doi.org/10.1002/dneu.20733

Lamb, T.D. (2013) Evolution of phototransduction, vertebrate photoreceptors and retina. *Progress in Retinal and Eye Research*, 36(September), 52–119. https://doi.org/10.1016/j.preteyeres.2013.06.001

Lamba, A. & Zernicka-Goetz, M. (2023) Chapter six—the role of polarization and early heterogeneities in the mammalian first cell fate decision. *Current topics in developmental biology* vol. 154, 169–196 https://doi.org/10.1016/bs.ctdb.2023.02.006
Lange, K. (2011) Fundamental role of microvilli in the main functions of differentiated cells: outline of an universal regulating and signaling system at the cell periphery. *Journal of Cellular Physiology*, 226(4), 896-927. https://doi.org/10.1002/jcp.22302

Lavrov, A.I., Bolshakov, F.V., Tokina, D.B. & Ereskovsky, A.V. (2022) Fine details of the choanocyte filter apparatus in asconoid careous sponges (Porifera: Calcarea) revealed by ruthenium red fixation. *Zoology*, 150(February), 125984. https://doi.org/10.1016/j.zool.2021.125984

Leadbeater, B.S.C. & Carr, M. (2022) Significance of the nudiform and tectiform modes of silica deposition, lorica assembly and cell division in choanoflagellates as exemplified by *Stephanoeca Diplocostata* and *Enibas Spp. Protist*, 173(6), 125923. https://doi.org/10.1016/j.protis.2022.125923

Leadbeater, B.S.C. (1983) Life-history and ultrastructure of a new marine species of Proterospongia (Choanoflagellida). *Journal of the Marine Biological Association of the United Kingdom*, 63(01), 135-160.

Leijnse, N., Barooji, Y.F., Arastoo, M.R., Sønder, S.L., Verhagen, B., Wullkopf, L., Erler, J.T., et al. (2022) Filopodia rotate and coil by actively generating twist in their actin shaft. *Nature Communications*, 13(1), 1636. https://doi.org/10.1038/s41467-022-28961-x

Leon, F., Espinoza-Esparza, J.M., Deng, V., Coyle, M.C., Espinoza, S. & Booth, D.S. (2024) Cell-type-specific expression of a DCYTB ortholog enables the choanoflagellate *Salpingoeca Rosetta* to utilize ferric colloids. *biorXiv*. https://doi.org/10.1101/2024.05.25.595918

Levin, T.C. & King, N. (2013) Evidence for sex and recombination in the choanoflagellate *Salpingoeca rosetta*. *Current Biology*, 23(21), 2176-2180. https://doi.org/10.1016/j.cub.2013.08.061

Leys, S.P., Cheung, E. & Boury-Esnault, N. (2006) Embryogenesis in the glass sponge *Oopsacas minuta*: formation of syncytia by fusion of blastomeres. *Integrative and Comparative Biology*, 46(2), 104-117. https://doi.org/10.1093/icb/icj016

Leys, S.P. & Hill, A. (2012) The physiology and molecular biology of sponge tissues. *Advances in Marine Biology*, 62, 1-56. https://doi.org/10.1016/B978-0-12-394283-8.00001-1

Li, C.-B., Wang, Z.-D., Zheng, Z., Hu, L.-L., Zhong, S.-Q. & Lei, L. (2011) Number of blastomeres and distribution of microvilli in cloned mouse embryos during compaction. *Zygote*, 19(3), 271-276. https://doi.org/10.1017/S0967199410000377

Liang, X., Qiu, X., Dionne, G., Cunningham, C.L., Pucak, M.L., Peng, G., Kim, Y.-H., Lauer, A., Shapiro, L. & Müller, U. (2021) CIB2 and CIB3 are auxiliary subunits of the mechanotransduction channel of hair cells. *Neuron*, 109(13), 2131-2149.e15. https://doi.org/10.1016/j.neuron.2021.05.007

Liao, I.J.-Y., Lu, T.-M., Chen, M.-E. & Luo, Y.-J. (2023) Spiralian genomics and the evolution of animal genome architecture. *Briefings in Functional Genomics*, 22(6), 498-508. https://doi.org/10.1093/bfgp/elad029

Lim, K.B., Bu, W., Goh, W.I., Koh, E., Ong, S.H., Pawson, T., Sudhaharan, T. & Ahmed, S. (2008) The Cdc42 effector IRSp53 generates filopodia by coupling membrane protrusion with actin dynamics. *Journal of Biological Chemistry*, 283(29), 20454-20472. https://doi.org/10.1074/jbc.M710185200

Liu, L., Wang, K., Liu, J., Wei, Y., Liu, W., Zhang, P., Hu, J. & Li, B. (2019) Starvation effect on the morphology of microvilli in HeLa cells. *Biochemical and Biophysical Research Communications*, 514(4), 1238-1243. https://doi.org/10.1016/j.bbrc.2019.05.073

Longo, F.J. (1987) Actin-plasma membrane associations in mouse eggs and oocytes. *Journal of Experimental Zoology*, 243(2), 299-309. https://doi.org/10.1002/jez.1402430215

Lorente-Cánovas, B., Eckrich, S., Lewis, M.A., Johnson, S.L., Marcotti, W. & Steel, K.P. (2022) Grxcr1 regulates hair bundle morphogenesis and is required for normal mechanoelectrical transduction in mouse cochlear hair cells. *PLoS ONE*, 17(3), e0261530. https://doi.org/10.1371/journal.pone.0261530

Mah, J.L., Christensen-Dalsgaard, K.K. & Leys, S.P. (2014) Choanoflagellate and choanocyte collar-flagellar systems and the assumption of homology. *Evolution & Development*, 16(1), 25-37. https://doi.org/10.1111/ede.12060

Majstoravich, S., Zhang, J., Nicholson-Dykstra, S., Linder, S., Friedrich, W., Siminovitch, K.A. & Higgs, H.N. (2004) Lymphocyte microvilli are dynamic, actin-dependent structures that do not require Wiskott-Aldrich syndrome protein (WASp) for their morphology. *Blood*, 104(5), 1396-1403. https://doi.org/10.1182/blood-2004-02-0437

Mallavarapu, A. & Mitchison, T. (1999) Regulated actin cytoskeleton assembly at filopodium tips controls their extension and retraction. *The Journal of Cell Biology*, 146(5), 1097-1106

Martin, P., Bozovic, D., Choe, Y. & Hudspeth, A.J. (2003) Spon- taneous oscillation by hair bundles of the bullfrog's sacculus. *Journal of Neuroscience*, 23(11), 4533-4548. https://doi.org/10.1523/JNEUROSCI.23-11-04533.2003

Mattila, P.K. & Lappalainen, P. (2008) Filopodia: molecular architecture and cellular functions. *Nature Reviews Molecular Cell Biology*, 9(6), 446-454. https://doi.org/10.1038/nrm2406

McGhee, J. (2007) The C. elegans intestine. *WormBook*. Online resource. https://doi.org/10.1895/wormbook.1.133.1

McGrath, J., Roy, P. & Perrin, B.J. (2017) Stereocilia morphogenesis and maintenance through regulation of actin stability. *Seminars in Cell & Developmental Biology*, 65(May), 88-95. https://doi.org/10.1016/j.semcdb.2016.08.017

Medalia, O., Beck, M., Ecke, M., Weber, I., Neujahr, R., Baumeister, W. & Gerisch, G. (2007) Organization of actin networks in intact filopodia. *Current Biology*, 17(1), 79-84. https://doi.org/10.1016/j.cub.2006.11.022

Meenderink, L.M., Gaeta, I.M., Postema, M.M., Cencer, C.S., Chinowsky, C.R., Krystofiak, E.S., Millis, B.A. & Tyska, M.J. (2019) Actin dynamics drive microvillar motility and clustering during brush border assembly. *Developmental Cell*, 50(5), 545-556.e4. https://doi.org/10.1016/j.devcel.2019.07.008

Miihkinen, M., Grönloh, M.L.B., Popović, A., Vihinen, H., Jokitalo, E., Goult, B.T., Ivaska, J. & Jacquemet, G. (2021) Myosin-X and talin modulate integrin activity at filopodia tips. *Cell Reports*, 36(11), 109716. https://doi.org/10.1016/j.celrep.2021.109716

Mikrjukov, K.A. & Mylnikov, A.P. (2001) A study of the fine structure and the mitosis of a lamellicristate amoeba, *Micronuclearia Podoventalis* Gen. et Sp. Nov. (Nucleariidae, Rotosphaerida). *European Journal of Protistology*, 37(1), 15-24. https://doi.org/10.1078/0932-4739-00783

Milenkovic, V.M., Brockmann, M., Stöhr, H., Weber, B.H.F. & Strauss, O. (2010) Evolution and functional divergence of the anoctamin fam- ily of membrane proteins. *BMC Evolutionary Biology*, 10(1), 319. https://doi.org/10.1186/1471-2148-10-319

Miles, A., Blair, C., Emili, A. & Tropepe, V. (2021) Usher syndrome type 1-associated gene, pcdh15b, is required for photoreceptor struc- tural integrity in zebrafish. *Disease Models & Mechanisms*, 14(12), dmm048965. https://doi.org/10.1242/dmm.048965

Miura, S., Sato, K., Kato-Negishi, M., Teshima, T. & Takeuchi, S. (2015) Fluid shear triggers microvilli formation via mechanosensitive acti- vation of TRPV6. *Nature Communications*, 6(1), 8871. https://doi.org/10.1038/ncomms9871

Montoro, D.T., Haber, A.L., Rood, J.E., Regev, A. & Rajagopal, J. (2020) A synthesis concerning conservation and divergence of cell types across epithelia. *Cold Spring Harbor Perspectives in Biology*, 12(11), a035733. https://doi.org/10.1101/cshperspect.a035733

Morales, F.C., Takahashi, Y., Kreimann, E.L. & Georgescu, M.-M. (2004) Ezrin-Radixin-Moesin (ERM)-binding phosphoprotein 50 organizes ERM proteins at the apical membrane of polarized epithelia. *Proceedings of the National Academy of Sciences*, 101(51), 17705-17710. https://doi.org/10.1073/pnas.0407974101

Murphy, F. & Moiseff, A. (2019) Anatomy of the stemmata in the Pho- turis firefly larva. *Journal of Comparative Physiology A*, 205(1), 151-161. https://doi.org/10.1007/s00359-018-01312-2
Mylnikov, A.P., Tikhonenkov, D.V., Karpov, S.A. & Wylezich, C. (2019) Microscopical studies on *Ministeria vibrans* Tong, 1997 (Filasterea) highlight the cytoskeletal structure of the common ancestor of Filasterea, Metazoa and Choanoflagellata. *Protist*, 170(4), 385–396. https://doi.org/10.1016/j.protis.2019.07.001

Narayanan, P., Chatterton, P., Ikeda, A., Ikeda, S., Corey, D.P., Ervasti, J.M. & Perrin, B.J. (2015) Length regulation of mechanosensitive stereocilia depends on very slow actin dynamics and filament-severing proteins. *Nature Communications*, 6(1), 6855. https://doi.org/10.1038/ncomms7855

Nekliudova, U.A., Schwaha, T.F., Kotenko, O.N., Gruber, D., Cyran, N. & Ostrovsky, A.N. (2019) Sexual reproduction of the placental brooder Celleporella hyalina (Bryozoa, Cheilostomata) in the White Sea. *Journal of Morphology*, 280(2), 278–299. https://doi.org/10.1002/jmor.20943

Nichols, S.A., Roberts, B.W., Richter, D.J., Fairclough, S.R. & King, N. (2012) Origin of metazoan cadherin diversity and the antiquity of the classical cadherin/β-catenin complex. *Proceedings of the National Academy of Sciences of the United States of America*, 109(32), 13046–13051. https://doi.org/10.1073/pnas.1120685109

Nicholson-Dykstra, S.M. & Higgs, H.N. (2008) Arp2 depletion inhibits sheet-like protrusions but not linear protrusions of fibroblasts and lymphocytes. *Cell Motility and the Cytoskeleton*, 65(11), 904–922. https://doi.org/10.1002/cm.20312

Nielsen, C. (2008) Six major steps in animal evolution: are we derived sponge larvae? *Evolution & Development*, 10(2), 241–257. https://doi.org/10.1111/j.1525-142X.2008.00231.x

Nordström, K., Wallén, J.S. & Nilsson, D. (2003) A simple visual system without neurons in jellyfish larvae. *Proceedings of the Royal Society of London. Series B: Biological Sciences*, 270(1531), 2349–2354. https://doi.org/10.1098/rspb.2003.2504

Norekian, T.P. & Moroz, L.L. (2019) Neural system and receptor diversity in the ctenophore. *Journal of Comparative Neurology*, 527(12), 1986–2008. https://doi.org/10.1002/cne.24633

Okamoto, C.T. (2017) Regulation of transporters and channels by membrane-trafficking complexes in epithelial cells. *Cold Spring Harbor Perspectives in Biology*, 9(11), a027839. https://doi.org/10.1101/cshperspect.a027839

Oldenbourg, R., Katoh, K. & Danuser, G. (2000) Mechanism of lateral movement of filopodia and radial actin bundles across neuronal growth cones. *Biophysical Journal*, 78(3), 1176–1182. https://doi.org/10.1016/S0006-3495(00)76675-6

Orbach, R. & Su, X. (2020) Surfing on membrane waves: microvilli, curved membranes, and immune signaling. *Frontiers in Immunology*, 11(September), 2187. https://doi.org/10.3389/fimmu.2020.02187

Ott, E., Hoff, S., Indorf, L., Ditengou, F.A., Müller, J., Renschler, G., Lienkamp, S.S., Kramer-Zucker, A., Bergmann, C. & Epting, D. (2023) A novel role for the chloride intracellular channel protein Clic5 in ciliary function. *Scientific Reports*, 13(1), 17647. https://doi.org/10.1038/s41598-023-44235-y

Pacentine, I., Chatterjee, P. & Barr-Gillespie, P.G. (2020) Stereocilia rootlets: actin-based structures that are essential for structural stability of the hair bundle. *International Journal of Molecular Sciences*, 21(1), 324. https://doi.org/10.3390/ijms21010324

Park, J., Karplus, K., Barrett, C., Hughey, R., Haussler, D., Hubbard, T. & Chothia, C. (1998) Sequence comparisons using multiple sequences detect three times as many remote homologues as pairwise methods¹. *Journal of Molecular Biology*, 284(4), 1201–1210. https://doi.org/10.1006/jmbi.1998.2221

Parra-Acero, H., Harcet, M., Sánchez-Pons, N., Casacuberta, E., Brown, N.H., Dudin, O. & Ruiz-Trillo, I. (2020) Integrin-mediated adhesion in the unicellular holozoa *Capsaspora owczarzaki*. *Current Biology*, 30(21), 4270–4275.e4. https://doi.org/10.1016/j.cub.2020.08.015

Parra-Acero, H., Ros-Rocher, N., Perez-Posada, A., Kożyczczowska, A., Sánchez-Pons, N., Nakata, A., Suga, H., Najle, S.R. & Ruiz-Trillo, I. (2018) Transfection of *Capsaspora owczarzaki*, a close unicellular

relative of animals. *Development*, 145(10), dev162107. https://doi.org/10.1242/dev.162107

Pasha, S.N., Meenakshi, I. & Sowdhamini, R. (2016) Revisiting myosin families through large-scale sequence searches leads to the discovery of new myosins. *Evolutionary Bioinformatics Online*, 12(August), 201–211. https://doi.org/10.4137/EBO.S39880

Pelaseyed, T. & Bretscher, A. (2018a) Regulation of actin-based apical structures on epithelial cells. *Journal of Cell Science*, 131(20), jcs221853. https://doi.org/10.1242/jcs.221853

Pelaseyed, T. & Bretscher, A. (2018b) Regulation of actin-based apical structures on epithelial cells. *Journal of Cell Science*, 131(20), jcs221853. https://doi.org/10.1242/jcs.221853

Peña, J.F., Alié, A., Richter, D.J., Wang, L., Funayama, N. & Nichols, S.A. (2016) Conserved expression of vertebrate microvillar gene homologs in choanocytes of freshwater sponges. *EvoDevo*, 7(July), 13. https://doi.org/10.1186/s13227-016-0050-x

Peng, A.W., Belyantseva, I.A., Hsu, P.D., Friedman, T.B. & Heller, S. (2009) Twinfilin 2 regulates actin filament lengths in cochlear stereocilia. *The Journal of Neuroscience: The Official Journal of the Society for Neuroscience*, 29(48), 15083–15088. https://doi.org/10.1523/JNEUROSCI.2782-09.2009

Petersen, K. J., Goodson, H. V., Arthur, A. L., Luxton, G. W. G., Houdusse, A., & Titus, M. A. (2016) MyTH4-FERM myosins have an ancient and conserved role in filopod formation. *Proceedings of the National Academy of Sciences*, 113(50). https://doi.org/10.1073/pnas.1615392113

Pinette, J.A., Mao, S., Millis, B.A., Krystofiak, E.S., Faust, J.J. & Tyska, M.J. (2019) Brush border protocadherin CDHR2 promotes the elongation and maximized packing of microvilli in vivo. *Molecular Biology of the Cell*, 30(1), 108–118. https://doi.org/10.1091/mbc.E18-09-0558

Poncelet, G. & Shimeld, S.M. (2020) The evolutionary origins of the vertebrate olfactory system. *Open Biology*, 10(12), 200330. https://doi.org/10.1098/rsob.200330

Postema, M.M., Grega-Larson, N.E., Meenderink, L.M. & Tyska, M.J. (2019) PACSIN2-dependent apical endocytosis regulates the morphology of epithelial microvilli. *Molecular Biology of the Cell*, 30(19), 2515–2526. https://doi.org/10.1091/mbc.E19-06-0352

Postema, M.M., Grega-Larson, N.E., Neininger, A.C. & Tyska, M.J. (2018) IRTKS (BAIAP2L1) elongates epithelial microvilli using EPS8-dependent and independent mechanisms. *Current Biology: CB*, 28(18), 2876–2888.e4. https://doi.org/10.1016/j.cub.2018.07.022

Pozdnyakov, I.R., Sokolova, A.M., Severtzov, A.N., Ereskovsky, A.V. & Karpov, S.A. (2017) Kinetid structure of choanoflagellates and choanocytes of sponges does not support their close relationship. *Protistology*, 11(4): 248–264. https://doi.org/10.21685/1680-0826-2017-11-4-6

Ramachandran, K., Ansel, M., Brunet, T. & Dey, G. (2024) Origin and evolution of microvilli—code. GitHub. Online resource. 2024. https://github.com/kausthubhr/microvilli phylogenetics

Ramalho, J.J., Sepers, J.J., Nicolle, O., Schmidt, R., Cravo, J., Michaux, G. & Boxem, M. (2020) C-terminal phosphorylation modulates ERM-1 localization and dynamics to control cortical actin organization and support lumen formation during *Caenorhabditis elegans* development. *Development*, 147(14), dev188011. https://doi.org/10.1242/dev.188011

Rieger, R.M. (1976) Monociliated epidermal cells in gastrotricha: significance for concepts of early metazoan evolution. *Journal of Zoological Systematics and Evolutionary Research*, 14(3), 198–226.

Rix, L., Ribes, M., Coma, R., Jahn, M. T., de Goeij, J. M., van Oevelen, D., Escrig, S., Meibom, A., & Hentschel, U. (2020) Heterotrophy in the earliest gut: a single-cell view of heterotrophic carbon and nitrogen assimilation in sponge-microbe symbioses. *The ISME Journal*, 14(10), 2554–2567. https://doi.org/10.1038/s41396-020-0706-3

Robles, E., Huttenlocher, A. & Gomez, T.M. (2003) Filopodial calcium transients regulate growth cone motility and guidance through local
activation of calpain. *Neuron*, 38(4), 597–609. https://doi.org/10.1016/S0896-6273(03)00260-5

Ros-Rocher, N. & Brunet, T. (2023) What is it like to be a choanoflagellate? Sensation, processing and behavior in the closest unicellular relatives of animals. *Animal Cognition*, 26(6), 1767–1782. https://doi.org/10.1007/s10071-023-01776-z

Ros-Rocher, N., Kidner, R.Q., Gerdt, C., Sean Davidson, W., Ruiz-Trillo, I. & Gerdt, J.P. (2023) Chemical factors induce aggregative multicellularity in a close unicellular relative of animals. *Proceedings of the National Academy of Sciences*, 120(18), e2216668120. https://doi.org/10.1073/pnas.2216668120

Rost-Roszkowska, M.M., Świątek, P., Kszuk, M., Główczyk, K. & Bielecki, A. (2012) Morphology and ultrastructure of the midgut in Piscicola geometra (Annelida, Hirudinea). *Protoplasma*, 249(4), 1037–1047. https://doi.org/10.1007/s00709-011-0337-7

Ruiz-Trillo, I. (2014) Ministeria Vibrans 3 [Scanning Electron Micrograph]. Flickr. Onlin resource. https://www.flickr.com/photos/121041113@N07/14474052808/in/photostream/

Ruiz-Trillo, I., Roger, A.J., Burger, G., Gray, M.W. & Franz Lang, B. (2008) A phylogenomic investigation into the origin of metazoa. *Molecular Biology and Evolution*, 25(4), 664–672. https://doi.org/10.1093/molbev/msn006

Sahly, I., Dufour, E., Schietroma, C., Michel, V., Bahloul, A., Perfettini, I., Pepermans, E., et al. (2012) Localization of usher 1 proteins to the photoreceptor calyxal processes, which are absent from mice. *The Journal of Cell Biology*, 199(2), 381–399. https://doi.org/10.1083/jcb.201202012

Salvini-Plawen, L.V. (1978) On the origin and evolution of the lower metazoa. *Journal of Zoological Systematics and Evolutionary Research*, 16(1), 40–87.

Sauvanet, C., Wayt, J., Pelaseyed, T. & Bretscher, A. (2015) Structure, regulation, and functional diversity of microvilli on the apical domain of epithelial cells. *Annual Review of Cell and Developmental Biology*, 31(1), 593–621. https://doi.org/10.1146/annurev-cellbio-100814-125234

Saville-Kent, W. (1880) A manual of the infusoria: including a description of all known flagellate, ciliate, and tentaculiferous protozoa, British and foreign, and an account of the organization and the affinities of the sponges, vol. 1–2. London, United Kingdom: D. Bogue.

Schultz, J. & Terhoeven, N. (2013) The bilaterian roots of cordon-bleu. *BMC Research Notes*, 6(September), 393. https://doi.org/10.1186/1756-0500-6-393

Sebé-Pedrós, A., Burkhardt, P., Sánchez-Pons, N., Fairclough, S.R., Franz Lang, B., King, N. & Ruiz-Trillo, I. (2013) Insights into the origin of metazoan filopodia and microvilli. *Molecular Biology and Evolution*, 30(9), 2013–2023. https://doi.org/10.1093/molbev/mst110

Sebé-Pedrós, A., Degnan, B.M. & Ruiz-Trillo, I. (2017) The origin of metazoa: a unicellular perspective. *Nature Reviews. Genetics*, 18(8), 498–512. https://doi.org/10.1038/nrg.2017.21

Sebé-Pedrós, A., Grau-Bové, X., Richards, T.A. & Ruiz-Trillo, I. (2014) Evolution and classification of myosins, a paneukaryotic whole-genome approach. *Genome Biology and Evolution*, 6(2), 290–305. https://doi.org/10.1093/gbe/evu013

Sepers, J.J., Ramalho, J.J., Kroll, J.R., Schmidt, R. & Boxem, M. (2022) ERM-1 phosphorylation and NRFL-1 redundantly control lumen formation in the C. elegans intestine. *Frontiers in Cell and Developmental Biology*, 10(February), 769862. https://doi.org/10.3389/fcell.2022.769862

Shabardina, V., Kashima, Y., Suzuki, Y. & Makalowski, W. (2020) Emergence and evolution of ERM proteins and merlin in metazoans. *Genome Biology and Evolution*, 12(1), 3710–3724. https://doi.org/10.1093/gbe/evz265

Sharkova, M., Aparicio, G., Mouzaaber, C., Zolessi, F.R. & Hocking, J.C. (2024) Photoreceptor calyxal processes accompany the developing outer segment, adopting a stable length despite a

dynamic core. *Journal of Cell Science*, 137(7), jcs261721. https://doi.org/10.1242/jcs.261721

Sharkova, M., Chow, E., Erickson, T. & Hocking, J.C. (2022) The morphological and functional diversity of apical microvilli. *Journal of Anatomy*, 242(3), 327–353. https://doi.org/10.1111/joa.13781

Sogabe, S., Hatleberg, W.L., Kocot, K.M., Say, T.E., Stoupin, D., Roper, K.E., Fernandez-Valverde, S.L., Degnan, S.M. & Degnan, B.M. (2019) Pluripotency and the origin of animal multicellularity. *Nature*, 570(7762), 519–522. https://doi.org/10.1038/s41586-019-1290-4

Solé, M., Lenoir, M., Manuel Fontuño, J., Durfort, M., van der Schaar, M. & André, M. (2016) Evidence of Cnidarians sensitivity to sound after exposure to low frequency noise underwater sources. *Scientific Reports*, 6(1), 37979. https://doi.org/10.1038/srep37979

Staun-Ram, E. & Shalev, E. (2005) Human trophoblast function during the implantation process. *Reproductive Biology and Endocrinology*, 3(1), 56. https://doi.org/10.1186/1477-7827-3-56

Steinmetz, P.R.H. (2019) A non-bilaterian perspective on the development and evolution of animal digestive systems. *Cell and Tissue Research*, 377(3), 321–339. https://doi.org/10.1007/s00441-019-03075-x

Theofanopoulou, C., Gastaldon, S., O'Rourke, T., Samuels, B.D., Messner, A., Martins, P.T., Delogu, F., Alamri, S. & Boeckx, C. (2017) Self-domestication in homo sapiens: insights from comparative genomics. *PLOS ONE*, 12(10), e0185306. https://doi.org/10.1371/journal.pone.0185306

Tikhonenkov, D.V., Hehenberger, E., Esaulov, A.S., Belyakova, O.I., Mazei, Y.A., Mylnikov, A.P. & Keeling, P.J. (2020) Insights into the origin of metazoan multicellularity from predatory unicellular relatives of animals. *BMC Biology*, 18(1), 39. https://doi.org/10.1186/s12915-020-0762-1

Tilney, L.G., Tilney, M.S. & Cotanche, D.A. (1988) Actin filaments, stereocilia, and hair cells of the bird cochlea. V. How the staircase pattern of stereociliary lengths is generated. *The Journal of Cell Biology*, 106(2), 355–365.

Tonucci, F.M., Ferretti, A., Almada, E., Cribb, P., Vena, R., Hidalgo, F., Favre, C., Tyska, M.J., Kaverina, I. & Larocca, M.C. (2018) Microtubules regulate brush border formation. *Journal of Cellular Physiology*, 233(2), 1468–1480. https://doi.org/10.1002/jcp.26033

Triana-Garcia, P.A., Nevitt, G.A., Pesavento, J.B. & Teh, S.J. (2021) Gross morphology, histology, and ultrastructure of the olfactory rosette of a critically endangered indicator species, the Delta Smelt, Hypomesus Transpacificus. *Journal of Comparative Physiology A*, 207(5), 597–616. https://doi.org/10.1007/s00359-021-01500-7

Tsubouchi, A., Caldwell, J.C. & Daniel Tracey, W. (2012) Dendritic filopodia, ripped pocket, NOMPC, and NMDARs contribute to the sense of touch in drosophila larvae. *Current Biology*, 22(22), 2124–2134. https://doi.org/10.1016/j.cub.2012.09.019

Ubelmann, F., Chamaillard, M., El-Marjou, F., Simon, A., Netter, J., Vignjevic, D., Nichols, B.L., et al. (2013) Enterocyte loss of polarity and gut wound healing rely upon the F-actin–severing function of villin. *Proceedings of the National Academy of Sciences*, 110(15), E1380–E1389. https://doi.org/10.1073/pnas.1218446110

Ullrich-Lüter, E.M., Dupont, S., Arboleda, E., Hausen, H. & Arnone, M.I. (2011) Unique system of photoreceptors in sea urchin tube feet. *Proceedings of the National Academy of Sciences*, 108(20), 8367–8372. https://doi.org/10.1073/pnas.1018495108

Vélez-Ortega, A.C., Freeman, M.J., Indzhykulian, A.A., Grossheim, J.M. & Frolenkov, G.I. (2017) Mechanotransduction current is essential for stability of the transducing stereocilia in mammalian auditory hair cells. *eLife*, 6, e24661. https://doi.org/10.7554/eLife.24661

Velle, K.B. & Fritz-Laylin, L.K. (2020) Conserved actin machinery drives microtubule-independent motility and phagocytosis in Naegleria. *Journal of Cell Biology*, 219(11), e202007158. https://doi.org/10.1083/jcb.202007158

Venkatachalam, M.A., Bernard, D.B., Donohoe, J.F. & Levinsky, N.G. (1978) Ischemic damage and repair in the rat proximal tubule: differ-
ences among the S1, S2, and S3 segments. *Kidney International*, 14(1), 31–49. https://doi.org/10.1038/ki.1978.87

Verpy, E., Leibovici, M., Michalski, N., Goodyear, R.J., Houdon, C., Weil, D., Richardson, G.P. & Petit, C. (2011) Stereocilin connects outer-hair-cell stereocilia to one another and to the tectorial membrane. *The Journal of Comparative Neurology*, 519(2), 194–210. https://doi.org/10.1002/cne.22509

Wan, K.Y. (2018) Coordination of eukaryotic cilia and flagella. *Essays in Biochemistry*, 62(6), 829–838. https://doi.org/10.1042/EBC20180029

Wang, T., Ling, A.H., Billings, S.E., Hosseini, D.K., Vaisbuch, Y., Kim, G.S., Atkinson, P.J., et al. (2024) Single-cell transcriptomic atlas reveals increased regeneration in diseased human inner ear balance organs. *Nature Communications*, 15(1), 4833. https://doi.org/10.1038/s41467-024-48491-y

Wang, X., Liu, S., Cheng, Q., Qu, C., Ren, R., Du, H., Li, N., et al. (2023) CIB2 and CIB3 regulate stereocilia maintenance and mechanoelectrical transduction in mouse vestibular hair cells. *Journal of Neuroscience*, 43(18), 3219–3231. https://doi.org/10.1523/JNEUROSCI.1807-22.2023

Weinbaum, S., Duan, Y., Satlin, L.M., Wang, T. & Weinstein, A.M. (2010) Mechanotransduction in the renal tubule. *American Journal of Physiology—Renal Physiology*, 299(6), F1220–F1236. https://doi.org/10.1152/ajprenal.00453.2010

Wilson, N.F. & Snell, W.J. (1998) Microvilli and cell-cell fusion during fertilization. *Trends in Cell Biology*, 8(3), 93–96. https://doi.org/10.1016/S0962-8924(98)01234-3

Wolpert, L. & Mercer, E.H. (1963) An electron microscope study of the development of the blastula of the sea urchin embryo and its radial polarity. *Experimental Cell Research*, 30(2), 280–300. https://doi.org/10.1016/0014-4827(63)90300-8

Wright, B.A., Kvansakul, M., Schierwater, B. & Humbert, P.O. (2022) Cell polarity signalling at the birth of multicellularity: what can we learn from the first animals. *Frontiers in Cell and Developmental Biology*, 10, 1024489. https://www.frontiersin.org/articles/10.3389/fcell.2022.1024489

Wyart, C., Carbo-Tano, M., Cantaut-Belarif, Y., Orts-Del'Immagine, A. & Böhm, U.L. (2023) Cerebrospinal fluid-contacting neurons: multimodal cells with diverse roles in the CNS. *Nature Reviews Neuroscience*, 24(9), 540–556. https://doi.org/10.1038/s41583-023-00723-8

Xiong, J., Feng, J., Yuan, D., Zhou, J. & Miao, W. (2015) Tracing the structural evolution of eukaryotic ATP binding cassette transporter superfamily. *Scientific Reports*, 5(1), 16724. https://doi.org/10.1038/srep16724

Yang, X., Qi, J., Zhang, L., Tan, F., Huang, H., Xu, C., Cui, Y., Chai, R. & Wu, P. (2023) The role of espin in the stereocilia regeneration and protection in Atoh1-overexpressed cochlear epithelium. *Cell Proliferation*, 56(11), e13483. https://doi.org/10.1111/cpr.13483

Zhai, X., Du, H., Shen, Y., Zhang, X., Chen, Z., Wang, Y. & Xu, Z. (2022) FCHSD2 is required for stereocilia maintenance in mouse cochlear hair cells. *Journal of Cell Science*, 135(16), jcs259912. https://doi.org/10.1242/jcs.259912

Zhang, D.-S., Piazza, V., Perrin, B.J., Rzadzinska, A.K., Collin Poczatek, J., Wang, M., Prosser, H.M., Ervasti, J.M., Corey, D.P. & Lechene, C.P. (2012) Multi-isotope imaging mass spectrometry reveals slow protein turnover in hair-cell stereocilia. *Nature*, 481(7382), 520–524. https://doi.org/10.1038/nature10745

Zhang, Y., Wang, Y., Feng, X., Zhang, S., Xu, X., Li, L., Niu, S., et al. (2021) Oocyte-derived microvilli control female fertility by optimizing ovarian follicle selection in mice. *Nature Communications*, 12(1), 2523. https://doi.org/10.1038/s41467-021-22829-2

Zhao, H., Pykäläinen, A. & Lappalainen, P. (2011) I-BAR domain proteins: linking actin and plasma membrane dynamics. *Current Opinion in Cell Biology*, 23(1), 14–21. https://doi.org/10.1016/j.ceb.2010.10.005

Zhou, K., Sumigray, K.D. & Lechler, T. (2015) The Arp2/3 complex has essential roles in vesicle trafficking and transcytosis in the mammalian small intestine. *Molecular Biology of the Cell*, 26(11), 1995–2004. https://doi.org/10.1091/mbc.E14-10-1481

Zhou, L.-Y., Jin, C.-X., Wang, W.-X., Song, L., Shin, J.-B., Du, T.-T. & Wu, H. (2023) Differential regulation of hair cell actin cytoskeleton mediated by SRF and MRTFB. *eLife*, 12, e90155. https://doi.org/10.7554/eLife.90155

Zhu, H., Li, M., Zhao, R., Li, M., Chai, Y., Zhu, Z., Yang, Y., et al. (2022) In situ structure of intestinal apical surface reveals nanobristles on microvilli. *Proceedings of the National Academy of Sciences*, 119(24), e2122249119. https://doi.org/10.1073/pnas.2122249119

Zou, J., Zheng, T., Ren, C., Askew, C., Liu, X.-P., Pan, B., Holt, J.R., Wang, Y. & Yang, J. (2014) Deletion of PDZD7 disrupts the usher syndrome type 2 protein complex in cochlear hair cells and causes hearing loss in mice. *Human Molecular Genetics*, 23(9), 2374–2390. https://doi.org/10.1093/hmg/ddt629

Zwaenepoel, I., Naba, A., Da Cunha, M.M.L., Del Maestro, L., Formstecher, E., Louvard, D. & Arpin, M. (2012) Ezrin regulates microvillus morphogenesis by promoting distinct activities of Eps8 proteins. *Molecular Biology of the Cell*, 23(6), 1080–1095. https://doi.org/10.1091/mbc.e11-07-0588

## SUPPORTING INFORMATION

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**How to cite this article:** Ansel, M., Ramachandran, K., Dey, G. & Brunet, T. (2024) Origin and evolution of microvilli. *Biology of the Cell*, 116, e2400054. https://doi.org/10.1111/boc.202400054