**Interdisciplinary Synergy to Reveal Mechanisms of Annexin-Mediated Plasma Membrane Shaping and Repair**

**Poul Martin Bendix 1,\*, Adam Cohen Simonsen 2,\*, Christo**ff**er D. Florentsen 1, Swantje Christin Häger 3, Anna Mularski 2, Ali Asghar Hakami Zanjani 2, Guillermo Moreno-Pescador 1, Martin Berg Klenow 2, Stine Lauritzen Sønder 3, Helena M. Danielsen 1, Mohammad Reza Arastoo 1, Anne Sofie Heitmann 3, Mayank Prakash Pandey 2, Frederik Wendelboe Lund 2, Catarina Dias 3, Himanshu Khandelia 2,\* and Jesper Nylandsted 3,4,\***


Received: 2 April 2020; Accepted: 19 April 2020; Published: 21 April 2020

**Abstract:** The plasma membrane surrounds every single cell and essentially shapes cell life by separating the interior from the external environment. Thus, maintenance of cell membrane integrity is essential to prevent death caused by disruption of the plasma membrane. To counteract plasma membrane injuries, eukaryotic cells have developed efficient repair tools that depend on Ca2<sup>+</sup>- and phospholipid-binding annexin proteins. Upon membrane damage, annexin family members are activated by a Ca2<sup>+</sup> influx, enabling them to quickly bind at the damaged membrane and facilitate wound healing. Our recent studies, based on interdisciplinary research synergy across molecular cell biology, experimental membrane physics, and computational simulations show that annexins have additional biophysical functions in the repair response besides enabling membrane fusion. Annexins possess different membrane-shaping properties, allowing for a tailored response that involves rapid bending, constriction, and fusion of membrane edges for resealing. Moreover, some annexins have high affinity for highly curved membranes that appear at free edges near rupture sites, a property that might accelerate their recruitment for rapid repair. Here, we discuss the mechanisms of annexin-mediated membrane shaping and curvature sensing in the light of our interdisciplinary approach to study plasma membrane repair.

**Keywords:** annexin; plasma membrane repair; membrane curvature; membrane curvature sensing; membrane shaping; interdisciplinary research; cell rupture; membrane damage

### **1. Introduction**

Membranes of eukaryotic cells play fundamental roles in organizing cellular compartments and cluster specific molecules to generate signaling platforms. Beyond defining the physical cell boundary, the plasma membrane serves a dual purpose: it allows the communication and exchange of solutes with the extracellular environment, while guarding the cell from its surroundings. In addition, the plasma membrane facilitates cell–cell interactions and adhesion and regulates the architecture of the cell. Thus, it is crucial that the plasma membrane is kept intact to ensure cell survival.

The plasma membrane is composed of both structural and signaling lipids, as well as embedded proteins. The major structural lipids are glycerophospholipids (such as phosphatidylserine and phosphatidylcholine), sterols, and sphingolipids (e.g., sphingomyelin) [1]. Of note, the enrichment of sphingolipids and sterols, like cholesterol, aids in resisting mechanical stress [2]. Plasma membrane topology is shaped by the geometrical properties of its lipids, in combination with both integrated and peripheral proteins, including components of the cytoskeleton [3]. Glycerophospholipids with small head groups, such as phosphatidylethanolamines and phosphatidic acid, are cone-shaped and have the propensity to induce membrane curvature, thus lipid shape is intimately related to membrane shape [3,4]. Moreover, the surface charge of membranes influences its interaction with proteins, including curvature inducing regulators, and is determined by the head group charge of individual lipids [5,6]. This is exemplified by the family of annexin proteins that are characterized by their capacity to bind negatively charged phospholipids in a Ca2+-dependent manner, to exert their various functions in membrane organization, trafficking, and repair [7].

Unlike prokaryotic cells, which are shielded by a cell wall, mammalian cells are primarily protected by their plasma membrane and appear more susceptible to cell injuries. To counteract the eminent threat that membrane damage would impose on cellular homeostasis, robust repair mechanisms have evolved to ensure that membrane holes are sealed within the timeframe of tens-of-seconds to maintain cell integrity [8,9]. If disruptions to the plasma membrane are not repaired rapidly, the resultant pronounced osmotic and ionic imbalances would lead to cell death. Skeletal muscle and cardiac muscle cells are classical examples of cells that, due to their high mechanic activity, need to cope with extensive plasma membrane disruptions [10]. Similarly, epithelial cells in the gut and skin fibroblasts are exposed to both mechanical and chemical stimuli and experience regular wounding [11]. However, it seems that all cell types can potentially experience plasma membrane injury and have the capacity to repair lesions [12].

Deficiency in plasma membrane repair is associated with several diseases. The best established link between repair deficiency and disease phenotype is observed in muscular dystrophies, such as dysferlin-deficient muscular dystrophy, where a failure to repair repeated membrane lesions leads to progressive muscle wasting [13]. Inversely, our research supports that enhanced plasma membrane repair plays a role in helping metastatic breast cancer cells cope with the physical stress imposed during invasion [14,15].

The importance of Ca2<sup>+</sup> for membrane repair was recognized in 1930 by Heilbrunn, who discovered that sea urchin eggs failed to repair upon mechanical damage when the surrounding solution was depleted of Ca2<sup>+</sup> [16]. Despite the scarce knowledge regarding membranes and cell compartmentalization at the time, this discovery pointed to Ca2<sup>+</sup> as a fundamental trigger of plasma membrane repair. In intact resting cells the intracellular free Ca2<sup>+</sup> concentrations in the cytoplasm are in the nanomolar range (~100 nM), while extracellular Ca2<sup>+</sup> concentrations are in the millimolar range [17]. This steep Ca2<sup>+</sup> gradient is important for signaling events and is established by actively pumping Ca2<sup>+</sup> into the extracellular space and intracellular Ca2<sup>+</sup> stores, which is also crucial to prevent Ca2<sup>+</sup> cytotoxicity. Upon plasma membrane injury, the Ca2<sup>+</sup> gradient leads to a rapid and pronounced flux of Ca2<sup>+</sup> into the cell's cytoplasm, which poses a threat to the cell [18,19]. However, this rise in Ca2<sup>+</sup> acts at the same time as the triggering event for plasma membrane repair mechanisms.

### **2. Annexins in Repair**

Several functions have been reported for annexins, and most of them include their involvement in membrane trafficking events [20]. Interestingly, during the last two decades more evidence regarding their role in plasma membrane repair has emerged [20–23]. Most proteins that play a role in plasma membrane repair have the ability to bind Ca2<sup>+</sup> and become activated upon injury-induced Ca2<sup>+</sup> influx. However, the recruitment of repair components to the injured plasma membrane can also be facilitated via complex formation with other Ca2<sup>+</sup>-binding proteins. This is exemplified by recent data showing a link between the Ca2<sup>+</sup>-dependent, phospholipid-binding protein annexin A7 (ANXA7), and the shedding of damaged membrane during repair facilitated by the endosomal sorting complex required for transport III (ESCRT-III) complex [22]. In detail, ANXA7 localizes to the injury site where it forms a complex with apoptosis-linked gene 2 (ALG-2) and ALG-2-interacting protein X (ALIX) [23]. ANXA7 facilitates thereby anchoring of ALG-2 to the damaged plasma membrane, where ALG-2 can assemble the ESCRT-III complex and drive shedding of damaged membrane [22].

Accumulating evidence indicates that annexins are instrumental for coping with plasma membrane injuries, although their exact mode of action is not well characterized. Annexins are a family of proteins (in mammals: ANXA1-11 and ANXA13) which share a structurally conserved C-terminal domain and the functional property of being able to bind anionic phospholipids in a Ca2+-dependent manner. Binding to membranes is mediated by their core domain, which consists of four characteristic structural annexin repeats [24]. The N-terminal domain varies in length and composition and is responsible for the interaction with other proteins, giving rise to different functional properties [24].

Work on dysferlin-deficient mouse muscle cells pioneered the hypothesis that annexins function in plasma membrane repair. ANXA1 and ANXA2 were found to associate with dysferlin upon muscle cell membrane injury. Since ANXA1 and ANXA2 were known to cause aggregation and fusion of liposomes in vitro [25], Lennon et al. proposed that the annexins could aid repair by facilitating fusion of vesicles with the injured plasma membrane [26]. Wounding experiments, using laser injury, confirmed that ANXA1 localizes to the injury site and that inhibition of ANXA1 function impedes plasma membrane repair [27].

Involvement of ANXA1 and ANXA6 could also be detected in microvesicle shedding of streptolysin O (SLO) pores [28]. Furthermore, data from the Protein Data Bank (PDB), where annexins have been crystallized, show that annexins have varying Ca2<sup>+</sup> binding affinity, which adds another level of complexity to plasma membrane repair mechanisms. In line with this, ANXA6 was shown to have higher Ca2+ sensitivity than ANXA1, which led to differential recruitment patterns depending on the extent of the membrane lesions [28]. The importance of ANXA6 in muscle cell plasma membrane repair was recognized when mutations of the ANXA6 gene were found to negatively impact the phenotype of dysferlinopathy (a group of rare muscular dystrophies with recessive mutations in the dysferlin gene). Focal laser injury experiments revealed that ANXA6 localizes to the site of injury within seconds forming a repair cap [29,30]. Later work showed that ANXA6 localization to the injury site in muscle cells is actin-dependent and coincides with recruitment of ANXA1, ANXA2, and ANXA5 [31].

Even though the function of annexins in plasma membrane repair was initially merely explained by the ability of annexins to aid vesicle fusion, research now indicates that other mechanisms play a role for annexin-mediated wound closure. ANXA5, for example, was shown to assemble into 2D protein arrays around the injury site, preventing wound expansion [32,33]. To that end, using supported membrane models we have identified membrane binding and shaping features of other annexins that are independent of vesicle fusion events, pointing to a direct effect of annexins on membranes [34]. For example, ANXA4 was shown to generate a curvature force at free membrane edges because of its Ca2<sup>+</sup>-dependent homo-trimerization, while binding of ANXA6 led to a constriction force [35]. Therefore, it is likely that annexins possess different membrane-shaping properties, allowing for a tailored response that meets the needs of repair. In concert with other described processes, such as actin remodeling, annexins lead to efficient and rapid repair.

Interestingly, certain annexins, which are central components in plasma membrane repair, also display strong affinity for high membrane curvatures as we have shown recently in Moreno-Pescador et al. [36]. This suggests that membrane shaping and curvature sensing are coupled in a feedback loop where initial curvature generation by annexins leads to further recruitment of more annexins. It is therefore essential that the effect of biophysical cues, like membrane curvature, are thoroughly investigated for a comprehensive understanding of the activity and function of annexins.

### **3. Interdisciplinary Approaches to Address Annexin Function**

Given their complexity and rapid dynamics, detailed mechanisms of annexin-mediated membrane shaping and repair can only be resolved by incorporating sophisticated physical methods for assessing biophysical parameters like curvature and tension. Thus, interdisciplinary research within biophysics, molecular simulations, and molecular and cellular biology is needed to understand these mechanisms. This approach proved successful, as exemplified in recent studies showing that annexin family members share a common ability of membrane curvature induction, which seems to be important for their function in repair [24,25]. Here, the impact of annexins on planar membranes with stable, free edges are relevant for studying the membrane conformation near a hole.

The annexin core domain has a convex shape at its membrane-binding interface [21]. Thus, membrane-association of annexins can generally be expected to induce spontaneous membrane curvature and possibly produce membrane shape changes. The shape changes generated by annexins depend crucially on the initial membrane geometry and on the presence of free membrane edges in particular. Thus, the use of membrane model systems prepared with specific geometries allow the systematic study of the interplay between annexin binding, membrane shape, and shape remodeling.

Supported planar membranes in a stacked conformation are prepared using spincoating with the secondary membranes existing as isolated patches on the primary membrane. The patches have stable, free edges at their borders and have minimal interactions with the solid support due to the presence of the primary membrane [26]. These membrane patches serve as useful models for the plasma membrane near the injury site and address the experimental challenge that a membrane containing a hole is often unstable and the hole is short-lived. The planar patches allow out-of-plane bending, away from the supported surface and can be used for monitoring shape changes induced by different proteins. To this end, the impact of different annexin family members on the morphology of membrane patches with free edges were recently examined [24].

Notably, the annexin members ANXA4 and ANXA5 were both found to induce rolling of the patch starting from the free membrane edges at the patch border. Complete roll-up of a cell-sized membrane patch (50–100 μm) occurs over a time scale of 5–10 s. This result shows that membrane binding of ANXA4 and ANXA5 under physiological conditions induces spontaneous curvature. The curvature generation can be expected to also occur near a plasma membrane hole when Ca2<sup>+</sup> briefly enters the cytoplasm and cytosolic annexin binds to the negatively charged internal leaflet.

In order to model curvature generation around a plasma membrane hole, it has proven useful to describe the system using theoretical modeling of the membrane curvature energy.

According to the classical theory by Helfrich [27], the curvature elastic energy *H*curve of a membrane with area A can be written as:

$$H\_{\text{curve}} = \int\_{A} \left[ \frac{1}{2} k\_c (\overline{\mathbf{c}} - \mathbf{c}\_0)^2 + k\_G \overline{\mathbf{c}\_G} \right] dA \tag{1}$$

where *kc* is the mean curvature elastic modulus [J], *kG* is the gaussian curvature elastic modulus [J], and *c*<sup>0</sup> is the spontaneous curvature [*m*<sup>−</sup>1]. The local curvature of the membrane is described by the two principal radii of curvature, *R*<sup>1</sup> and *R*2. The mean curvature is defined as: *c* = <sup>1</sup> *<sup>R</sup>*<sup>1</sup> <sup>+</sup> <sup>1</sup> *<sup>R</sup>*<sup>2</sup> and the gaussian curvature as: *cG* = <sup>1</sup> *R*1 1 *R*2 .

*Hcurve* describes the curvature energy associated with a membrane of a general shape, for example a curved membrane near a plasma membrane hole. The spontaneous curvature co, is a quantity describing the tendency of a membrane to spontaneously curve with a curvature radius (*1*/*co*). The effect of annexin binding to a membrane is potentially complex, but it can approximately be modeled as a non-zero value of *co*. When describing a membrane containing a hole, there will additionally exist a tension force (edge tension) associated with the formation of the free edge. In a simple picture, the edge tension acts to contract the hole while the curvature energy (*Hcurve* equation) acts oppositely, by bending the membrane out-of-plane and increasing the edge radius. As previously shown [28], an equilibrium configuration is possible that balances the curvature and tension energies to create a stable neck-like shape of the membrane near the hole. We propose that ending of the membrane near a plasma membrane hole, via a mechanism as described above, plays a functional role in the plasma membrane repair process. In a cellular system, membrane re-shaping is envisioned to involve the concerted action of several annexins plus other repair proteins to rapidly bend, constrict, and finally seal the hole (Figure 1A–C).

Members of the family of human annexins were shown to induce distinctly different morphologies in the planar membrane patches [24]. This was observed despite the fact that the annexins all contain a membrane-binding core domain, which is highly conserved. In addition to large scale (cooperative) rolling as induced by ANXA4 and ANXA5, rolling in a fragmented morphology was observed for ANXA3 and ANXA13. Rolling was not observed for ANXA1 and ANXA2, which instead both induced a blebbing/folding type morphology of the membrane patch (Figure 1D,E). ANXA7 and ANXA11 induce rolling, in addition to the generation of lens-shaped membrane inclusions containing the protein and phosphatidylserine lipids. In total, the morphologies induced by annexins in membrane patches correlate well with a dendrogram of their amino acid sequences [24]. This points to an important functional role of the N-terminal annexin domain in reshaping membranes.

A deeper insight into the interplay between molecular curvatures and the rich polymorphic membrane shapes, which can be induced by annexins, will require theoretical simulations and also development of assays for studying membrane shaping in 3D, e.g., surrounding a membrane hole, and to study curvature sensing by this large class of proteins.

**Figure 1.** Binding of annexins (green) to a planarmembrane patch with free edges and adhesion energy wad inducing spontaneous curvature and a rolling morphology of the patch (**A**). Translation to the geometry of a membrane hole (**B**) where the edge tension τ and the spontaneous curvature c0 acts to create a stable neck conformation. Example of blebbing/folding morphologies induced by ANXA1 and ANXA2 (**C**) and examples of fluorescence data for patches (POPC: 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, POPS: (1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-L- serine), 9:1 ratio, DiDC18) showing blebbing (**D**) and rolling (**E**).

More specifically, the recent developments in super-resolution microscopy, like stochastic optical reconstruction microscopy (STORM) [29] and stimulated emission depletion (STED) [30], will be valuable for investigating the shape evolution of the injured site. STORM has been used to image the cortical actin of cells with great detail [31] and could be used for resolving the rearrangement of cortical actin, which is known to regulate both plasma membrane shape and tension. Finally, faster imaging modes like STED or high speed-atomic force microscopy (AFM) could capture the steps in the formation of a hole and the subsequent membrane healing.

### **4. Annexins Are Recruited by Membrane Curvature**

Proteins with the ability to shape membranes into highly curved structures often have the ability to sense high membrane curvatures. Such proteins include the well-studied Bin/amphiphysin/Rvs BAR domain containing proteins, which have all been verified as both curvature generators and sensors of specific membrane curvatures that correlate with their molecular shape [32–34]. The curvature sensing ability of annexins has only recently been recognized [35,36] despite the convex shape of the conserved membrane binding core domain, present in all annexins. Naively, one could expect that the convex shape, of the membrane binding domain, would imply that all annexins could be potential sensors of negative membrane curvatures. However, looking at the non-trivial curvature generation by different annexins this argument could be too simple, in particular considering the complex protein–protein interactions, variable Ca2<sup>+</sup> sensitivity, and the variability in the N-terminal domains amongst the annexins.

Our studies, using isolated giant plasma membranes vesicles (GPMVs), have shown that both ANXA4 [36] and ANXA5 [35] are efficient sensors of negative membrane curvatures. Optical manipulation of the GPMVs was used to extract nanotubes which exhibit extreme membrane curvatures as shown in several studies on giant unilamellar vesicles (GUVs) [34]. Using GPMVs isolated from living cells expressing ANXAs coupled to GFP circumvents the step of protein encapsulation used in studies with artificial vesicles and also allows the curvature sensing to be investigated in a more physiologically-relevant environment with complex lipid and protein composition (See Figure 2A–C). Using optical manipulation, nanotubes were extracted from the vesicles (Figure 2D–F) and protein sorting between the nanotube and the vesicle membrane was quantified by the relative sorting parameter, S:

$$S = \frac{\left(I\_{\text{prot}} / I\_{\text{mem}}\right)\_{\text{tube}}}{\left(I\_{\text{prot}} / I\_{\text{mem}}\right)\_{\text{vesicle}}} \tag{2}$$

where *S* is a measure of the relative density of protein on the tube (*Iprot*)tube versus the quasi-flat surface on the vesicle, (*Iprot*)vesicle. The intensity from a lipid analog in the membrane, (*Imem*)tube scales with the tube diameter and (*Imem*)vesicle is used for normalization to account for possible different concentrations of membrane dye in different vesicles.

As shown in Figure 2E,G the density of ANXA5 was found to increase significantly within nanotubes extracted from GPMVs which were derived from HEK293T cells. The increase in density of ANXA5 ranged up to 10–15 times higher than the density on the quasi-flat region on the vesicle membrane (Figure 2G) although with significant heterogeneity in the sorting as shown in Figure 2E,G,H. Interestingly, the curvature sensing of the cognate protein ANXA2 did show a majority of sorting values below 1 indicating a slightly negative affinity for the curvatures within the nanotubes (Figure 2F–G). These results indicate that proteins from the annexin family can differ remarkably in curvature sensing despite the similarly shaped membrane binding core domain. Furthermore, we recently found that ANXA4 senses membrane curvature with a similar effect as ANXA5 and the curvature sensing was dependent on the ability of ANXA4 to form trimers [36]. The curvature sensing and membrane rolling induced by ANXA4 and ANXA5, but not ANXA2, could together indicate that trimerization is critical for membrane curvature sensing and curvature induction. Beyond these results, future experiments should test whether the two dimensional curvature of annexins can discriminate between spherical and cylindrical curvatures in similar assays as used in Li et al. [37].

**Figure 2.** Membrane curvature sensing of ANXA2 and ANXA5 measured in lipid nanotubes extracted from Giant Plasma Membrane Vesicles (GPMVs). (**A**) GPMVs are derived from HEK293T cells expressing ANXA2-GFP or ANXA5-GFP. (**B**) GFP tagged annexins bind to the inner side of the lipophilic carbocyanine DiD labeled membrane. (**C**) Images of DiD (red) labeled GPMVs containing ANXA5-GFP (green). (**D**) Schematic of optical manipulation of the vesicles to form nanotubes with radius ~50nm and length 10μm. (**E**) Overlay image of a GPMV and nanotube containing GFP tagged ANXA5 and DiD membrane label. (**F**) Overlay image of a GPMV and nanotube containing GFP tagged ANXA2 and membrane label DiD. (**G**) Quantification of curvature sorting for ANXA2 and ANXA5, respectively. The dashed line represents Sorting = 1 corresponding to no sorting. \*\*\* *p* = 0.004. (**H**) The Sorting values from (**G**) plotted as a histogram which reveals significant heterogeneity in the Sorting by ANXA5. Reproduced with permission from [35].

Consistent with the rolling and sensing experiments discussed above multiscale simulations have revealed that ANXA4 trimers both induce and sense membrane curvatures [36]. All-atom (AA) molecular dynamics (MD) simulations provide molecular details of the interactions of annexins with lipid membranes of various compositions. Coarse-grained MD simulations, where multiple heavy atoms in a molecule are represented by single beads, are used to analyze the interactions of multiple annexin molecules with a bilayer surface, and the micron-scale scaffolding of annexins on a membrane surface [38]. Large-scale membrane conformations induced by annexins are probed by Monte-Carlo simulations, where the membrane is modeled as a fluid triangulated surface, and the annexin molecules are modeled as a vector field that induced local curvature changes in the membrane [36]. The three different computational techniques constitute an inter-coupled multi-scale simulation strategy where annexin-membrane interactions are investigated from the molecular level all the way to mesoscale fluctuations in membrane shape.

We used all-atom simulations to calculate the annexin-induced curvature upon POPC (1-Palmitoyl-2-oleoylphosphatidylcholine):POPS (1-Palmitoyl-2-oleoylphosphatidylserine) (4:1 ratio) lipid membranes in the presence of Ca2+. Simulations of ANXA4 reveal that the ANXA4 trimer induces a significant negative average mean curvature of 0.0024 <sup>±</sup> 0.0002 nm−<sup>1</sup> on the bilayer. The curvature is calculated by fitting the coordinates of the lipid head group phosphorus atoms by a two-dimensional Fourier series. The simulations also reveal a significant accumulation of PS lipids below the annexin trimer surface, which is to be expected because Ca+<sup>2</sup> ions cross-link between anionic amino acids on the annexin trimer and anionic POPS lipids [39]. It is worth noting that although such lipid reorganization can be observed in all-atom simulations on time-scales of hundreds of nanoseconds, the diffusion of lipids in the membrane plane is slow, and is best examined by coarse-grained simulations on longer time scales. The trimer of ANXA5 has a similar impact on bilayer properties, while the effect of

monomers of ANXA4 and ANXA5 differ in their interaction with the bilayer surface, compared to their trimer forms (unpublished data). Future studies along the lines described in the previous section are needed to test the theoretical predictions in experimental model systems using GUVs with controlled membrane curvatures and known protein densities.

The curvature calculated from the all-atom MD simulations is fed as a parameter into the Monte-Carlo simulations, which show that ANXA4 trimers accumulate on the inner membrane of highly curved nanotubes, and are desorbed from the less curved membrane surface (Figure 3). The data are in excellent agreement with the curvature-sensing experiments presented in Figure 2, which show a significant increase in the concentration of ANXA5 within nanotubes compared to the flat GPMV surface. The Monte-Carlo method described here simulates the distribution of annexins on a closed membrane surface. We are currently developing a framework of the Monte-Carlo simulations where the effect of annexins onto a free membrane edge can be investigated. Such a setup will open the possibility of investigating the effect of multiple annexins on the shape evolution of a free membrane edge, bringing us even closer to simulating the mechanism of membrane repair.

**Figure 3.** (**A**) An overview of the molecular dynamics (MD) procedure, with a simple form of force field and velocity-verlet integration algorithm. The integration timestep in all-atom MD is usually 2 fs. (**B**) Initial simulation setup of the ANXA4 trimer near a POPC:POPS (4:1) bilayer. (**C**) Final snapshot showing the indentation of the membrane. (**D**) Top view of the final snapshot. (**E**) The 2D curvature profile for a surface passing through the center of the membrane in panel C. (**F**) Monte-Carlo simulation snapshot of ANXA4 protein affinity for a membrane patch (flat) and a highly curved nanotube generated by pulling a vertex from a flat membrane. Note that the proteins are depicted on the outer surface for clarity.

### **5. Annexin Sca**ff**olding**

Some members of the annexin family have been shown to form large scale protein scaffolds at high protein densities. These scaffolds have been imaged by atomic force microscopy (AFM) and consist of a regular lattice of trimers. High-speed AFM has even shown the dynamics of the protein lattice [40]. Protein immobility has also been measured for annexin bound plasma membrane nanotubes indicating that protein lattices can also form in highly curved regions. The functional role of these lattices in membrane repair remains unknown, however it was recently shown that ANXA5 can change the physical properties of the membrane [39]. Additionally, another function could be to provide friction to the membrane and stabilize the membrane rupture against further expansion [41].

### **6. Approaches to Inflict Damage to the Plasma Membrane**

Many cell studies aimed at studying plasma membrane resealing use ablation laser to inflict spatial damage to the cell membrane. Here, a single and localized wound can be induced at the plasma membrane of cultured cells and repair can be followed using fast time-lapse imaging [13,41,42]. The main advantages include user's control over the extent of damage by adjusting the laser power and the ability to assess cellular repair kinetic and monitor the action of fluorescently-tagged proteins involved in repair (Figure 4) [43]. A possible drawback is that membrane damage triggered by UV-ablation laser induces local high temperatures at the injured membrane, which can potentially affect membrane proteins and lipids and create thermally-induced diffusion and denaturation artifacts. However, we and others have demonstrated tempo-spatial recruitment of repair proteins occurs within 10–45s of laser-induced plasma membrane injury [28,44], indicating that the repair machinery is not disabled by the collateral thermal damage. Importantly, complimentary methods to induce membrane injury, such as the use of glass bead injury, detergents, and scraping, show that the same repair proteins are recruited to damaged membrane (including annexins, actin, ESCRT III) as with laser injury and needed for repair [14,45]. This suggests that laser-induced plasma membrane injury can provide insight into repair mechanisms that are of biological significance, since all injuries (artificial and, potentially, physiological/pathological) converge the point of Ca2<sup>+</sup> influx, which is the key stimulus for plasma membrane repair mechanisms.

An alternative method for inflicting cell injury is to use thermoplasmonic [46]. Irradiation of plasmonic nanostructures using near infrared (NIR) light results in an extremely localized temperature increase which can be used to disrupt plasma membranes. This strategy has been used for fusion of cells [47] and membrane vesicles [48]. Both pulsed lasers or continuous wave lasers can be used in combination with plasmonic nanoparticles for disrupting membranes [49–52]. We envision that this technique will provide a fruitful approach for investigation of plasma membrane repair in the future in particular when combined with predictions from theoretical calculation on the stability of membrane holes decorated with annexins scaffolds.

**Figure 4.** Plasma membrane injury inflicted by UV-laser ablation injury. (**A**) Schematic of ablation laser injury to monitor annexin behavior during plasma membrane repair. Ca2<sup>+</sup> influx through the wounded membranes activates and enables annexin to bind and seal the hole by bending membrane and glue membrane edges together. (**B**) Sequential images from time-lapse movie of a MCF-7 breast carcinoma cell showing translocation behavior of ANXA4-RFP to the site of damage (white arrow) upon laser injury.

### **7. Concluding Remarks**

This synergy fostered by interdisciplinary research achieves its full potential when different perspectives are integrated to comprehensively understand a complex cellular phenomenon. Here, we describe the development of new complementary strategies that, when combined, have elucidated mechanisms underlying membrane shaping and repair. Our recent findings, which could only be revealed through interdisciplinary collaboration, show that there is more to annexins than previously anticipated. In particular there are many unanswered questions in relation to the complex interaction between annexins and curved membranes, and how physical cues aid in membrane healing. We surmise that, a combination of novel membrane assays and high resolution imaging together with multiscale simulations will provide major progress in resolving the enigmatic process of membrane repair in the future.

**Author Contributions:** All authors contributed with manuscript writing and editing of this article. All authors have read and agreed to the published version of the manuscript.

**Funding:** Work presented here is supported by the Novo Nordisk Foundation Interdisciplinary Synergy Grant (NNF18OC0034936) and the Lundbeck Foundation (R218-2016-534).

**Acknowledgments:** We thank Dr. Weria Pezeshkian (University of Groningen) for providing the Monte-Carlo simulation snapshot in Figure 3F, and mapping the ANXA all-atom MD into a vertex-based model.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Exploring Biased Agonism at FPR1 as a Means to Encode Danger Sensing** †

**Jieny Gröper 1,2, Gabriele M. König 3, Evi Kostenis 3, Volker Gerke 1,2, Carsten A. Raabe 1,4,\* and Ursula Rescher 1,2,\***


Received: 31 March 2020; Accepted: 21 April 2020; Published: 23 April 2020

**Abstract:** Ligand-based selectivity in signal transduction (biased signaling) is an emerging field of G protein-coupled receptor (GPCR) research and might allow the development of drugs with targeted activation profiles. Human formyl peptide receptor 1 (FPR1) is a GPCR that detects potentially hazardous states characterized by the appearance of N-formylated peptides that originate from either bacteria or mitochondria during tissue destruction; however, the receptor also responds to several non-formylated agonists from various sources. We hypothesized that an additional layer of FPR signaling is encoded by biased agonism, thus allowing the discrimination of the source of threat. We resorted to the comparative analysis of FPR1 agonist-evoked responses across three prototypical GPCR signaling pathways, i.e., the inhibition of cAMP formation, receptor internalization, and ERK activation, and analyzed cellular responses elicited by several bacteria- and mitochondria-derived ligands. We also included the anti-inflammatory annexinA1 peptide Ac2-26 and two synthetic ligands, the W-peptide and the small molecule FPRA14. Compared to the endogenous agonists, the bacterial agonists displayed significantly higher potencies and efficacies. Selective pathway activation was not observed, as both groups were similarly biased towards the inhibition of cAMP formation. The general agonist bias in FPR1 signaling suggests a source-independent pathway selectivity for transmission of pro-inflammatory danger signaling.

**Keywords:** bias analysis; G protein-coupled receptor (GPCR); formyl peptide receptor 1; danger-associated molecular pattern (DAMP); pathogen-associated molecular pattern (PAMP); annexin A1 peptide Ac2-26

### **1. Introduction**

A key feature of the eukaryotic immune defense is its capability to sense danger signals. Microbes are a source of potentially deleterious infections, and consequently, 'pathogen-associated molecular patterns' (PAMPs) and 'danger-associated molecular patterns' (DAMPs) represent chemical signatures that are sensed and transduced via the corresponding 'pattern recognition receptors' (PRRs) [1,2].

One such molecular pattern is the characteristic N-formylated methionine at the N-terminus of bacterial proteins. This modified amino acid is not utilized for the initiation of eukaryotic protein translation. However, mitochondria, which are considered to represent endosymbionts of bacterial origin [3], initiate protein biosynthesis with N-formylated methionine and might release detectable levels of formylated peptides in situations of enhanced cell death or trauma [4,5]. Therefore, the appearance of N-formylated peptides signals potentially hazardous states caused by either bacterial threats (PAMPs) or tissue destruction (DAMPs).

In higher eukaryotes, this unique pattern is sensed by the family of formyl peptide receptors (FPRs), which belong to the superfamily of G protein-coupled receptors (GPCRs) [5,6]. Thus, a shared sensor system detects bacteria-derived ligands and those that are liberated by non-infectious tissue destruction from mitochondria. The corresponding formylated peptides elicit strong signals via the activation of formyl peptide receptor 1 (FPR1) [7,8]. Here, we addressed whether human FPR1 is capable to decode the actual source of threat via different, i.e., ligand-specific signaling, and consequently modify the elicited cellular responses. We hypothesized that these layers of signal information are encoded by distinguishing ligand perception and potentially are governed by biased agonism. This emerging concept in GPCR-mediated signal transduction [9] emphasizes that ligands selectively stabilize specific receptor conformations that ultimately favor one (or more) signaling pathways out of the many the receptor is linked to. The final cellular response elicited by the receptor/ligand interaction is therefore biased towards specific signaling pathways [9–11]. We also considered that the different classes of agonists might group due to their efficacies and potencies for the same cellular pathways, i.e., apart from the selective activation of entirely different signaling pathways. In this scenario, different classes of agonists would not be associated with their own unique profile of qualitatively distinct cellular responses but instead would cause the same effect, as revealed by similar bias factors, yet on a different scale.

In general, agonist-activated GPCRs stimulate a broad set of heterotrimeric αβγ guanine nucleotide-binding proteins (G proteins), which in turn regulate adenylyl cyclase or phospholipase C (PLC) activities, ultimately leading to effective changes in second messengers cyclic AMP (cAMP) and inositol triphosphate (IP3) levels [12]. Apart from regulating enzymes to control the generation of second messengers, key intracellular signaling pathways are also activated via the GPCR signaling axis. For instance, GPCR-linked stimulation of the mitogen-activated protein kinase (MAPK) pathway, which plays important functional roles in, e.g., the regulation of (chronic) inflammation or even the development of cancer [13,14], is well established. Receptor internalization, which is linked with desensitization and signal termination, is regulated via GPCR/agonist interactions. Notably, emerging novel findings suggest that this mechanism might also be linked to prolonged receptor signaling [15,16].

Here, we compared the signaling profiles of representative microbial, endogenous, and synthetic FPR1 agonists. Our results reveal that the FPR1 activators cluster depending on their origin: the bacterial formylated peptides are strong, potent, and efficacious superagonists, whereas the mitochondrial peptides are less effective. Bias calculation uncovered that although the agonists operate on different levels as defined by their logistic parameters, FPR1 signal transmission generally is biased toward inhibition of cAMP formation.

### **2. Materials and Methods**

### *2.1. FPR1 Ligands and Reagents*

Formylated peptides corresponding to the N-termini of the human mitochondrially encoded proteins NADH:ubiquinone oxidoreductase core subunit 2 (MT-ND2; fMNPLAQ), NADH:ubiquinone oxidoreductase core subunit 6 (MT-ND6; fMMYALF), and cytochrome b (CYTB; fMTPMRKTNPLMKLIN), the formylated pentapeptide fMIVIL from *Listeria monocytogenes*, and the gG-2p20 peptide GLLWVEVGGEGPGPT derived from the secreted glycoprotein sgG-2 of herpes simplex virus type 2 (HSV-2) were custom-synthesized (Biomatik, Cambridge, ON, Canada). The acetylated peptide Ac2-26 corresponding to the N-terminus of human annexin A1 (AcAMVSEFLKQAWFIENEEQEYVQTVK) and the synthetic agonist W-peptide (WKYMVm) were purchased from Tocris (Wiesbaden-Nordenstadt, Germany). The prototype FPR1 agonist fMLF derived from *E. coli* was purchased from Sigma. Stock solutions were prepared as indicated in Table A1. The mouse monoclonal anti-FLAG antibody M1 (Sigma-Aldrich, Darmstadt, Germany), which recognizes the FLAG epitope only when present at the very N-terminus of the FPR1 receptor, i.e., after successful cleavage of the hemagglutinin signal sequence (see below) in the endoplasmic reticulum (ER), was labeled with DyLight488 antibody labeling kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer's protocol. Pertussis toxin (PTX) from *Bordetella pertussis* was purchased from Tocris, the Gα<sup>q</sup> inhibitor FR900359 (FR, formerly known as UBO-QIC), a cyclic depsipeptide from the plant *Ardisia crenata sims sims* was purified following a previously published protocol [17]. Reversed-phase high-performance liquid chromatography separation of the FR-containing fraction (column: YMC C18 Hydrosphere, 250 × 4.6 mm, 3 μm; MeOH:H2O (8:2), 0.7 mL min<sup>−</sup>1) afforded FR with a purity of 95%. For G protein inhibition experiments, cells were pretreated for 16 h with 100 ng/mL PTX or for 1 h with 1 μM FR preincubation in cell culture medium at 37 ◦C.

### *2.2. FPR1-Encoding Plasmid, HeLa-FPR1- Cell Line, and Cell Culture Conditions*

The FPR1 expression vector, containing the N-terminally FLAG-tagged human FPR1, was generated as previously described [18]. The FPR1 coding sequence was PCR-amplified from a cDNA library representing human total leukocyte RNA (Takara Bio, Saint-Germain-en-Laye, France). The FLAGepitope was introduced immediately upstream to the original FPR1 start codon and is preceded by a cleavable influenza hemagglutinin signal sequence to facilitate cell surface presentation. This tagged FPR1 CDS was transferred into the mammalian expression vector pcDNA3.1 (-) (Thermo Fisher Scientific) via XhoI and EcoRI restriction sites. HeLa cells cultured in Dulbecco's modified Eagle's medium (DMEM, Sigma), supplemented with 10% standardized fetal bovine serum (FBS Superior, Biochrom, Cambridge, UK), 100 U/mL penicillin, and 0.1 mg/mL streptomycin) at 37 ◦C in a 7% CO2 atmosphere were transfected using Lipofectamine 2000 (Thermo Fisher Scientific). Clonal lines were selected with 800 ng/mL geneticin (G418, AppliChem, Darmstadt, Germany).

### *2.3. FPR1 Expression Analysis in Parental and Recombinant HeLa Cells by qPCR and Immunofluorescence Microscopy*

qRT-PCR was employed to confirm that parental, i.e., non-transfected HeLa cells do not express members of the FPR family at detectable levels and to confirm FPR1 expression in the stably expressing HeLa-FPR1 cell lines. Total RNA from HeLa cells was isolated with the RNeasy mini kit (Qiagen, Hilden, Germany) according to the manufacturers' instructions; 1 μg of RNA starting material was converted into cDNA with the high-capacity cDNA reverse transcription kit and random hexamer primers (Thermo Fisher Scientific). Subsequent qPCR analysis was performed with QuantiTect primer assays (Qiagen) for FPR1 (Hs\_FPR\_1\_SG, QT00199745) and custom-designed sets of primers (Microsynth, Lindau, Germany) for amplification of FPR2 (for: 5 -TTGGTTTCCCTTTCAACTGG-3 rev: 5 -AGACGTAAAGCATGGGGTTG-3 ) and FPR3 (for: 5 -GGTTGAACGTGTTCATTACC -3 rev: 5 -TGGTTTCTGTGAATTTTGGC-3 ). Housekeeping genes actin (Hs\_ACTB\_1\_SG, QT00095431) and glyceraldehyde 3-phosphate dehydrogenase (Hs\_GAPDH\_2\_SG, QT01192646) served as references. All qPCR reactions were conducted with the Brilliant III Ultra-Fast SYBR Green qPCR Master Mix (Agilent Technologies, Santa Clara, CA, USA). Four independent cell samples were analyzed in technical replicates and amplified for 45 cycles on a CFX 384 real-time PCR cycler. The PCR amplification was analyzed with the CFX Manager Software v.2.1 (Bio-Rad, Hercules, CA, USA).

Expression and correct localization of tagged FPR1 were confirmed by immunofluorescence imaging. HeLa-FPR1 cells were cultured on glass coverslips and fixed with 4% paraformaldehyde for 10 min at room temperature. After incubation with anti-FLAG M1 antibody (diluted 1:100 in 2% BSA in PBS containing Ca2<sup>+</sup> and Mg2<sup>+</sup> (PBS++, Sigma) for 60 min, cells were treated with anti-mouse Alexa 594-coupled secondary antibody (Invitrogen, Carlsbad, CA, USA) for an additional 45 min at room

temperature. To visualize cell nuclei, Hoechst 33,342 stain (Thermo Fisher Scientific, diluted 1:100) was added during the incubation with the secondary antibody. Oregon Green 488 conjugated wheat germ agglutinin (Invitrogen, WGA, 5 μg/mL in Hank's balanced salt solution containing Ca2<sup>+</sup> and Mg2<sup>+</sup> for 10 min at room temperature) was used to label the plasma membrane. Samples were imaged with a LSM800 (Zeiss, Oberkochen, Germany) confocal microscope using a 63× objective.

### *2.4. Flow Cytometric Analysis of Agonist-Induced Receptor Internalization*

HeLa-FPR1 cells were treated with vehicle or agonists diluted in internalization medium (IM; DMEM, 20 mM HEPES, 1 mg/mL BSA, pH 7.2) for 15 min at 37 ◦C. Cells were washed in PBS (Sigma), and detached in PBS/5 mM EDTA for 3 min at 37 ◦C. For the detection of the cell surface receptor pool, cells were washed with ice-cold PBS containing 5% BSA and 1 mM CaCl2 and subsequently incubated with 5 μg/mL DyLight488-conjugated anti-FLAG M1 antibody for 45 min. 7AAD (eBioscience, San Diego, CA, USA) allowed the exclusion of compromised cells. Median fluorescence intensities (MFI) of 10,000 viable cells per condition were measured with a Guava easyCyte flow cytometer and the InCyteTM Software (Merck-Millipore, Darmstadt, Germany). Agonist-induced internalization was defined as the difference between the MFI of vehicle-treated controls and the MFI detected in agonist-treated cells. For each measurement, agonist-induced internalization was normalized to the mean internalization induced by 10−<sup>4</sup> M W-peptide, which consistently represented the maximum system response for internalization.

### *2.5. HTRF-Based Quantification of cAMP Levels*

A competitive immunoassay based on time-resolved measurement of fluorescence resonance energy transfer (HTRF, homogeneous time-resolved fluorescence) between a cryptate-labeled specific antibody (donor) and a d2-coupled cAMP acceptor molecule (cAMP-Gi Kit, Cisbio, Codolet, France) was used to measure cyclic AMP (cAMP) formation in cells, as described elsewhere earlier [19]. In brief, cells cultured as described above on 96 well plates (50k cells/well) were incubated with 5 μM of the adenylyl cyclase activator forskolin (Sigma) in complete medium supplemented with 500 μM of the phosphodiesterase inhibitor 3-isobutyl-1-methylxanthine (IBMX, Sigma), together with the agonists at the indicated concentrations, for 30 min at 37 ◦C. Samples were transferred onto a 384 well low volume plate, conjugates were added, and samples were analyzed with the CLARIOstar reader (BMG Labtech, Ortenberg, Germany) (200 flashes/well, integration start 60 μsec, integration time 400 μsec, settling time 100 μsec). Luminescence signals were expressed as the ratio of 10,000× (acceptor signal/donor signal). Values were normalized to the maximum system output, which was obtained as forskolin-induced cAMP formation.

### *2.6. HTRF-Based Quantification of MAPK*/*ERK Phosphorylation Levels*

The advanced Phospho-ERK1/2 (Thr202/Tyr204) plate-based assay (Cisbio) was used to measure ERK activation through HTRF using a sandwich assay format of a donor-coupled antibody and an acceptor antibody as described before in [20]. Briefly, cells grown on 96 well plates (45k cells/well) were serum-starved for 2 h and stimulated with the agonists at the indicated concentrations for 5 min at 37 ◦C. Lysates were transferred to a 384 well low volume plate, incubated with the antibodies for 4 h at room temperature, and luminescence was recorded with a CLARIOstar reader (BMG Labtech) (200 flashes/well, integration start 60 μsec, integration time 400 μsec, settling time 100 μsec). HTRF ratios were normalized to maximum system output obtained through stimulation with 100 nM phorbol 12-myristate 13-acetate (PMA).

### *2.7. HTRF-Based Quantification of IP1 Levels*

The IP-One-Gq assay (Cisbio), that detects inositol monophosphate, a stable downstream metabolite of IP3 induced by phospholipase C activation, was utilized to establish FR pretreatment conditions, as described elsewhere earlier [21]. In brief, cells grown overnight on 384 well plates (5k cells/well) were pretreated with 1 μM FR for 1 h at 37 ◦C prior to stimulation with 100 μM phospholipase C activator 2,4,6-trimethyl-N-[3-(trifluoromethyl)phenyl]benzenesulfonamide (m-3M3FBS, Tocris) for 2 h at 37 ◦C. LiCl present in the stimulation buffer provided in the kit prevented the degradation of IP1. After addition of the conjugates, samples were incubated for 1 h at room temperature and read on a CLARIOstar plate reader (BMG Labtech) (200 flashes/well, integration start 60 μsec, integration time 400 μsec, settling time 100 μsec).

### *2.8. Curve Fitting*

For each individual curve, ligand concentrations were log-transformed, normalized, and expressed as fractions of the maximum system response per pathway. Concentration-response curves were analyzed with Graphpad Prism 6 and the in-built four parameters sigmoidal model with a Hill coefficient of 1. EC50 values were derived from individual sigmoidal curve fits. If the data fitting was ambiguous without further constraints (very weak partial agonists, no inflection point), the minimum was set to zero and the highest experimental response was considered to represent Emax. The ROUT method [22] was used to detect and eliminate outliers. An agonist-induced response was defined by a curve span >3 SEM. If no fitting was possible (i.e., no detection of agonist-elicited responses), the 'no response' (NR) label was assigned. To analyze the relationship between data, linear regression and Spearman's correlation coefficients were calculated (Graphpad Prism 8).

Bias was calculated with the operational model [23]. The logarithm of the activity ratio *E*max/EC50 [24–26] served as a surrogate for the actual transduction coefficient log(t/KA) [27] and was calculated individually for each agonist and pathway. To visualize the ligand "texture", agonist-specific ΔlogR values were plotted for the analyzed pathways. To compare the relative effectiveness of an agonist at a given pathway with the reference agonist, ΔlogR values were calculated as differences of the respective pathway-specific logR of the agonist of interest and the endogenous MT-ND6 peptide as the reference agonist. To rank the relative signaling preferences of a ligand for one pathway over another, ΔΔlogR values were calculated.

### **3. Results**

To address in a systematic fashion whether FPR1 agonists differ in their corresponding signaling fingerprints, we established a heterologous FPR1 expression system [18]. The resulting transgenic HeLa-FPR1 cell line was analyzed via qRT-PCR to confirm the stable expression of FPR1 (Figure 1a). As parental HeLa cells did not express any detectable levels of FPR1-3, this experimental design enabled monitoring specific FPR1-mediated responses [8,28]. The correct localization and incorporation of the FLAG-tagged receptor in the plasma membrane were validated by immunofluorescence imaging (Figure 1b). In order to avoid potentially distorting influences resulting from agonist effects other than those elicited via the FPR1 signaling axis, we used parental HeLa cells as a negative control. No signal could be observed when these cells were stained with the M1 antibody, and none of the tested agonists caused any significant response for the analyzed pathways, thereby confirming the specificity of our results (Figure A1).

### *3.1. FRET-Based Analysis of Agonist-Induced Changes in cAMP Levels*

Agonist-induced cellular responses mediated via FPR1 previously have been reported to depend on Gα<sup>i</sup> for signal transduction [29]. We, therefore, resorted to a FRET-based system to analyze agonist-mediated changes in cAMP de novo generation. Notably, none of the analyzed FPR1 agonists caused any detectable increase of cellular cAMP levels, thereby confirming that the tested ligands did not trigger FPR1 responses via Gα<sup>s</sup> (Figure A2). To stimulate the cellular cAMP formation to its maximum, we utilized forskolin, a known activator of adenylyl cyclase activity [30] and determined the inhibitory potential of the different FPR1 agonists (exemplary raw data are shown in Figure A3a). Relative potencies and efficacies of the analyzed agonists differed significantly in our assay (Figure 2). Bacterial peptides [31,32] and the W-peptide [15,33,34], formed a distinct group of agonists with

high potencies and efficacies, mitochondrial peptides—on the other hand—were characterized by significantly lower potencies. However, differences in the respective efficacies for bacterial and mitochondrial peptides were not as pronounced. Notably, the annexin A1 peptide Ac2-26 [35], which displayed the lowest potency in our assay, was still able to suppress de novo cAMP formation to a level similar to the mitochondrial agonists. Interestingly, gG2p20, an exogenous ligand-derived from herpes virus [36], and the synthetic small molecule FPRA14 [37,38] mimicked the mitochondrial peptides to some extent. To analyze agonist profiles in more mechanistic detail, we investigated whether the magnitude of the agonist-elicited responses was linked with the respective potencies. Generally, EC50 values of the agonists displayed a very strong negative monotonic relationship with their respective ability to elicit Emax, as demonstrated by the Spearman's rank correlation coefficient (r = −0.917, *p* = 0.001, n 9).

**Figure 1.** Evaluation of FPR1 (formyl peptide receptor 1) expression and localization in the heterologous HeLa expression system. (**a**) qRT-PCR revealed that none of the three FPR paralogs is expressed in parental HeLa cells (WT), whereas FPR1 is readily detectable in the HeLa-FPR1 transgenic cell line. Housekeeping genes ACTB (beta actin) and GAPDH (glyceraldehyde 3-phosphate dehydrogenase) served as internal controls. (**b**) Confocal immunofluorescence microscopy with the M1 antibody (red channel) confirmed the plasma membrane localization of FLAG-tagged FPR1. The plasma membrane was stained with WGA (green channel), and nuclei were visualized using Hoechst stain (blue channel). Upper panel: HeLa-FPR1 cell line; lower panel: WT HeLa cells, scale bar, 5 μm.

**Figure 2.** Concentration–response curves for FPR1-mediated inhibition of cAMP production. Cells were stimulated with forskolin (FSK) and the respective FPR1 agonists. Responses recorded 30 min after agonist addition were normalized to the maximum system response obtained with forskolin. Data points represent the mean ± SEM of at least 7 independent measurements.

### *3.2. FRET-Based Analysis of Agonist-Induced Changes in MAPKinase Activation*

The MAPKinase cascade is a common effector of GPCR activation [39–41]. As shown in Figure 3, all agonists were able to increase ERK1/2 phosphorylation (exemplary raw data are shown in Figure A3b). Interestingly, the mitochondrial agonist MT-ND6 performed comparably to the bacterial agonists and the W-peptide, whereas the other mitochondrial peptides, as well as Ac2-26, only weakly activated ERK1/2 phosphorylation. As in the case of the adenylyl cyclase inhibition, gG2p20 and the synthetic small molecule FPRA14 displayed almost identical profiles. Spearman's rank correlation uncovered a strong negative monotonic correlation between EC50 and Emax values (r = −0.8, *p* = 0.014, n 9).

**Figure 3.** Concentration–response curves for FPR1-mediated phosphorylation of ERK1/2 on Thr202/Tyr204. Responses monitored 5 min after the addition of agonist were normalized to the maximum system response, which was obtained with PMA (phorbol 12-myristate 13-acetate). Data points represent the mean ± SEM of at least 5 independent measurements.

### *3.3. Agonist-Mediated Internalization of FPR1*

Activated GPCRs usually are removed from the cell surface and are internalized into endosomes [15,16]. To test the potential of our agonists to induce FPR1 internalization, we determined the agonist-dependent decrease in FPR1 cell surface presentation, based on the detection of the N-terminal FLAG-epitope by flow cytometry [42,43]. The actual amount of internalized receptor was calculated via the difference of cell surface signals measured in untreated and agonist-treated samples after 15 min of agonist addition. Administration of W-peptide, bacterial peptides fMLF and fMIVIL, or the endogenous mitochondrial peptide MT-ND6 decreased the cell surface-associated FPR1 pool in a concentration-dependent manner. In contrast to that, the mitochondria-derived peptides MT-ND2, CYTB, the synthetic agonist FPRA14, and the annexin A1 peptide Ac2-26 did not elicit detectable internalization and were therefore deemed non-responders. Importantly, no significant difference was observed between the two formylated ligands fMLF and MT-ND6, indicating that the source of signal (PAMP vs. DAMP) was not encoded by these ligands (Figure 4).

**Figure 4.** Concentration–response curves for agonist-induced FPR1 internalization. Receptor internalization was calculated as the difference between FLAG signals of unstimulated cells and signals recorded 15 min after agonist addition. Results were normalized to the maximum system response obtained with 10−<sup>4</sup> M W-peptide. Data points represent the mean <sup>±</sup> SEM of 6 independent measurements.

### *3.4. G protein Dependency of Agonist-Mediated Responses*

We considered that the correlation for Emax and EC50 values for individual agonists and pathways, as revealed by the Spearman's rank coefficients, might—albeit indirectly—provide insights into common effector proteins interacting with the receptor to transduce agonist-elicited responses. We detected a very strong positive monotonic correlation between EC50 values for cAMP inhibition and ERK phosphorylation (r = 0.933, *p* = 0.001, n 9). A similar trend could also be established for the corresponding Emax values, which again revealed a strong positive correlation (0.733, *p* = 0.031, n 9; for an overview of logEC50 and EmaxA values, see Figure A4 and Table A2.). To directly assess whether the different molecular pathways are driven by Gαi-protein-dependent signal transduction, we repeated our experiments in the presence of the Gαi-specific inhibitor pertussis toxin (PTX) at a concentration that completely abolished the inhibition of cAMP formation induced by W-peptide (Figure A4a). PTX treatment of HeLa-FPR1 cells did not interfere with ERK activation per se, however, agonist-mediated ERK1/2 activation was suppressed, thus strongly implying that both pathways, (at least) in our cellular system, relied on the Gαi-protein (Figure 5). In stark contrast, receptor internalization was not dependent on Gα<sup>i</sup> activation, as agonist-evoked FPR1 internalization was not affected in PTX-pretreated cells (Figure 6a) and was equally undisturbed in cells pretreated with the Gα<sup>q</sup> inhibitor FR (Figure 6b).

**Figure 5.** FPR1-mediated ERK1/2 phosphorylation is abolished in PTX (pertussis toxin)-pretreated cells. Cells were pretreated for 16 h with 100 ng/mL PTX and subsequently stimulated with agonist concentrations eliciting the respective Emax. Results were normalized to the maximum system response obtained with PMA. Data points represent the mean ± SEM of 6 independent measurements.

**Figure 6.** Agonist-induced FPR1 internalization is not dependent on Gα-mediated signal transmission. (**a**) Cells were pretreated either for 16 h with PTX or (**b**) for 1 h with FR and subsequently stimulated with agonist concentrations eliciting the respective Emax. Results were normalized to the 10−<sup>4</sup> M W-peptide-induced maximum response. Data points represent the mean <sup>±</sup> SEM of 6 independent measurements.

### *3.5. Analysis of Agonist Bias on FPR1*

For the identification and analysis of ligand bias [23] associated with the FPR1 signaling axis, we selected the mitochondria-derived MT-ND6 as endogenous reference agonist for all tested pathways. For each agonist and pathway, we determined the corresponding logR (the logarithm of the activity ratio) values [24] and normalized the activities to MT-ND6 (ΔlogR=logRagonist-logRMT-ND6, see Table I) [23]. Figure 7 summarizes the resulting agonist ranking for each pathway. Overall, two agonist clusters were established: bacterial ligands, along with the W-peptide, which resembles a conserved spatial structure of bacterial agonists [44], constituted the group of "high-performers". Endogenous mitochondria-derived agonists, the herpes virus-derived peptide, the small synthetic agonist FPRA14, and annexin A1 peptide Ac2-26 represented the second group of considerably lower activity (Figure 7a). Interestingly, the endogenous agonist MT-ND6 featured balanced characteristics, with a profile "in-between" both groups. Graphical representations of the intrinsic activity profiles, i.e., in terms of ΔlogR, further highlight the tendency of our tested ligands to segregate into two clusters, which was observed independently of pathway but in relation to the agonist origin (Figure 7b). However, the segregation was less pronounced when the magnitude of the respective response was taken into consideration (Figure 8, Figure A5, and Table A2). Analysis of pathway preferences, i.e., in terms of ΔlogR values, revealed that FPR1 activation was generally biased towards the inhibition of cAMP formation compared to the activation of the MAPKinase pathway or receptor internalization (Figure 9). However, the direct comparison of the weak agonist Ac2-26 with the strong agonist fMIVIL impressively revealed that bias was realized at different logistic levels (Figure 9).

**Figure 7.** Relative agonist profiles compared to the endogenous reference agonist. (**a**) ΔlogR values were calculated from logR values with the mitochondrial peptide MT-ND6 as reference agonist. Data points represent the mean and 95% confidence intervals. (**b**) Relative agonist activities corresponding to the ΔlogR values for the respective pathways are depicted in radial graphs. Each radius is displayed in the logarithmic scale.

**Figure 8.** Graphical representation of the agonist profiles. Relative agonist activities corresponding to the ΔlogR values and the logistic parameters (**a**) EC50 and (**b**) EmaxA for the respective pathways are depicted in radial graphs. Each radius for ΔlogR is displayed in the logarithmic scale, radiuses for Emax are scaled linearly.

**Figure 9.** Comparison of agonist pathway preferences. ΔΔlogR values were calculated from ΔlogR values. Data points represent the mean and 95% confidence intervals.

### **4. Discussion**

FPR1 and the corresponding large repertoire of FPR1 agonists, derived from various cellular and pathogenic sources, constitute a powerful system for the detection of insults that are linked with tissue destruction under infectious and sterile conditions, such as pathogenic challenges or trauma caused by burns or injury [9,45]. Because overactivation of the innate immune response is often correlated with excessive and deleterious tissue damage, e.g., in influenza A virus (IAV) infection, targeting the FPR family might represent a novel approach to balance innate immunity. Indeed, our previous studies revealed the advantageous use of the host-derived anti-inflammatory N-terminal annexin A1 peptide Ac-2-26, a pan-FPR agonist [46] to counteract viral load and mortality in a preclinical murine IAV infection model [47], thus encouraging the development of novel FPR-based therapies.

Generally, the often observed signaling diversity elicited by GPCR agonists that are acting on the same receptor is based on the ligand preference for certain receptor conformational states linked to a subset of all possible signaling responses. This diversity has been termed "functional selectivity" and has led to the concept of "biased agonism" and helps to explain different regulatory outcomes via the activation of the same receptor [9,11]. Therapeutically, bias analysis might help to identify compounds that direct receptor signaling toward desired responses, thus aiding in the development of novel therapeutics with an effective pharmacological profile that avoids activation of unwanted signaling pathways and hence side effects [48,49]. Indeed, a few biased agonists have been developed and are currently in various stages of clinical trials; this is true even for FPR-targeting compounds [50].

To further explore the potential of biased agonism acting on FPRs, we selected human FPR1, the founding member of the FPR family [8], for broader analysis. FPR1 represents the sensor platform for short formylated peptides, a pattern commonly associated with bacterial PAMPs and mitochondria-derived DAMPs [51,52]. However, the preference for such modified peptides is not exclusive, and even non-formylated derivatives (such as Ac2-26) are known to activate FPR1 effectively. The structural diversity of FPR1 agonists led us to hypothesize that potentially different classes of agonists might group, based on their origin, i.e., as PAMPs, otherwise endogenous ligands or DAMPs, thus enabling the receptor to decode the actual source of danger and in turn to channel the cellular responses.

Commonly, agonists are classified according to their ability to invoke the maximum receptormediated response of a given pathway. These agonists represent "full" agonists, whereas "partial" agonists only elicit a fraction of the cellular responses caused by a full agonist. This classification is intuitive and seems to be well suited to describe the properties of most ligands; it inherently suffers, however, from the disadvantage that potentially better—yet unidentified—agonists (evoking a higher response) cannot be accounted for. An alternative approach classifies agonist efficacies in relation to an endogenous reference agonist of high efficacy. Therefore, some agonists might be identified, which are even capable to elicit stronger cellular responses than those associated with the reference agonist; consequently, these ligands are referred to as "superagonists", although this term still has to be defined in broader detail [53]. The molecular explanation for this phenomenon might lie in the observation that

simultaneous binding of an efficacious agonist and a G protein is required to induce the full receptor response [54,55]. Our results imply that bacterial agonists function as bona fide superagonists at FPR1.

Of note, FPR1 agonist clusters clearly grouped based on ligand efficacies and potencies; these logistic properties were strongly correlated as revealed by the analysis "within" as well as "across" Gαi-dependent pathways. Moreover, our results argued in favor of a shared signal transmission selectivity for a given pathway across these structurally unrelated agonist classes. This was most obvious in the case of agonist-mediated ERK activation [56,57], which was entirely G protein-dependent, because in no instance did we observe Gαi-independent activation of the ERK signaling cascade. Yet, lack of evidence for G protein-independent signals, as far as ERK phosphorylation is concerned, does by no means exclude that FPR1 agonists display preferences for supposedly distinct subsets of FPR1-elicited signaling pathways over others. Internalization might—at first glance—appear G protein-independent. However, the most likely scenario probably is that endocytosis requires an active receptor conformation rather than an active signaling pathway and therefore occurs even when G proteins are precluded from active signal transmission. It is therefore perhaps not surprising that particularly high-efficacy ligands are the most effective at causing receptor internalization. It also cannot be ruled out that a given cellular environment is hard-wired for a set of pathways [58]. Hence, the decoding capacity of our heterologous expression system might cause the cells' inability to distinguish the source of these ligands, leading to so-called "system bias". Our data suggests that peptide-based FPR1 pharmacotherapy might be worthwhile exploring, however, peptide aggregation is a huge challenge [59].

Based on the bias calculations, all of the agonists preferentially inhibited cAMP over ERK phosphorylation. cAMP is a prominent second messenger in the PKA signaling pathway and a potent regulator of immunity. Because increasing and decreasing cAMP levels are correlated to dampening or stimulating immune responses, respectively, the cellular cAMP balance is considered a bona fide druggable target [60]. Surprisingly, the Ac2-26 peptide was also biased toward adenylyl cyclase inhibition, similar to the endogenous mitochondrial peptides, and the agonistic profile did not resonate with the established anti-inflammatory properties.

The bias in FPR1 activation toward inhibition of cAMP production may signal imminent danger, regardless of the source. However, similar bias factors can be associated with entirely different logistic parameters, and therefore, might cause different levels of physiological response. Our data do not support a selective activation of entirely different signaling pathways, as typically associated with biased agonists. Rather, our findings identified that the actual differences of the danger signals are encoded by the different levels of response (described by the logistic parameters EC50 and the maximum agonist-elicited response Emax).

Another means by which GPCR signaling can be diversified is the organization of GPCR-based signaling platforms via homo- and hetero-oligomerization [61]. Indeed, there is emerging functional evidence for FPR higher-order structures [62,63]. Whether such supramolecular sensor complexes are able to decode additional information is certainly an important line of future research.

**Author Contributions:** Conceptualization, C.A.R. and U.R.; methodology, J.G., C.R. and U.R.; validation, C.A.R., J.G. and U.R.; formal analysis, J.G.; investigation, J.G.; resources, U.R., G.M.K., E.K.; writing—original draft preparation and methodology, J.G., V.G., C.A.R. and U.R.; writing—review and editing, J.G., C.A.R., U.R., G.M.K., E.K.; visualization, J.G.; supervision, C.A.R.; project administration, U.R.; funding acquisition, V.G., U.R. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the GERMAN RESEARCH FOUNDATION (DFG), CRC1009 "Breaking Barriers", Project A06 (VG., U.R.), and CRC 1348 "Dynamic Cellular Interfaces", Project A11 (U.R). E.K. and G.M.K. were supported by the DFG-funded Research Unit FOR2372 with the grants KO 1582/10-1 and KO 1582/10-2 (to E.K), as well as KO 902/17-1 and KO 902/17-2 (to G.M.K.).

**Acknowledgments:** We thank Rod Flower, Terry Kenakin, and Henry Showell for thought-provoking impulses, insightful discussions and comments.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

**Appendix A**

**Figure A1.** Agonist effects are specific for the FPR1 signaling axis. Parental HeLa cells were subjected to the assays analyzing (**a**) internalization, (**b**) cAMP formation, and (**c**) MAPKinase activation, at agonist concentrations eliciting the respective Emax response. (**a**) A representative result, (**b**), (**c**) mean response ± SEM, n 6.

**Figure A2.** Concentration-response curves for FPR1-mediated stimulation of cAMP production. None of the agonists increased intracellular cAMP levels, in line with the reported FPR1-mediated activation of Gαi. Mean response ± SEM, n 3.

**Figure A3.** Exemplary raw data of concentration-response curves for FPR1-mediated stimulation of (**a**) cAMP production and (**b**) MAPKinase activation. In addition to the W-peptide measurements, data points for the cAMP standards and the ERK positive and negative controls included in the kit are shown. Data were normalized to the maximum output as obtained with forskolin and PMA.

**Figure A4.** Analysis of G protein dependency. (**a**) PTX-mediated inhibition of FPR1-mediated changes in cAMP generation. (**a**) Maximum FPR1-mediated inhibition of forskolin-induced cAMP formation was completely prevented in cells pretreated for 16 h with 100 ng/mL PTX. (**b**) Effect of 1 μM FR preincubation for 1 h on IP-1 production induced by the phospholipase C activator m-3M3FBS. Data points represent the mean ± SEM.

**Figure A5.** Depiction of logEC50 and EmaxA values of each pathway. Ligands are arranged according to the intensity of the response they elicited in the respective pathway. Data are expressed as mean ± SEM after outlier removal.


**Table A1.** Ligand stock solutions. Concentrations of stock solutions were calculated based on the solubility of the compound as indicated by the manufacturer and its molecular weight (M). Stock solutions were prepared using the solvents recommended by the manufacturer.



### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Review* **Annexin A2 in Inflammation and Host Defense**

### **Valentina Dallacasagrande and Katherine A. Hajjar \***

Department of Pediatrics, Weill Cornell Medicine, 1300 York Avenue, New York, NY 10065, USA; vad2010@med.cornell.edu

**\*** Correspondence: khajjar@med.cornell.edu

Received: 31 May 2020; Accepted: 17 June 2020; Published: 19 June 2020

**Abstract:** Annexin A2 (AnxA2) is a multifunctional calcium2<sup>+</sup> (Ca2+) and phospholipid-binding protein that is expressed in a wide spectrum of cells, including those participating in the inflammatory response. In acute inflammation, the interaction of AnxA2 with actin and adherens junction VE-cadherins underlies its role in regulating vascular integrity. In addition, its contribution to endosomal membrane repair impacts several aspects of inflammatory regulation, including lysosome repair, which regulates inflammasome activation, and autophagosome biogenesis, which is essential for macroautophagy. On the other hand, AnxA2 may be co-opted to promote adhesion, entry, and propagation of bacteria or viruses into host cells. In the later stages of acute inflammation, AnxA2 contributes to the initiation of angiogenesis, which promotes tissue repair, but, when dysregulated, may also accompany chronic inflammation. AnxA2 is overexpressed in malignancies, such as breast cancer and glioblastoma, and likely contributes to cancer progression in the context of an inflammatory microenvironment. We conclude that annexin AnxA2 normally fulfills a spectrum of anti-inflammatory functions in the setting of both acute and chronic inflammation but may contribute to disease states in settings of disordered homeostasis.

**Keywords:** annexin A2; inflammation; infection; adherens junction; angiogenesis; macroautophagy

### **1. Introduction**

Inflammation is defined as the local host response to injury caused by infectious or noninfectious agents; it results in elimination or compartmentalization of the inciting agent, clearance of necrotic cells, and repair of damaged tissue [1,2]. Initially, host cells recognize danger-associated molecular patterns (DAMPs) or pathogen-associated molecular patterns (PAMPs) through germline-encoded pattern-recognition receptors (PRRs) expressed mainly in monocytes, macrophages, neutrophils, and dendritic cells [1]. The four known classes of PRR molecules include transmembrane proteins, such as Toll-like (TLRs) and C-type lectin receptors (CLRs), as well as cytoplasmic proteins, such as retinoic acid-inducible gene (RIG)-like receptors (RLRs) and nucleotide-binding oligomerization domain-like receptors (NLRs) [3]. These pathways induce the release of proinflammatory cytokines, which increase vascular permeability, thus facilitating the extravasation of immune cells into the damaged tissue, and chemokines, which recruit supplemental immune cells that phagocytose and kill pathogens [1]. Inflammation that resolves within days is classified as acute, whereas chronic inflammation may persist for months to years. Chronic inflammation underlies the pathogenesis of an array of disease entities, including diabetes, asthma, arthritis, cancer, atherosclerosis, vasculitis, and inflammatory bowel disease.

The annexins are Ca2+-regulated, phospholipid- and membrane-binding proteins that are named after the Greek word "*annex*", meaning to attach or bridge because of their ability to link membranes to other membranes or other structures [4]. All but one (A6) of the twelve annexin family members (A1–A11 and A13) identified in vertebrates have a highly homologous core domain (~30–35 kilodaltons) containing four multi-alpha helical repeats with potential Ca2<sup>+</sup>-binding activity and an N-terminal

domain (~3 kilodaltons), which is specific for each family member [5,6]. The annexins are expressed ubiquitously throughout the phylogenetic tree and are evolutionarily ancient.

Annexin A2 (AnxA2) is one of the most extensively studied members of the annexin superfamily [6]. AnxA2 is produced by a wide spectrum of cell types, including endothelial, trophoblast, epithelial, and tumor cells, as well as innate immune cells, such as macrophages, monocytes, and dendritic cells. AnxA2 may exist in either monomeric or heterotetrameric form. The heterotetramer (A2•S100A10)2 is composed of two copies each of AnxA2 and protein S100A10, also called p11. AnxA2 is present in the cytoplasm and on cell surfaces, and its functions are largely location specific. On the endothelial cell surface, the (A2•S100A10)2 complex binds components of the fibrinolytic system, plasminogen and tissue plasminogen activator (tPA), accelerating the activation of the serine protease plasmin [7,8]. As the primary fibrinolytic protease, plasmin enables fibrin breakdown and angiogenesis [9,10]. Intracellularly, AnxA2 seems to fulfill many functions, including organization of specialized membrane microdomains, facilitation of vesicle budding, and regulation of additional membrane dynamic events such as fusion, endocytosis, endosomal biogenesis, and membrane repair [4,5,11–21].

In view of this wide array of functions, we have developed a working model depicting the disparate anti- and pro-inflammatory roles of AnxA2 at the various stages of inflammation (Figure 1). In the initial phase, AnxA2 limits vascular permeability, thereby modulating recruitment of leukocytes and their subsequent release of inflammatory mediators. AnxA2 also supports internal membrane repair, thus modulating inflammasome activation, and participates in the biogenesis of the phagophore in autophagy, thereby facilitating the removal of pathogens and damaged organelles. Later, AnxA2 enables angiogenesis and tissue healing.

**Figure 1.** Annexin A2 in inflammatory responses—working model. Under homeostatic conditions (top arrow), AnxA2 plays an anti-inflammatory role in response to injury or infection. In the immediate response to injury, AnxA2 maintains vascular integrity, thereby preventing edema and extravasation of blood cells. Within minutes to hours, AnxA2 also protects internal membranes, such as those delimiting the lysosome, thus preventing inflammasome activation and tempering cytokine production. AnxA2 is also necessary for biogenesis of the phagophore, a double membrane structure that engulfs and destroys intracellular pathogens and allows the cell to adapt to environmental stresses in the process of macroautophagy. In the later stages of inflammation, AnxA2 likely promotes angiogenesis and wound healing by supporting cell surface fibrinolytic activity. In disease states, however (bottom arrow), AnxA2 may be co-opted to perform pro-inflammatory actions. On the plasma membrane, AnxA2 appears to serve as a site for bacterial adhesion and entry into cells, and may enable viral infection, replication, and release. In chronic inflammatory states, excessive angiogenesis that is sustained by AnxA2 may induce tissue damage, as in the diabetic retina, or may support the progression of cancer. In addition, some tumor cells may utilize cell surface AnxA2 to generate protease activity necessary for migration and metastasis within a pro-inflammatory micromilieu.

### **2. Annexin A2 and Infection**

Infection results from the interaction of a pathogen (e.g., bacterium, fungus, or virus) with a host organism, leading to tissue invasion, proliferation of the pathogen, production of toxins, and cellular damage [22,23]. The first response of the mammalian host to infection is inflammation, mediated by the innate immune system, and is not specific to the inciting agent. The ensuing adaptive immune response is highly specific to the invading pathogen and typically provides long-lasting protection [23]. Annexin A2 fulfills a number of protective roles during pathogenic infection (Table 1).

In a model of *Klebsiella* pneumonia in mice, AnxA2 appeared to reduce infection-associated inflammation [24]. In *Anxa2-*/*-* mice infected intranasally with *Klebsiella* pneumonia, the level of pro-inflammatory cytokines was significantly elevated, and *Anxa2-*/*-* mice exhibited 100% mortality, versus 100% survival for *Anxa2*+/+ mice at 50 h post injection. Additionally, peritoneal macrophages lacking AnxA2 showed worsening of the TLR4-triggered inflammatory response. These investigators concluded that AnxA2 has a role in limiting inflammation by promoting anti-inflammatory signals [24].

Annexin A2 also protects the host in a murine model of*Cryptococcal* infection.*Cryptococcus neoformans* is an encapsulated budding yeast that, unlike others, can replicate under acidic conditions; its virulence depends upon its interaction with host macrophages [25]. In this infection, *Anxa2-*/*-* bone-marrow-derived macrophages were less efficient than control macrophages at phagocytosing yeast cells, resulting in a lower frequency of non-lytic exocytosis. In addition, the *Cryptococcus* capsule was enlarged in *Anxa2-*/*-* -deficient macrophages, possibly impacting nonlytic exocytosis, and potentially reflecting the ability of AnxA2 to promote intracellular membrane interactions and vesicle adhesion to the cell membrane [6,26–31]. In vivo, mice lacking AnxA2 showed a lower survival rate when infected with *Cryptococcus*, reflecting a dysregulated inflammatory response [25].

On the other hand, there is evidence that cell surface annexin AnxA2 may be co-opted to facilitate infection by invading bacteria such as *Pseudomonas aeruginosa* [32], *Escherichia coli* [33], *Salmonella typhimurium* [34], and *Rickettsial* species [35]. *P. aeruginosa*, an organism that causes chronic pulmonary infection in patients with cystic fibrosis, anchors to AnxA2 on respiratory epithelial cells, undergoes internalization, and causes apoptotic cell death and release of pro-inflammatory cytokines [32]. In this instance, AnxA2 enables the initiation of productive infection.

Similarly, *E. coli*, particularly the EspL2 strain, reorganizes the cytoskeleton of the host epithelial cells with the participation of annexin A2, which is known to regulate actin dynamics at the bacterium–membrane contact site [36,37]. AnxA2 is recruited to the membrane adhesion site by *E. coli*, where its F-actin bundling activity is enhanced [33,38]. AnxA2 also promotes cytoskeletal rearrangements during host cell invasion by *Salmonella. S. typhimurium*, a common cause of gastroenteritis, uses the contact-dependent type 3 secretion system of the host cell to extensively remodel actin. AnxA2 is engaged in various mechanisms driven by actin rearrangement (e.g., endocytosis, cell–cell adhesion, and membrane ruffling) [34]. AnxA2 participates in reorganization of the actin cytoskeleton during *Salmonella* invasion; depletion of annexin A2 and S100A10 reduced bacteria invasion, probably because of the ability of these proteins to bind the large phosphoprotein AHNAK at the bacteria entry site [34].

In addition, annexin A2 acts as an adhesion receptor in *Rickettsial* infections [35]. In these disorders, bacteria adhere to and invade vascular endothelial cells in a manner that must be strong enough to overcome the shear stress of flowing blood. In human umbilical vein endothelial cells, immunofluorescence–confocal microscopy revealed colocalization of AnxA2 and *Rickettsial* organisms on the external face of the plasma membrane. The numbers of *Rickettsia* adherent to wild-type mouse brain microvascular endothelial cells was significantly increased compared to those associated with cells from AnxA2-null mice. In-vivo findings corroborated the hypothesis that host endothelial cell AnxA2 serves as an adhesion receptor for *Rickettsial* species. In fact, using a plaque assay and with confocal imaging, He et al. showed an increase in *Rickettsia* in circulating blood and a concomitant decrease on the endothelial cell surface in AnxA2-null mice [35].

Finally, annexin A2 appears to be co-opted during the life cycle of at least 13 human viruses. Reported roles include attachment, penetration by receptor-mediated endocytosis or direct membrane fusion, replication, assembly, and release [39]. For example, the (A2•S100A10)2 tetramer was identified as a central mediator in human papillomavirus (HPV) attachment and intracellular trafficking, leading to progression of genital cancers. Suppression of AnxA2 altered the entry of HPV into target cells, whereas antibodies against S100A10 did not have the same effect [40]. In addition, the infection was significantly reduced when the (A2•S100A10)2 complex was eliminated, a result that was not seen when S100A10 alone was deleted [41,42].


**Table 1.** Role of annexin A2 in pathogen–host cell interactions in infection.

### **3. Annexin A2 and Regulation of Vascular Integrity**

One of the earliest responses to inflammation is a loss of vascular integrity, which facilitates the invasion of innate immune cells into the affected tissue. The maintenance of blood vessel integrity is an active process that, if interrupted, can cause hemorrhage, edema, and progressive inflammation. Vascular permeability is regulated by endothelial–endothelial cell junctions [43]. Homotypic interactions between vascular endothelial cadherin (VE-cad), a major constituent of the adherens-type junction, are regulated by Src-mediated tyrosine phosphorylation of VE-cad, which induces opening of the junction [44–46].

Annexin A2 interacts directly with VE-cad and is essential to maintaining VE-cad within cell junctions [45]. AnxA2 and S100 A10 interact independently with VE-cad via actin filaments, and vascular endothelial growth factor (VEGF) treatment disconnects AnxA2 and actin from the VE-cad complex, increasing vasculature permeability. Phosphorylation of the adherens junctions can be induced by either loss of Src homology phosphatase 2 (SHP2) or destabilization of cholesterol drafts, both of which implicate annexin A2 in vascular integrity [45]. Annexin A2 also regulates tyrosine phosphorylation of VE-cad in the pulmonary microvasculature. Under hypoxia, *Anxa2-*/*-* mice, unlike control *Anxa2*+/+ animals, showed a significant increase in Src-related phosphorylation of VE-cad. *Anxa2-*/*-* , but not *Anxa2*+/+, mice also displayed an acute inflammatory response with infiltration of neutrophils into the lung parenchyma and development of pulmonary edema. With VE-cad and SHP2, AnxA2 forms a complex, which is disrupted in the absence of AnxA2, thus preventing dephosphorylation of VE-Cad and inducing vascular leak [47].

In addition, in-vitro and in-vivo studies support a role for annexin A2 in maintaining the blood–brain barrier. Analysis of *Anxa2-*/*-* mice showed fewer tight junctions containing zonulin-1 and claudin-5 and fewer adherens junctions containing VE-cad. In cultured primary human brain microvascular endothelial cells, AnxA2 seems to promote F-actin and VE-cad interactions by binding Robo4 and contributing to its complex formation with paxillin. This process appears to be essential for maintaining vasculature integrity in the central nervous system [48].

### **4. Annexin A2 and Recruitment of Inflammatory Cells**

It is likely that annexin A2 is important for the recruitment of some classes of leukocytes to sites of inflammation. AnxA2 may interact with CD44 in the chemotaxis of neutrophil-like cells in response to complement factor 5a in vitro, and anti-AnxA2 appears to block this activity in human neutrophils [49]. At the same time, the level of expression of AnxA2 in freshly isolated human neutrophils appears to be low compared to other circulating leukocytes, especially human monocytes and monocyte-derived macrophages, where it is highly expressed [50]. In fact, anti-AnxA2 IgG impairs cytokine-directed monocyte migration through the extracellular matrix [50] This migratory activity also requires tPA-dependent activation of plasminogen to the serine protease plasmin. In addition, an in-vitro wound healing assay revealed that loss of AnxA2 expression in intestinal epithelial cells led to increased cell-matrix adhesion via a β integrin, and reduced cell migration [51], but whether this mechanism applies to inflammatory cells is unknown. Nevertheless, the full extent of the actions of AnxA2 in inflammatory cell recruitment in vivo remains to be determined.

### **5. Annexin A2 in Inflammasome Dynamics**

As an adaptive response, inflammation must be tightly regulated; inadequate inflammation can lead to persistent infection, whereas excessive or prolonged inflammation can cause longstanding disease, such as chronic arthritis, neurodegeneration, inflammatory bowel disease, or metabolic syndrome (atherosclerosis, type 2 diabetes and obesity) [52,53]. Inflammasomes are large intracellular protein complexes that sense inflammatory triggers in macrophages, monocytes, dendritic cells, neutrophils, and epithelial cells. They activate caspase-1 in response to both pathogens and host-derived signals of cellular stress including leakage of lysosomal cathepsins into the cytosol, damage to mitochondria, and production of reactive oxygen species [52].

As an adaptive response, inflammation must be tightly regulated; inadequate inflammation can lead to persistent infection, whereas excessive or prolonged inflammation can cause longstanding disease, such as chronic arthritis, neurodegeneration, inflammatory bowel disease, or metabolic syndrome (atherosclerosis, type 2 diabetes and obesity) [52,53]. Inflammasomes are large intracellular protein complexes that sense inflammatory triggers in macrophages, monocytes, dendritic cells, neutrophils, and epithelial cells. They activate caspase-1 in response to both pathogens and host-derived signals of cellular stress including leakage of lysosomal cathepsins into the cytosol, damage to mitochondria, and production of reactive oxygen species [52].

In humans with joint replacement devices, artificial articular surfaces may generate nonbiodegradable wear debris particles, which, upon phagocytosis by dendritic cells, damage the endolysosomal limiting membrane. Damage to membranes leads to discharge of lysosomal cathepsins and H<sup>+</sup> ions into the cytosol and activation of the NLRP3 inflammasome [54]. In the cytosol of dendritic cells exposed to wear debris particles, AnxA2 was profoundly downregulated, and both AnxA2 and S100A10 translocated to the endosome, to promote resealing of damaged membranes. These findings confirmed the hypothesis that AnxA2 is involved in endolysosomal membrane repair, thus limiting inflammasome activation [54].

On the other hand, upon infection with *Anaplasma phagocytophilum*, a *Rickettsial* organism, annexin AnxA2 null macrophages showed evidence of impaired inflammasome activation. Interleukin-1β secretion, caspase-1 activation, and NLRC4 oligomerization were all reduced compared to wild-type macrophages under the same conditions. Moreover, *Anxa2-*/*-* -infected mice were significantly more susceptible to infection than wild-type mice, indicating that AnxA2 helps this pathogen evade the NLRC4 inflammasome-based host defense system [55].

### **6. Annexin A2 and Macroautophagy**

Once the innate immune cells have identified the pathogens or damaged cells during the inflammatory response, the process of macroautophagy may be initiated. Macroautophagy is a cell survival mechanism, in which certain cytoplasmic constituents (damaged organelles, misfolded proteins, or bacteria) are degraded in a lysosome-derived, double-membrane-enclosed vacuole, and then recycled to maintain cellular homeostasis [56–58]. In inflammatory macroautophagy, the process begins when a pre-autophagosomal membrane elongates and forms an autophagosome by engulfing a portion of the cytosol. Autophagosomes initially fuse with endosomes to form amphisomes, and then ultimately fuse with lysosomes to degrade their contents [59]. Macroautophagy is associated with a number of human pathologies including cancer [60], myopathies [61], diabetes [62], neurodegenerative processes [63], and infectious disease [63,64].

Annexin A2 participates in macroautophagy by interacting with the autophagy-related protein Atg16, especially in the biogenesis of Atg16L-positive vesicles. In primary dendritic cells, which have the highest rate of plasma membrane turnover and are crucial for immunosurveillance, proteomic analyses identified a population of vesicles positive for both AnxA2 and Atg16L. Comparison of cells from *Anxa2*+/+ and *Anxa2-*/*-* mice demonstrated that AnxA2 facilitates Atg16L-positive vesicle fusion, a fundamental step in phagophore formation and elongation. Thus, AnxA2 appears to link Atg16L to membrane vesicles [15].

Macroautophagy is upregulated in response to a wide range of chronic inflammatory disorders and other stress conditions. These include persistent infections, such as tuberculosis, neurodegenerative disorders, such as Parkinson's disease, and chronic inflammatory disorders, such as Crohn's disease [65]. Under starvation conditions, for example, macroautophagy correlates with increased expression of annexin A2 in cultured cells and in mouse brain through a transcriptional pathway involving Jun N-terminal kinase (JNK) and c-Jun. Biogenesis of autophagosomes was enhanced in cells fed exogenous AnxA2 and abrogated in AnxA2-null mice. Autophagosome formation may be regulated by the effect of AnxA2 on actin-mediated trafficking of the transmembrane protein Atg9a, as AnxA2 and Atg9a were found to colocalize with actin filaments, leading to the conclusion that AnxA2 anchors actin on Atg9a-positive vesicles [66].

In another study, activation of autophagy during oxygen–glucose deprivation was reported to be influenced by AnxA2 in human retinal endothelial cells. Knockdown of AnxA2 attenuated the initiation of autophagy, reduced cell viability, and fostered cell apoptosis [67]. In another study, *Anxa2-*/*-* mice sustained a more severe *Pseudomonas aeruginosa* infection, which spread more readily to the lung and other organs. This appeared to correlate with an inability to clear the infection when annexin A2 is silenced and with the role of AnxA2 in regulating autophagosome formation through the Akt1–mTOR–ULK1/2 signaling pathway [68].

Macroautophagy is also central to the pathogenesis of osteoarthritis, a form of chronic joint inflammation. Autophagy may be protective in normal cartilage, as expression of ULK1, Beclin1 and LC3, the primary genes regulating the autophagy pathway, is decreased in osteoarthritic cartilage and chondrocytes [69]. In an in-vivo study with green fluorescent protein (GFP)-conjugated LC3 (GFP-LC3)-transgenic reporter mice, which allow one to monitor autophagy, there was a decrease in autophagic activity associated with a size and number reduction of autophagosomes [70]. These studies confirmed that age-related osteoarthritis correlates with autophagy, underlining a possible role for AnxA2 in this chronic inflammatory disorder.

### **7. Annexin A2 in Angiogenesis**

Angiogenesis is the process by which endothelial cells, derived from pre-existing vasculature, proliferate and migrate to create new blood vessels. It is distinct from vasculogenesis, in which

endothelial cells or their precursors coalesce to form new vasculature [71]. Under physiologic conditions, blood vessels undergo constant turnover, in which the rate of vessel degeneration is counterbalanced by the rate of blood vessel renewal. Insufficient angiogenesis is associated with myocardial infarction, stroke, preeclampsia, and neurodegeneration, whereas excessive vascular growth supports inflammation, obesity-associated disorders, development and spread of malignant tumors, and vascular-based ocular disorders such as diabetic retinopathy [72]. During inflammation, secreted pro-angiogenic factors promote neovascularization. The formation of new blood vessels, in turn, facilitates infiltration of innate immune cells, thus perpetuating the inflammatory process [73]. Additionally, upregulation of the inflammatory response caused by the persistence of the instigating agent may also facilitate angiogenesis through endothelial cell proliferation and migration [74].

Early in angiogenesis, pro-angiogenic signals activate endothelial cells, which migrate into the extracellular matrix, proliferate, elongate and create a lumen. The (A2•S100A10)2 complex, residing on the endothelial cell surface, converts plasminogen into plasmin, which can then activate a cascade of matrix metalloproteases. Matrix metalloproteases (MMP) can proteolyze components of the basement membrane, thus liberating endothelial cells and permitting their directed migration [72]. Based on these observations, it has been hypothesized that AnxA2 may be involved in excessive, pathological angiogenesis across a range of disease states [71].

Several in-vivo studies demonstrate a role for annexin A2 in postnatal angiogenesis. In the corneal pocket assay, for example, AnxA2-deficient mice displayed a decreased capability of generating new blood vessels in the cornea stimulated with fibroblast growth factor [75]. In wild-type mice, this response was blocked when mice were placed on a high-methionine diet; methionine is metabolized to homocysteine, which modifies AnxA2 by forming a covalent adduct with cysteine 8 in the N-terminal tail region. Interestingly, when hyperhomocysteinemic wild-type mice were treated with intravenous AnxA2, normal corneal angiogenesis ensued [76]. In the Matrigel implant assay, similarly, neovessel formation was significantly impaired in *Anxa2-*/*-* mice. Unlike *Anxa2-*/*-* mice, wild-type mice showed neovascularization of implanted Matrigel plugs. Addition of a peptide that mimics the N-terminal domain of annexin A2 and blocks tPA binding reduced the number of von Willebrand factor positive cells by 80%, whereas a scrambled control peptide had no effect [75].

In the oxygen-induced retinopathy (OIR) model of angiogenesis, 7-day-old neonatal mice are subjected to a 75% O2 environment for 5 days and then returned to room air (21% O2) for an additional 5-day recovery period. Immunohistologic analysis of retinas revealed a significant reduction of both neovascular tufts and tuft cell nuclei in *Anxa2-*/*-* mice. Under OIR conditions, the accumulation of fibrin in the perivasculature was extensive in *Anxa2-*/*-* retinas, but almost indiscernible in *Anxa2*+/+ retinas. Treatment of pups with ancrod, a fibrinogen-depleting agent, upon return to room air almost completely eliminated fibrin in both groups of mice, while at the same time doubling the extent of neovascularization in *Anxa2-*/*-* but not *Anxa2*+/+ mice. On the other hand, pups treated with tranexamic acid, an inhibitor of fibrinolysis, drastically increased fibrin deposition and lowered neovascularization in *Anxa2*+/+ but not *Anxa2-*/*-* retinas. In addition, it was established that hypoxia inducible factor-1, a master hypoxia-responsive transcription factor, directly regulates AnxA2 gene expression and that AnxA2 binding with S100A10 promotes neovascularization by enhancing plasmin activation and fibrin remodeling. Together, these data confirmed that both fibrinolysis and angiogenesis were impaired under ablation of annexin A2 [77]. Similarly, AnxA2 appears to be upregulated by vascular endothelial growth factor, and blockade of AnxA2 in ischemic mice inhibits retinal neovascularization [78]. In choroidal neovascularization, moreover, the AnxA2-binding agent, TM601, a synthetic form of chlorotoxin, significantly suppressed neovascularization by inducing apoptosis in endothelial cells [79]. Together, these studies indicate that the annexin A2 fibrinolytic system is a key regulator of angiogenesis.

### **8. Annexin A2 and Tumor Progression**

The correlation between inflammation, angiogenesis, and cancer is well known. In fact, hypoxiainduced release of VEGF, a feature of both cancer progression and tumor angiogenesis, increases cellular

proliferation, invasion and metastasis [74]. The involvement of annexin A2 in tumor progression may be related to the overproduction of plasmin on the surface of aggressive cancer cells, on the surface of endothelial cells, or both [80]. In addition, AnxA2 may facilitate the recruitment of pro-angiogenic inflammatory cells into the tumor microenvironment, which may in turn promote tumor progression.

Overexpression of annexin A2 has been observed in many malignancies, including aggressive breast cancer. AnxA2 was found to be elevated in stromal cells and epithelial cells of invasive ductal carcinoma (MDA-MB231) but not in a less aggressive cell line (MCF-7). MDA-MB231 cells, unlike MCF-7 cells, had the ability to activate plasminogen, leading to the hypothesis that plasmin generating capacity correlates with breast cancer cell migration and aggressiveness, implicating AnxA2 as a key mediator in metastasis. They also reported that blocking AnxA2 with angiostatin, a competitor for plasminogen binding on endothelial cell surface, or an anti-AnxA2 monoclonal antibody hindered MDA-MB231 invasion and migration. Immunohistochemical analyses of human breast cancer tissues revealed AnxA2 and elevated tPA on the surface of cancer cells, but not normal cells, as well as evidence of inflammation within the tumor. Quantitative analysis of the microvascular density showed a correlation between new blood vessel formation and the annexin A2 expression pattern [81–83]. Furthermore, immunoneutralization of AnxA2 impaired plasmin-mediated activation of MMP-2/9 in the breast tumor microenvironment [84]. Together, these data suggest that AnxA2 plays a pivotal role in breast cancer invasion and angiogenesis.

Annexin A2 expression is significantly elevated in glioblastoma, a highly vascularized and exceedingly aggressive brain tumor [85,86]. Intracerebral inoculation of canine glioma cells into the brains of athymic rats showed that AnxA2 is present in clusters of tumor cells surrounding dilated tumor vessels. Overexpression of AnxA2 correlated with high expression of VEGF and platelet-derived growth factor (PDGF), suggesting that the level of AnxA2 reflected invasiveness [87]. In addition, the involvement of AnxA2 in glioma angiogenesis was highlighted by the demonstration that tumors in wild-type mice displayed significantly higher microvascular density and more dilated blood vessels than tumors in *Anxa2-*/*-* mice [88].

Increased expression of AnxA2 appears to correlate with tumor invasiveness in renal cell carcinoma [89,90], hepatocellular carcinoma [91], colorectal cancer [92] and lung cancer [93]. These data suggest that AnxA2 may be a potential target in cancer therapeutics. In these latter studies, expression of AnxA2 has not yet been shown to correlate with activation of plasmin, as it has in breast cancer and glioblastoma. In colorectal cancer, upregulation of AnxA2 is regulated by TGF-β, which induces epithelial–mesenchymal transition [92]. In addition, silencing of AnxA2 in lung cancer stem cells in mice induced a reduction in tumor weight, which correlated with the loss of both β-catenin and S100A100, suggesting that AnxA2 may directly or indirectly regulate metastasis [93]. The mechanism for these effects, however, remains poorly understood.

### **9. Conclusions**

Annexin A2 appears to have a spectrum of functions across a multitude of inflammatory disorders. In early infection, according to our working model, AnxA2 may support infection by acting as an anchor protein that promotes adhesion and internalization of bacteria and viruses, regulating actin dynamics at adhesion sites, and enabling virus assembly. AnxA2 may also help recruit leukocytes to some sites of inflammation. AnxA2 also maintains the integrity of adherens junctions early in the inflammatory process by regulating phosphorylation of VE-cadherin. Numerous studies demonstrate that AnxA2 is required for optimal endosomal membrane stabilization and autophagosome biogenesis and promotes membrane repair during inflammation and infection. Cell-surface AnxA2 in complex with S100A10, finally, is a key factor in both malignant and non-malignant angiogenesis, and it contributes to tumor cell invasion and metastasis within an inflammatory microenvironment. We conclude, therefore, that annexin A2 has a primarily anti-inflammatory role, although it occasionally facilitates pathogen activity. Many of these actions have been elucidated through the use of the *Anxa2-*/*-* mouse,

which has become a useful tool for understanding the role of AnxA2 in vivo (Table 2). Taken together, these activities identify annexin A2 as a robust biomarker and potential therapeutic target.


**Table 2.** Studies in *Anxa2-*/*-* mice revealing a role of annexin A2 in inflammation.

**Author Contributions:** V.D. and K.A.H. conceptualized the manuscript. V.D. wrote the initial draft, and K.A.H. provided further editing. Both authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by the Belfer Diabetes Fund at Weill Cornell Medicine.

**Acknowledgments:** We thank Dena Almeida, Min Lucy Luo, Deyin Doreen Hsing, and Hana Lim, who provided valuable comments on the manuscript.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).
