The biology of a cell is the sum of many highly dynamic processes, each orchestrated by a plethora of proteins and other molecules. Microscopy is an invaluable approach to spatially and temporally dissect the molecular details of these processes. Hundreds of genetically encoded imaging tools have been developed that allow cell scientists to determine the function of a protein of interest in the context of these dynamic processes. Broadly, these tools fall into three strategies: observation, inhibition and activation. Using examples for each strategy, in this Cell Science at a Glance and the accompanying poster, we provide a guide to using these tools to dissect protein function in a given cellular process. Our focus here is on tools that allow rapid modification of proteins of interest and how observing the resulting changes in cell states is key to unlocking dynamic cell processes. The aim is to inspire the reader's next set of imaging experiments.

Cell biology involves understanding cellular processes at the molecular level. For a given pathway or process, we would like to know what molecules are involved and how they work together to generate a particular function. All cellular processes are highly dynamic, and microscopy is a powerful approach to investigate the molecular players in a particular pathway, reporting the spatial and temporal details that are not revealed by biochemical approaches. Cell scientists now have many wonderful genetically encoded imaging tools that can be deployed to unpick the functions of the molecules in a process, with many more methods being generated all the time. Here, we focus on tools that allow us to dynamically probe protein function in cellular processes that we have classified into three different strategies: observation – tools that allow us to observe protein function in as close to native state as possible; inhibition – tools that allow us to inhibit protein function and observe the effect on cellular processes; and activation – tools that allow us to activate proteins to confer function or reconstitute a process inside the cells.

With all three strategies, the approach used needs to be tuned to the timing of the cellular process under investigation. Any switches in cellular state must occur at least as rapidly as the process being studied (Box 1). Common approaches used are chemogenetic (using a chemical compound together with a compatible genetically encoded protein domain) or optogenetic (using light to trigger a change in a protein domain), often to induce heterodimerisation (Box 2). Generally, protein domains can be fused to one another in order to build a new protein with the desired characteristics to investigate your protein of interest (POI). Here, and in the accompanying poster, we provide a non-comprehensive overview of these tools, highlighting exemplar uses for each strategy and covering design principles to consider when designing your own experiments.
Box 1. The importance of timing

The tools and methods outlined in this article are described as rapidly acting. This is because each is sufficiently fast to allow imaging of a cell in the unperturbed state, before applying the respective method, switching state and observing the resulting behaviour. This possibility distinguishes them from slower-acting methods, such as RNAi or gene knockout, where a correlation of the observation with cellular states is not practical. Moreover, the timescale of many cellular processes is shorter than that of slower-acting methods (see poster section ‘Timing’); therefore, rapidly acting methods are the only feasible way to switch between the perturbed and unperturbed states.

A good example here is the study of cell division. The cell cycle might take 24 h, with M phase taking up to 1 h. If a slow-acting method such as RNAi is used, where the POI is degraded over 24-72 h, the function of the protein is difficult to discern as the cell goes through progressive mitoses with reduced levels of the protein. Nevertheless, with proteins that are rapidly turned over, this approach may still be useful. It is also possible that loss of the protein through RNAi has additional unintended consequences. A rapid inactivation method, where the protein is switched off immediately before (or during) mitosis, thus allows a more unambiguous interpretation of phenotype (Royle, 2013). Note that even the most rapid methods described here (optogenetics) can also work on longer timescales, so they should not be discounted as options when studying slower cell biological processes.

Box 2. Inducible heterodimerisation

Heterodimerisation of two genetically encoded proteins can be induced using chemical or optical means. In the widest used chemogenetic method, the FK506-binding protein (FKBP) domain of FKBP12 (also known as FKBP1A) is fused to one protein, and the FKBP–rapamycin-binding (FRB) domain of mTOR is fused to another. Rapamycin can then be applied, which binds to the FKBP domain, and the resulting FKBP–rapamycin complex binds tightly to the FRB domain (Kd, 10 nM), causing heterodimerisation of the two proteins (Spencer et al., 1993). Rapamycin derivatives (rapalogs), such as AP21967, do not interact with endogenous domains that bind rapamycin and can be used with modified FRB domains, for example FRB(T2098L), to reduce the potentially confounding effects of rapamycin on cell physiology. One example of a reversible chemically induced heterodimerisation system involves the coupling of a modified FKBP domain [FKBP(F36V)] to a non-FRB domain (eDHFR) via the synthetic dimerising agent SLF′–TMP (Voß et al., 2015). An alternative widely used chemical heterodimerisation method employs GA3-AM, a gibberellin (plant hormone) derivative that binds to the receptor GIBBERELLIN INSENSITIVE DWARF 1 (GID1). GA3-AM binding causes a conformational change in GID1 that permits it to bind to GIBBERELLIC ACID INSENSITIVE (GAI); this interaction has been exploited to induce dimerisation between proteins tagged with GID1 and GAI domains. There are further variations on this principle (see poster section ‘Main oligomerisation tools’).

A myriad of optogenetic heterodimerisation systems have also been developed (Benedetti, 2021) – including CRY2–CIB1 (Kennedy et al., 2010), tunable light-interacting proteins (TULIPs; Strickland et al., 2012), improved light-inducible dimer (iLID; Guntas et al., 2015), eMags (Benedetti et al., 2020) and others (see poster section ‘Main oligomerisation tools’) ­– which are documented at OptoBase (https://www.optobase.org/; Kolar et al., 2018). Generally, these methods include a light-sensitive domain, which binds another domain upon illumination with light of a certain wavelength. Both chemical and optical modalities give the cell scientist control over heterodimerisation in their experiment. They can even be combined, as in the case of the DHFR–zapalog system (Gutnick et al., 2019), to chemically induce heterodimerisation and then, by use of light, dissociate the induced dimer.

To understand the dynamics of proteins in a cellular process, a number of approaches allow us to image cells in an analogous way to biochemical ‘pulse chase’ experiments, where a labelled compound (pulse) is chased with an unlabelled compound to visualise the spatiotemporal behaviour of the labelled POI. Fluorescent protein (FP) tags that undergo photoactivation, photoswitching, photobleaching or photoconversion allow us to label subsets of tagged proteins and study their behaviour. Two popular methods to do this are fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP). Here, a POI is tagged with an FP and bleached using concentrated illumination in a subregion of the cell (see poster section ‘Protein dynamics’). The dynamics of recovery, or the loss of cellular signal, can be analysed to understand the mobility of the POI (Chudakov et al., 2010). An alternative is to selectively label the cell surface population of receptors and monitor their trafficking following endocytosis. This is possible with extracellular HaloTags and the use of cell-impermeable dyes (Huet-Calderwood et al., 2017). HaloTag is a 33 kDa tag derived from a bacterial enzyme that can be covalently labelled with a fluorescent dye. Labelling can also be achieved using a chemical heterodimerisation approach (Box 2) to ‘FerriTag’ POIs so that they can be observed by both light microscopy and electron microscopy (EM) (Clarke and Royle, 2018). Here, an engineered ferritin particle (visible by EM) with multiple copies of an FRB domain and the red FP mCherry (visible by fluorescence microscopy) can be induced to attach to a POI tagged with an FKBP domain in live cells.

Trafficking of proteins can also be monitored by retention using selective hooks (RUSH), a system where a POI can be released synchronously from a holding compartment in the cell (Boncompain et al., 2012). With RUSH (see poster section ‘Protein dynamics’), a secretory pathway POI is tagged with a streptavidin-binding peptide (SBP) and is co-expressed with an endoplasmic reticulum (ER)-resident protein that is fused to streptavidin. This ER-resident protein may have, for example, a KDEL ER-retention signal, and because SBP binds tightly to streptavidin, the POI is trapped in the ER. The SBP–streptavidin interaction can be broken by adding excess biotin, which freely diffuses across all cell compartments and thus can be easily added to the medium while the cells are on the microscope stage. Here, biotin competitively displaces the SBP as it binds streptavidin with a higher affinity. The SBP-tagged POI is then released, allowing us to observe its secretion and quantify its trafficking dynamics. For example, RUSH has been useful for examining the effect of physical forces on secretion (Phuyal et al., 2022) and for observing anterograde trafficking of procollagen (McCaughey et al., 2019).

Reporters

A number of FP-based sensors can respond dynamically to changes in cellular state. One example is pHluorins, which are pH-sensitive mutants of GFP whose fluorescence is quenched in acidic environments (such as lysosomes) (see poster section ‘Reporters’). pHluorins have been used as reporters of exocytosis and endocytosis (Miesenböck et al., 1998; Merrifield et al., 2005). Another example is the fluorescent ubiquitylation-based cell cycle indicator (FUCCI), which switches from a red fluorescence in G1 phase to a green signal in S-G2-M phase of the cell cycle via the tagging of two licensing factors (Cdt1 and geminin) that are differentially degraded (Sakaue-Sawano et al., 2008) (see poster section ‘Reporters’). Genetically encoded Ca2+ indicators, such as the GCaMP family, fluoresce when Ca2+ binds to the fused calmodulin (Chen et al., 2013). These are widely used as reporters of neuronal activity. These reporters, like others described in this article, are readily transplantable and can be targeted to different cellular compartments using tried-and-tested targeting motifs (see poster section ‘Reporters’).

Fluorescence resonance energy transfer (FRET)-based sensors have been used as an alternative method for imaging Ca2+ signals because they can report changes in protein proximity, which may be altered by binding of Ca2+ (Miyawaki et al., 1997) or of different second messenger molecules. This rationale has been exploited to engineer other FRET-based reporters for cellular states that can be reported by protein proximity, such as tension. If two labelled protein domains are separated by an elastic linker, then FRET can be used as a readout of tension. This approach has been used to visualise the spatiotemporal organisation of forces in live-cell imaging experiments (Grashoff et al., 2010).

Protein interactions

Apart from reporting on intraprotein dynamics, FRET is commonly used to understand changes in protein–protein interactions (PPIs). If the donor and acceptor fluorophores are fused to two distinct POIs, their proximity can be reported through donor quenching or acceptor emission. If two POIs are sufficiently close for FRET to occur, they can be inferred to bind (Chudakov et al., 2010).

Dynamic relocalisation of proteins can also be used to determine whether two distinct POIs interact (see poster section ‘Protein interaction’). The rationale is that if one POI is rerouted to a different cellular location, for example the mitochondria, and the second POI is co-rerouted, this is evidence for PPI (Cheeseman et al., 2013). However, other tests are required to understand whether co-rerouting represents a direct PPI or an indirect PPI via another protein, or even whether it is the result of the two proteins being present on the same cellular structure, which has itself been relocated with the POIs (Larocque et al., 2020). Methods for dynamic relocalisation using chemogenetics or optogenetics are covered below. One non-inducible method to relocalise proteins includes tagging a POI with a targeting motif to ensure ectopic localisation; for example, targeting Golgin proteins to the mitochondria (Wong and Munro, 2014). Another example of this is the use of nanobodies targeted to a particular location in order to relocalise a POI that is tagged with GFP (Rothbauer et al., 2008; Derivery et al., 2015; Küey et al., 2019).

A further method to visualise PPIs is bimolecular fluorescence complementation (BiFC; see poster section ‘Protein interaction’). This involves tagging two POIs with fragments of an FP that individually are not fluorescent. If the two proteins come together, it allows the FP fragments they are tagged with to assemble into a functional FP, and the interaction between the POIs can then be observed by microscopy (Hu and Kerppola, 2003).

Inferring the function of a protein within a pathway by looking at what happens to this pathway when the POI is inhibited is a cornerstone of cell biology. To switch to an inhibitory state, two main mechanisms can be deployed: the levels of the POI can be decreased or the POI can be physically removed from its normal site of activity.

Decreasing protein levels

One mechanism for decreasing proteins levels is by leveraging a system used in plant cells referred to as auxin-induced degradation. Here, the plant auxin hormone indole-3-acetic acid (IAA) is used to promote the interaction between the Skp-Cullin-F-box-containing (SCF) complex and an auxin-inducible degron (AID) tag, which is fused to the POI (Verma et al., 2020) (see poster section ‘Protein degradation’). The SCF complex recruits an E2 ubiquitin-conjugating enzyme, which results in polyubiquitylation of the AID tag and elimination of the AID-tagged POI by the proteasome. This method acts with a half-life of 20–40 mins (Nishimura et al., 2009), making it far faster than RNAi but slower than other methods discussed here. A related (non-inducible) method, termed deGradFP, uses a nanobody that recognises GFP to target a GFP-tagged POI to be degraded (Caussinus et al., 2011).

Proteolysis-targeting chimeras (PROTACs) that recruit ubiquitin ligases directly to POIs have also been used for targeted destruction and have proven to be a very useful tool for cell biologists (Bondeson et al., 2015; Wang et al., 2021). Here, the PROTAC is designed to include a so-called ‘warhead’ – usually a small molecule that is known to bind selectively to the target – that is used to couple the chimera to the POI (see poster section ‘Protein degradation’). As this method does not require tagging of the POI or expression of genetically encoded tools, PROTACs potentially can be deployed in native tissues. Although this is a strength of the approach, it also has the limitation that the method can only be used where appropriate ‘warheads’ can be developed, and so it is less adaptable compared with other approaches. An alternative is to use a hydrophobic tag strategy to label a protein, via HaloTags, so that it is targeted for degradation (Neklesa et al., 2011). A further method to inactivate proteins by degradation is Trim-Away (Clift et al., 2017), where an antibody that binds a POI is used to target it for proteasomal degradation. This method works around tripartite motif-containing protein 21 (TRIM21), a ubiquitin ligase and a receptor that recognises the introduced antibody. As it relies on an introduced antibody and endogenous factors, this method is particularly useful in cells where expression of genetically encoded tools is difficult. Finally, note that a number of promising approaches to use light for direct protein inactivation [such as chromophore-assisted light inactivation (CALI) and KillerRed] have been described, but none are widely utilised yet, suggesting that they might be difficult to use in practice (Ankenbruck et al., 2018).

One of the main chemical heterodimerisation tools – the FKBP–rapamycin–FRB system – has also been successfully modified so that it can be used to induce protein degradation. This application uses a modified FKBP domain fused to the POI and a molecule called dTAG. The dTAG molecule binds to both a modified FKBP domain, FKBP(F36V), and an E3 ubiquitin ligase complex component, cereblon. Thus, upon addition of dTAG to cells, the FKBP(F36V)-tagged POI is degraded by the proteasome (Nabet et al., 2018; Scheffler et al., 2022) (see poster section ‘Protein degradation’).

Protein relocation

Relocating a protein using induced heterodimerisation is a rapid and effective way to inactivate it. The rationale is that, since the POI is no longer at its normal site of action, it cannot function and is therefore inactivated. In knocksideways approaches, an FKBP domain-tagged POI is rerouted from its site of action to the mitochondria using an FRB domain attached to the mitochondrial outer membrane protein Tom70p. Upon addition of rapamycin, the POI is trapped at the mitochondria (Robinson et al., 2010) (see poster section ‘Rerouting and relocation’). Typically, the endogenous POI is depleted using RNAi so that its function is solely dependent on the overexpressed FKBP domain-tagged POI (Cheeseman et al., 2013). Alternatively, the FKBP domain can be knocked-in at the endogenous locus using gene editing (Ryan et al., 2021). Rerouting is extremely rapid, acting on a timescale of seconds to minutes, depending on the intrinsic dynamics of the POI (Robinson et al., 2010). It therefore outperforms degradation methods for the study of cell processes operating on this timescale. In addition to the FKBP–rapamycin–FRB system, there are several other chemogenetic tools for inducing heterodimerisation (see poster section ‘Main oligomerisation tools’), and recently this technology has been further developed to induce chemically the heterotrimerisation of three POIs by splitting an FRB domain (Wu et al., 2020). Protein inhibition via knocksideways can also be achieved using optogenetics. For example, the microtubule-binding protein regulator of cytokinesis 1 (PRC1), tagged with the bacterial protein SspB, can be co-expressed with iLID, which is targeted to the plasma membrane (PM) by a CAAX motif. Blue light can then be used to inactivate PRC1 by removing it from microtubules to the PM by inducing binding of the SspB tag on PRC1 to the PM-anchored iLID (Jagrić et al., 2021).

There are practical differences between optogenetic and chemogenetic heterodimerisation systems that affect their application. First, the illumination used in optogenetic systems can be directed to subcellular regions in order to give tight spatiotemporal control, whereas chemogenetic systems are typically activated throughout the entire cell and usually across all cells in the cell culture vessel. Second, most chemically induced heterodimerisation systems have long dissociation times (typically hours), which mean they can be considered irreversible on the timescale of most cell processes. By contrast, optogenetic methods are reversible (with a typical dissociation time of ∼10 s), which allows the cell scientist to observe recovery from inhibition as well as the effect of inhibition itself. It is possible to combine both approaches to allow photocaging of chemical heterodimerisation (Ballister et al., 2015) or to add reversibility of chemical heterodimerisation using light (Gutnick et al., 2019).

In order to relocate a POI and cause inhibition, the binding that underlies the normal localisation of a protein must be labile enough to allow removal (Robinson et al., 2010). Integral membrane proteins, or proteins that bind with very high affinity, cannot be removed in this way. However, if the compartment or structure upon which the POI is located is itself movable, then the entire compartment can be relocalised. This form of inhibition has been demonstrated for the ER (being cleared to the plasma membrane in mitotic cells) and for trafficking vesicles, which have been captured on mitochondria (Ferrandiz et al., 2022; Larocque et al., 2021; Hirst et al., 2015).

The same chemogenetic and optogenetic heterodimerisation tools can be deployed to induce activation of a cellular process, termed hotwiring (see poster). Two classic examples are the induced movement of organelles by coupling them to motors (van Bergeijk et al., 2015) and the recruitment of small GTPases to specific locations to induce their activity (Inoue et al., 2005; Komatsu et al., 2010). In addition, the formation of clathrin-coated pits can be induced at the plasma membrane to trigger endocytosis on demand (Wood et al., 2017), which is achieved by inducible recruitment of a clathrin-binding protein to a PM anchor. This event can be induced using chemogenetics (FKBP–rapamycin–FRB) or optogenetics (Strickland et al., 2012; Wood et al., 2017). By varying the anchor, it is possible to create clathrin-coated pits on other intracellular membranes, even those that do not normally support pit formation (Küey et al., 2022). Note that there are many target sequences that can be used to anchor a protein to the desired compartment (for widely used examples, see poster). In many cases, there are multiple anchors that can achieve the same function. For example, the transmembrane proteins CD8a and CD4, or peripheral membrane targeting sequences from Fyn or GAP43, can all be used for triggering endocytosis (Wood et al., 2017). This rationale of activation using heterodimerisation has been exploited in many other creative ways, such as to reactivate the spindle assembly checkpoint (Ballister et al., 2014), to artificially trigger T-cell immune responses (James and Vale, 2012) and to induce selective cell death (Shkarina et al., 2022) (see poster).

Rapamycin-activated protease through induced dimerisation (RAPID)-release is a novel approach to protein activation that has been developed to study the nuclear import of histones (Apta-Smith et al., 2018). Here, the POI is initially tethered to mitochondria via an OMP25 tag. The POI fusion construct also contains an FRB domain and a cleavage domain that is cut by the tobacco vein mottling virus (TVMV) protease. An FKBP domain-tagged TVMV protease is co-expressed in this system. Rapamycin addition recruits the protease to the POI through formation of a protease–FKBP–rapamycin–FRB–POI complex. There, the protease cuts the cleavage domain and liberates the POI from the mitochondrially anchored remainder of the fusion construct. This allows the nuclear import of the released POI to be visualised (with a half-time of 5 mins; Apta-Smith et al., 2018) (see poster section ‘Hotwiring’). RAPID-release is similar to RUSH in allowing the on-demand release of a POI. However, because the initial location of the POI is ectopic rather than within the pathway of interest, we classify this method as an activation strategy rather than an observation strategy.

If it were possible to rapidly produce a POI in cells that do not express it, this could, in principle, be used to activate a pathway that the POI acts in. In practice, inducible expression is too slow (minutes to hours) to be useful for most dynamic cell processes (seconds to minutes). A useful alternative is to continually degrade the POI, before protecting it from degradation in an inducible manner. This can be achieved by tagging a POI with a modified FKBP domain, which results in the constitutive degradation of the tagged protein, before adding a compound (Shield-1) that binds to the modified FKBP domain and shields the tagged protein from degradation (Banaszynski et al., 2006) (see poster section ‘Protein stabilisation’).

Most cellular processes are in a dynamic equilibrium; therefore, upregulation of the activity of one protein can be used to inhibit the production of other proteins that it interacts with, resulting in the inhibition of downstream pathways. Two examples that defy our categorisation of inhibition and activation are the severing of microtubules using opto-katanin (Meiring et al., 2022) and the inhibition of endocytosis by phosphatidylinositol (4,5)-bisphosphate [PI(4,5)P2] depletion (Zoncu et al., 2007). In the first case, the microtubule-severing enzyme katanin is recruited to microtubules using the iLID optogenetic system, causing the localised disassembly of microtubules, which leads to inhibition of transport and other processes (Meiring et al., 2022). In the second example, the activation of inositol 5-phosphatase to reduce PI(4,5)P2 levels at the PM, using chemogenetics, leads to an inhibition of processes that depend on this phosphoinositide, such as endocytosis (Zoncu et al., 2007).

Designing and validating a construct

Almost all of the tools described here require the fusion of protein domains into a single construct for expression in cells. This modular approach is possible because proteins consist of one or more functional domains, which can be isolated and fused to other domains via short linker sequences, generating new constructs. The design and validation of each new construct is crucial to a successful experiment (see poster section ‘Design and validation of a construct’). In the simplest case, fusion of an FP to a POI, one must consider whether the FP is fused to the N terminus or C terminus of the POI. If both N- and C-termini of the POI are known to be important for the function of the protein, or if their orientation means that the tagging would not be functional (e.g. both C- and N-termini of a POI are within the lumen of an organelle, but you want to study a process that takes place in the cytosol), then internal insertion of an FP is a possible alternative. Previously, fusions were generated by trial and error, but now the structure prediction software AlphaFold can guide the design of fusions (Jumper et al., 2021) by identifying domain boundaries and loops where tags may be inserted. In addition, the length and nature of linker sequences is important. Generally, flexible linkers are favoured to allow for protein domains to be independent of one another, but they should only comprise inert amino acid sequences and not contain, for example, sequences for import into cell organelle membranes, which might mistarget the fusion protein inappropriately. Note that long linkers may self-cleave and that glycine-rich linkers are prone to aggregation (Gräwe and Stein, 2021).

Other issues to consider when designing a construct are the properties of the domains to be fused. What is the size of the domain relative to the size of the POI? For FPs, FPbase is a valuable resource that provides information about their properties (such as fluorescence spectra, multimericity and stability) (https://www.fpbase.org/; Lambert, 2019; Cranfill et al., 2016). How the construct will be expressed is a further variable. For some constructs, transient expression with a strong promoter [such as the cytomegalovirus (CMV) promoter] is fine, but for others the overexpression may cause problems and a lower-expressing promoter [such as the phosphoglycerate kinase 1 (PGK) promoter] may be necessary. Expression at endogenous levels by using gene editing to knock-in additional domains is likely to be the optimal solution (Ryan et al., 2021), but this requires substantially more work. Importantly, each construct must be carefully validated. The simple addition of even a small tag is likely to disrupt the function of a protein to some extent. It is therefore crucial to assess this impact before attempting to use the construct in experiments to test hypotheses about the functions of tagged POIs (see poster section ‘Design and validation of a construct’); for example, whether the localisation or dynamics of a POI are affected should be checked in the first instance. Further functional validation can be achieved by testing whether the expression construct can fully complement function in cells depleted of the endogenous POI, or whether the expressed protein still interacts with known binding partners.

In this Cell Science at a Glance and the accompanying poster, we have highlighted ways that genetically encoded tools can help us to observe cell biological processes and how they can be used to inhibit or activate POIs, allowing us to dissect their roles within these processes. We have reiterated the importance of timing when dealing with dynamic processes inside the cell, and that manipulating protein function must be done on a timescale that is at least as fast as the process under study. The past 20 years have seen an expansion of genetically encoded imaging tools, with new variants constantly being reported. Engineering and ‘remixing’ of domains is highly creative and has generated a diverse range of tools applicable to multiple aspects of cell biology. The possibilities are vast, and novel combinations continue to uncover new insights into cellular functions. It is an exciting time to be a cell biologist.

We thank all members of the lab for critical discussion.

Funding

Work in our lab is supported by Cancer Research UK (C25425/A27718), UK Research and Innovation - Biotechnology and Biological Sciences Research Council (BB/V003062/1), UK Research and Innovation - Medical Research Council (MR/N014294/1) and Human Frontier Science Program (RGP25/2022).

High-resolution poster and poster panels

A high-resolution version of the poster and individual poster panels are available for downloading at https://journals.biologists.com/jcs/article-lookup/doi/10.1242/jcs.260783#supplementary-data.

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Competing interests

S.J.R. is a director of The Company of Biologists but was not included in any aspect of the editorial handling of this article or peer review process. The authors declare no financial interests.