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. Author manuscript; available in PMC: 2026 Apr 28.
Published in final edited form as: J Cell Sci. 2025 Oct 24;138(20):jcs264137. doi: 10.1242/jcs.264137

DynAPs and cytoplasmic assembly of axonemal dyneins

John B Wallingford 1, Steven L Brody 2, Amjad Horani 3, Chanjae Lee 1
PMCID: PMC13112607  NIHMSID: NIHMS2166618  PMID: 41134057

Abstract

Motile cilia are microtubule-based organelles that generate fluid flow through coordinated beating, a process powered by axonemal dynein motors. Dyneins are pre-assembled in the cytoplasm by a suite of proteins called dynein axonemal assembly factors (DNAAFs). Genetic variants affecting either the motors or the assembly factors cause motile ciliopathy. In recent years, DNAAFs have been found to function in conjunction with heat shock protein (HSP) chaperone systems and organize with dynein subunits within cytoplasmic foci known as “dynein axonemal particles,” or DynAPs. In this Perspective, we provide our view on the assembly and potential function of DynAPs as well as their place within the broader context of motile ciliated cells.

Summary statement:

This Perspective discusses current work in the field of dynein axonemal particles (DynAPs), organelles found in motile ciliated cells that concentrate axonemal dynein subunits and chaperones required for their cytoplasmic preassembly before deployment to axonemes.

Introduction

Motile cilia are microtubule-based cellular projections that beat in a coordinated manner to generate fluid flow crucial for development and homeostasis. The beating of motile cilia is driven by an assemblage of axoneme-specific dynein motors acting collectively to slide axonemal microtubule doublets, the core structures of cilia. These multi-protein motors, called dynein arms owing to their extension from the axoneme, are subdivided into two broad categories of outer and inner dynein arms (ODAs and IDAs, respectively) based on their position on the axonemes. Dynein arms are massive heterodimeric or monomeric complexes, each composed of distinct combinations of heavy, intermediate and light dynein chain proteins (King, 2018). ODAs drive ciliary beating generally, whereas IDAs tune the shape of the cilia as they beat (Fig. 1B) (Kamiya and Yagi, 2014; King, 2018). ODAs and IDAs are further subdivided into classes, based largely on their proximodistal position along the axoneme (Dutcher, 2020; Grossman-Haham, 2023).

Fig. 1. DynAPs and multiciliated cells.

Fig. 1.

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A) Cross-sectional view of Xenopus multiciliated cells (MCCs) labeled with the outer dynein arm (ODA) subunit Dnai2, revealing the protein’s localization in apical axonemes and in dynein axonemal particles (DynAPs). Image by Chanjae Lee. B) Schematic of a MCC shows the position of the axonemes and DynAPs, as well as the positions of ODAs and inner dynein arms (IDAs) in the axoneme. The inset fluorescence microscopy images at bottom show colocalization of a DNAAF (Dnaaf5) and a chaperone (Hsp90ab1) throughout DynAPs and the partitioning of an ODA subunit (Dnai2) and IDA subunit (Wdr78) into sub-domains within DynAPs (sub-DynAPs). Images by Chanjae Lee .C) The DNAAF Lrrc6/Dnaaf11 labels DynAPs in a human airway MCC. Image reproduced from (Huizar et al., 2018).

Pathologic variants in genes encoding ODA subunits are a major cause of the motile ciliopathy syndrome known as primary ciliary dyskinesia (PCD; OMIM entry 244400, https://www.omim.org/entry/244400). This genetic disease, which is generally autosomal recessive, results in a syndrome of chronic sinopulmonary disease, bronchiectasis, situs anomalies, cardiac defects and infertility (Horani et al., 2016; Mitchison and Valente, 2017; Wallmeier et al., 2020). Because axonemal dynein motors are pre-assembled in the cytoplasm before deployment to cilia (Fowkes and Mitchell, 1998), PCD can also result from pathogenic variants in genes that encode any of several cytoplasmic proteins known collectively as dynein axonemal assembly factors (DNAAFs) (Box 1) (Desai et al., 2018; Fabczak and Osinka, 2019; Qiu and Roy, 2022). These DNAAF proteins are now identified by a unified, consensus nomenclature (Braschi et al., 2022).

Box 1:

Nomenclature Explainer

DNAAF: This term stands for “dynein axonemal assembly factor” (Mitchison et al., 2012), and can be confusing to the uninitiate because these factors act not in the axoneme but in the cytoplasm. The confusing phrasing that reflects the agreed-upon but awkward nomenclature for axonemal dynein subunits, in which each gene is named: “Dynein, axonemal (weight)(number).” For example, Dnah5 encodes dynein axonemal heavy chain five, and Dnali1 encodes dynein axonemal light-intermediate chain 1.

DynAP: This term stands for “dynein axonemal particle”, reflecting the nomenclature for DNAAFs. These are MCC-specific cytosolic foci described in Xenopus and humans that contain numerous DNAAFs (such as Ktu, Heatr2 and Lrrc6) and chaperones (such as Ruvbl1 and Ruvbl2) (Huizar et al., 2018).

kl-granule: These granules are named for the Drosophila genes kl-3 and kl-5, which encode ODA subunits (Fingerhut and Yamashita, 2020). These gene names reflect the complex nomenclature of Y-linked fertility-related genes in the fly (Brosseau, 1960).

R2HAD: This term stands for “Ruvbl1/Ruvbl2 hubs for axonemal dyneins”. This name has been given to foci in zebrafish that contain axonemal dynein heavy chain mRNAs and protein as well as Ruvbl1, Ruvbl2 and Lrrc6 (Li et al., 2024). It is our view that these foci should be called DynAPs.

The first DNAAF to be identified was kintoun (KTU, now called DNAAF2) (Omran et al., 2008), and human genetic and cell biological studies have now defined 19 DNAAFs (Desai et al., 2018; Fabczak and Osinka, 2019; Qiu and Roy, 2022). Of the 19 known DNAAFs, variants in 14 of the genes are causative of PCD (Table 1). Generally, biallelic disruption of any single DNAAF has been found to result in an absence of both ODAs and IDAs throughout the ciliary axoneme, but some studies suggests that at least some distinct DNAAFs may be dedicated for the assembly of specific subsets of dyneins (Lee et al., 2020; Yamaguchi et al., 2018).

Table 1:

Disease-related DNAAFs

DNAAF Synonym(s) Human Ciliopathy Reference
DNAAF1 LRRC50 (Duquesnoy et al., 2009; Loges et al., 2009)
DNAAF2 KTU (Omran et al., 2008)
DNAAF3 PF22, C19orf51 (Mitchison et al., 2012)
DNAAF4 DYX1C1 (Tarkar et al., 2013)
DNAAF5 HEATR2 (Horani et al., 2012)
DNAAF6 PIH1D3 (Olcese et al., 2017)
DNAAF7 ZMYND10 (Zariwala et al., 2013)
DNAAF11 LRRC6 (Horani et al., 2013; Kott et al., 2013)
DNAAF12 Officially LRRC56 (Bonnefoy et al., 2018)
DNAAF13 Officially SPAG1 (Knowles et al., 2013)
DNAAF16 Officially CFAP298 (Austin-Tse et al., 2013)
DNAAF17 Officially CFAP300, also C11ORF70 (Fassad et al., 2018; Höben et al., 2018)
DNAAF18 Officially DAW1, also WDR69 (Leslie et al., 2022)
DNAAF19 CCDC103 (Panizzi et al., 2012)
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The following DNAAFs have yet to be directly implicated in PCD or infertility in humans: DNAAF8 (Daap1, C16ORF7), DNAAF9 (C20ORF194, Shulin), DNAAF10 (WDR92, Monad), DNAAF14 (PIH1D1), DNAAF15 (PIH1D2)

Proteomic experiments initially revealed what is now a well-established link between DNAAFs and heat shock family chaperones (HSPs) (Fabczak and Osinka, 2019; Omran et al., 2008). Indeed, genetic ablation of the Hsp90 co-chaperones Ruvbl1 and Ruvbl2, scaffolding proteins that organize multimolecular complexes, results in defective dynein arm assembly in zebrafish and mice (Li et al., 2017; Zhao et al., 2013). It is now thought that DNAAFs function as part of a “chaperone relay” (Mali et al., 2018), in which distinct complexes of chaperones and DNAAFs function independently on different regions of the dynein polypeptide during protein folding. This process may or may not involve a sequential hand-off – or relay – of chaperones between DNAAFs (Cho et al., 2018; Olcese et al., 2017; Yamaguchi et al., 2018; Zur Lage et al., 2018).

In this Perspective, we review the interplay of DNAAFs, chaperones and dynein subunits and their colocalization into cytosolic foci we have termed “dynein axonemal particles”, or DynAPs (Fig. 1A, Box 1) (Huizar et al., 2018). We discuss the assembly and potential function of DynAPs, as well as similar membraneless organelles found in motile ciliated cells and other nomenclature that has recently been applied to these organelles.

Puncta, foci and DynAPs

Though it was not recognized as such at the time, the earliest evidence of discrete foci related to axonemal dynein assembly in ciliated cells came from work in zebrafish in 2008 (Kishimoto et al., 2008). Via immunostaining, Lrrc6 (now called Dnaaf11) was found to be present in cytosolic foci specifically in ciliated tissues in zebrafish. The cystic kidneys of lrrc6 mutant fish observed in the study were suggestive of a ciliary role for Lrrc6, but abnormal cilia function was not apparent in assays of cilia motility or axonemal structure (Kishimoto et al., 2008). Later, in a study using different lrrc6 mutant alleles, Lrrc6 was shown to be essential for cilia beating (Serluca et al., 2009), but the link to dynein assembly remained obscure because structural axoneme defects were not observed (Serluca et al., 2009). It seems likely that the subtlety of the phenotype or the incomplete penetrance of the alleles obscured the nature of the cell biological defect in these fish. Regardless, subsequent transmission electron microscopy (TEM) analysis of human patients with variants in LRRC6 finally linked the protein to axonemal dyneins (Fig. 1C) (Horani et al., 2013; Kott et al., 2012).

In 2012, the first definitive link between cytosolic foci and axonemal dynein assembly was made when HEATR2 (also called DNAAF5) was found in human cells to be essential for assembly of dynein arms and thus for cilia beating (Horani et al., 2012). HEATR2/DNAAF5 protein was also found in foci in the cytosol of multiciliated cells (MCCs). Importantly, these foci were shown to be distinct from other known organelles and did not colocalize with markers of lysosomes, endosomes or centrosomes (Horani et al., 2012).

The first link between these foci and more broadly acting chaperones arrived the following year, when Lrrc6 was found to colocalize in puncta with the Ruvbl2 ortholog, reptin, in zebrafish (Zhao et al., 2013). Ruvbl2 plays a broad role in protein folding (Dauden et al., 2021), yet its loss in mutant fish evoked a specific defect in axonemal dynein assembly (Zhao et al., 2013). Ruvbl2’s binding partner Ruvbl1 was later also found to colocalize in these cytosolic foci and to be essential for cilia beating in mice (Li et al., 2017).

In 2018, two papers revealed that colocalization of DNAAFs, chaperones and dynein subunits in cytosolic foci is a general feature of MCCs. First, we reported that in human airway cells, HEATR2/DNAAF5-containing foci also contain KTU/DNAAF2 and SPAG1 (also called DNAAF13), as well as the axonemal dynein subunit DNAI1 (Horani et al., 2018). We also found that DNAI1 colocalized in foci with LRRC6 (Huizar et al., 2018).

To better resolve these colocalizations, we turned to the very large MCCs found in the epidermis of Xenopus laevis embryos (Walentek and Quigley, 2017). In a study of nine distinct DNAAFs, six axonemal dynein subunits and seven broadly-acting chaperones, we found that all robustly colocalized in cytosolic foci (Fig. 1A) (Huizar et al., 2018). We further showed that foci labeled by these proteins did not colocalize with known markers of membrane-bound organelles or with other known foci-forming proteins (Huizar et al., 2018). We thus coined the term “dynein axonemal particles” (DynAPs) for these foci (Box 1).

It is important to note that although this work on Xenopus DynAPs involved low-level overexpression of DynAP-associated mRNAs, several lines of evidence demonstrate the findings’ veracity. First, DynAPs are readily observed in unmanipulated Xenopus MCCs using immunostaining for endogenous Ruvbl2 (Huizar et al., 2018), just as they are in fish and mammals (Li et al., 2017; Zhao et al., 2013). Second, in a screen of over 200 different proteins overexpressed in Xenopus MCCs, less than ten percent localized to cytosolic foci, and of these, only a single protein localized to DynAPs (the Hsp-related chaperone Stip1) (Tu et al., 2018). Third, overexpression of DynAP proteins consistently labels foci in MCCs, but it does not do so in the immediately adjacent mucus-secreting cells that receive precisely the same dose of mRNA. However, conversion of those neighboring cells into ciliated cells by mis-expression of ciliary transcription factors endows them with the ability to form cytosolic foci when DynAP proteins are also expressed (Huizar et al., 2018). Thus, DynAP formation is not related to mRNA overexpression generally but is a specific event occurring only with DynAP proteins expressed in the milieu of motile ciliated cells. Finally, it is essential to note that DynAPs are demonstrably distinct from other RNA-related organelles in Xenopus, as they are not labelled by markers of P-bodies, stress granules or other known RNA-containing foci (Huizar et al., 2018).

In a subsequent study, we found that Xenopus DynAPs are subdivided internally, with ODA and IDA dynein subunits occupying distinct regions within them. By contrast, DNAAFs and chaperones are present throughout DynAPs and are not partitioned into visible sub-regions (Lee et al., 2020). We refer to these sub-regions as “ODA sub-DynAPs” and “IDA sub-DynAPs” (Fig. 1B). Remarkably, even distinct classes of IDA partition into distinct compartments, with sub-DynAPs containing subunits of IDA-f being distinguishable from those containing IDA-a,c,d subunits (Lee et al., 2020).

In sum, it is our view that DynAPs should be defined as any membraneless organelle present in motile ciliated cells where dynein subunits, DNAAFs and broadly acting chaperones colocalize. Moreover, DynAPs are further partitioned into sub-DynAPs, in which the subunits of specific classes of ODA and IDAs are concentrated. Whether further sub-domains exist within sub-DynAPs is an important open question.

DynAPs are membraneless organelles that display some hallmarks of phase separation

In our original descriptions in human and Xenopus cells, we noted that DynAPs appear not to be membrane-bound and that they display “hallmarks of phase separation”. This conclusion was based on three findings. First, the foci were not labelled by any of several markers of known membrane-bound organelles (Horani et al., 2018; Huizar et al., 2018). Second, movies of DynAPs revealed they have relatively fluid shapes, which divide and re-fuse quickly (Huizar et al., 2018). Third, the rapid recovery of DNAAFs and chaperones within DynAPs in fluorescence recovery after photobleaching (FRAP) experiments was similar to that reported for other liquid-like organelles (Huizar et al., 2018). This led us to suggest that DynAPs may form as liquid-like biomolecular condensates (Banani et al., 2017).

However, FRAP experiments also revealed that dynein subunits were very stably retained inside DynAPs, suggesting the presence of a more stable, non-liquid component (Huizar et al., 2018). In fact, none of the dozens of constituent proteins in DynAPs have been shown to phase separate in vitro and it is important to note that in vivo analyses like FRAP provide only suggestions of the actual physical properties of organelles (McSwiggen et al., 2019). In an important commentary, Stephen King noted this issue and proposed an alternative interpretation of DynAP assembly: based on the massive size of dynein heavy chain proteins, he proposed that mRNAs and polysomes might simply be cross-linked by the assembly of the nascent multiprotein motors themselves, thus forming distinct foci (King, 2021).

Recent evidence discussed in more detail below suggests that DynAPs comprise a combination of liquid and solid-like features. Indeed, this mixture of behaviors for different constituent proteins in DynAPs is reminiscent of the known organization of stress granules, another type of cytosolic membraneless organelle (Jain et al., 2016; Wheeler et al., 2016).

In our view, the concept of DynAPs, as we originally defined it, is useful regardless of the underlying physical basis for these organelles’ assembly: DynAPs are sites of colocalization and high concentration of axonemal dynein subunits, DNAAFs and diverse chaperones.

DynAPs, kl-particles and R2HADs

RNA is a key component of many liquid-like organelles and the first suggestion that DynAPs contain RNA came from the observation that they are enriched for the RNA-binding protein G3bp1 (Huizar et al., 2018). Definitive evidence was obtained two years later, when a screen revealed that novel RNA-associated proteins localized to DynAPs in Xenopus, and staining with a general RNA dye revealed strong concentration of RNA in these organelles (Drew et al., 2020).

Even better evidence then came from a curious corner. A study of flagellar assembly in Drosophila sperm revealed that mRNAs encoding heavy chains of axonemal dyneins were present in discrete foci (Fingerhut et al., 2019). Those foci were designated kl-granules, reflecting the nomenclature for the Drosophila axonemal dyneins involved (Box 1). Fascinatingly, kl-granules resemble DynAPs by being strongly enriched in Ruvbl1 and Ruvbl2. Moreover, kl-granules partition mRNAs encoding ODA and IDA heavy chains into distinct sub-regions (Fingerhut and Yamashita, 2020), reflecting the localization of IDA and ODA proteins observed in sub-DynAPs (Lee et al., 2020). Lastly, loss of Ruvbl1 or Ruvbl2 disrupts the focal localization of the dynein heavy chain mRNAs, as well as their translation and the assembly of mature ODAs and IDAs (Fingerhut and Yamashita, 2020). Because motile flagella in Drosophila sperm form through a highly specialized mechanism, it is not entirely clear how kl-granules relate to DynAPs, but we believe their similarities are important and informative.

More directly relevant to MCCs are similar findings obtained recently in zebrafish, where the mRNAs for two ODA heavy chains were found to colocalize in foci that also contained the encoded ODA proteins (Li et al., 2024). This result suggested the foci are sites of active translation of ODA subunits. Indeed, disruption of translation of these ODA mRNAs led to dissolution of the foci. As in Xenopus (Huizar et al., 2018), these foci of ODA protein were found to be the same foci that contain Ruvbl2 and Lrrc6. The localization of Ruvbl2 and Lrrc6 to foci was also dependent upon translation (Li et al., 2024). Interestingly, mutation of Ruvbl1 disrupted the localization of Lrrc6 to foci, but the converse was not true; moreover, the DNAAF Pih1d3 acted farther downstream of Ruvbl1 and Lrrc6 (Li et al., 2024). These findings are consistent with the concept of DNAAFs acting in a “chaperone relay” (Mali et al., 2018).

Because they dissolve when translation is blocked, the foci characterized in zebrafish MCCs fit many of the parameters of the “dynein factories” model proposed for DynAPs by King (King, 2021). However, in one fascinating exception, they also reflect the phase separation model: hexanediol, which disrupts the weak multivalent interactions in phase-separated organelles, dispersed Ruvbl1, Ruvbl2 and Lrrc6 from foci but did not disperse dynein axonemal heavy chain 9 (Dnah9) (Li et al., 2024). This result perfectly reflects earlier FRAP observations showing that dynein subunits are highly stable within DynAPs, whereas chaperones and DNAAFs (including Ruvbl2 and Lrrc6) turn over rapidly (Huizar et al., 2018).

Li et al. termed the foci they observed in zebrafish MCCs “R2HADs” (Box 1) (Li et al., 2024), but because those authors chose not to mention DynAPs, it is difficult to know how they believe the two entities to be related. More work is warranted, but it seems likely that R2HADs are just DynAPs.

DynAPs and R2TP

New insights into how DynAPs might assemble and function can be gleaned from studies of the Hsp90-R2TP complex. This complex aids in assembly of diverse macromolecular machines, including RNA polymerases and small nucleolar ribonucleoproteins (snoRNPs), and is comprised of Ruvbl1, Ruvbl2, Hsp90 and the effectors Pihd1 and Rpap3 (Lynham and Houry, 2022). This complex is of interest in DynAP assembly because of the identification of a related R2TP-like complex, sometimes called R2SP, in which Pih1d1 and Rpap3 are replaced with Pih1d2 and Spag1 (Maurizy et al., 2018), two proteins now known to be DNAAFs (DNAAF15 and DNAAF13, respectively) (Knowles et al., 2013; Smith et al., 2022; Yamaguchi et al., 2018).

Like Pihd1, Ktu/Dnaaf2 also contains proteins interacting with HSP90 (PIH) domains (Omran et al., 2008) and Pih1d3 is known to be crucial for ODA and IDA assembly (Olcese et al., 2017; Yamaguchi et al., 2018). Indeed, a systematic survey of PIH proteins in zebrafish strongly suggests that R2TP, R2SPand other potential complexes combining Ruvbl1, Ruvbl2 and PIH domain proteins each play specific roles in assembling subsets of ODA and IDAs (Yamaguchi et al., 2018). Similarly complex patterns of loss of ODA or IDAs were also observed after knockdown of Daap1/Dnaaf8 in Xenopus (Lee et al., 2020).

In a recent preprint, R2TP was found to assemble co-translationally (Philippe et al., 2025), an important finding in light of the demonstration of co-translational assembly in DynAPs (Li et al., 2024). Moreover, the preprint showed that Ruvbl2 co-immunoprecipitated with the mRNAs of R2TP client proteins and that artificially driving Ruvbl2 into liquid-like foci also drove recruitment of client mRNAs into those foci (Philippe et al., 2025). This suggests that foci fluidity may be an important parameter for Ruvbl2 action in R2TP complexes, an idea that is also potentially relevant for DynAPs, as knockdown of Heatr2/Dnaaf5 in Xenopus was shown to alter the flux of other DNAAFs within DynAPs, affect DynAP size and disrupt ODA deployment (Huizar et al., 2018).

What are DynAPs and what do they do?

Although it may be too soon to say with certainty, R2HADs and kl-granules likely represent some flavor of DynAP. If this is the case, it seems probable that the more solid cores of these structures, containing mRNAs and nascent proteins of axonemal dynein subunits, are formed by co-translational crosslinking, as proposed by King and demonstrated by FRAP assays and experiments that manipulated translation (Huizar et al., 2018; King, 2021; Li et al., 2024). It also seems likely that DNAAFs and chaperones condense around these cores in a more fluid-like manner, as suggested by FRAP assays and by their dispersal in response to hexanediol treatment (Huizar et al., 2018; Li et al., 2024).

Interestingly, this view of DynAPs reflects the known constitution of stress granules. Both proteomic and imaging studies suggest that stress granules are composed of stable cores and more fluid-like shells (Jain et al., 2016; Wheeler et al., 2016). Stress granules, like DynAPs, are also enriched for Ruvbl1, Ruvbl2, HSP chaperones and G3bp1 (Jain et al., 2016). However, DynAPs do not contain all the proteins found in stress granules (e.g., Tia1). As with stress granules (Glauninger et al., 2022), if and how this “mixed” nature of DynAPs contributes to their function remains an open question.

One possibility is that DynAPs function by increasing the local concentration of specific proteins, thereby locally increasing the efficiency of biochemical reactions. For example, in liquid-like Cajal bodies, Ruvbl1 and Ruvbl2 are required for processing spliceosomal ribonucleoproteins (RNPs) (Cloutier et al., 2017). Modeling of Cajal bodies suggests that increasing local client concentrations can accelerate this process by 10-fold (Klingauf et al., 2006). Similar functions have been attributed to the distinct phase-separated domains within nucleoli (Feric et al., 2016). Precisely which reactions might be accelerated inside DynAPs is unknown, but could include translation itself, interactions between subunits during motor assembly, or transient interactions of nascent dyneins with chaperones. Directly testing these ideas in DynAPs has been difficult, albeit no less difficult than this has been in other membraneless organelles. Accordingly, it is also possible that DynAPs ‘do’ nothing at all – that their formation is no more than mere epiphenomena arising from the process of dynein translation and assembly.

Alternatively, DynAPs might perform some as-yet undefined function. Here, the known role of Ruvbl1, Ruvbl2 in processing spliceosomal RNPs is intriguing, because the spliceosome protein Srsf1 was recently found to act outside the nucleus to control translation of certain ODA and IDA subunits (Haward et al., 2021). Whether Srsf1 localizes to DynAPs is not known, but the related factor Sf3a3 does strongly localize to DynAPs in addition to its expected localization in nuclear foci (Lee et al., 2020).

DynAPs might also play a broader role in MCC protein homeostasis. Cytoplasmic Srsf1 also controls the translation of certain DNAAFs in addition to ODA and IDAs subunits (Haward et al., 2021). Moreover, Pih1d3/Dnaaf6 was recently found to interact with the RNA binding protein Larp6. These proteins localize together in DynAPs, but rather than controlling the machinery responsible for cilia beating, they control ciliogenesis. Larp6 binds to tubulin mRNAs and controls tubulin protein levels in MCCs (Earwood et al., 2024). To determine if DynAPs contain additional factors involved in MCC homeostasis, further exploration of the biochemistry within DynAPs will be a welcome future direction.

Finally, it is also possible that DynAPs play additional roles as platforms for the transport of IDAs and ODAs from the cytoplasm to the ciliary axoneme. Though this idea has not yet been explored in detail, we recently identified a set of proteins that provide an ‘address’ for binding of each of the IDAs to a CCDC39-CCDC40 dimer on the axoneme (Brody et al., 2025). Termed ciliary address recognition proteins (CARPs), it is possible that these proteins and their unique interactions with different IDAs are initiated in DynAPs en route to the intraflagellar transport system.

Other puncta, other foci

Notably, several other seemingly unrelated membraneless organelles have recently been implicated in ciliary biology. Of these, the most interesting are the cytoplasmic foci related to the function of the central pair apparatus of motile cilia (Fig. 1B, orange). The Hoatz protein acts in the cytoplasm to process enolase 4 (Eno4), which is a component of the central pair apparatus required for ciliary beating (Narita et al., 2020). Mice lacking Hoatz display cilia beating defects and excessive accumulation of Eno4-positive foci in the cytoplasm. Notably, the Hoatz interactome resembles the known roster of DynAP proteins, enriched for proteins factors related to HSP chaperones and RNA processing (Narita et al., 2020). Additionally, like the dynein arms and the central pair apparatus, radial spoke complexes (Fig. 1B, green) are also pre-assembled in the cytoplasm (Diener et al., 2011). Whether a membraneless organelle is involved in that process is unknown.

Finally, two other cilia-related membraneless organelles have been described. First, trinucleotide repeat-containing gene 6A (Tnrc6a) has been previously associated with microRNA-based gene silencing but was recently found to be highly expressed in MCCs (Liu et al., 2023). In these cells, Tnrc6a is present in two populations of cytosolic foci: a population of large basolaterally positioned P-bodies and a population of foci in the apical cytoplasm. These latter foci are also sites of nascent translation and loss of Tnrc6a leads to reduced levels of several proteins involved in centriole amplification (Liu et al., 2023). Second, the intrinsically disordered protein brain-expressed X-linked protein 1 (Bex1) was recently found to form foci near the base of non-motile primary cilia, where it helps to concentrate tubulin, thereby facilitating microtubule polymerization and contributing to primary ciliogenesis (Hibino et al., 2022).

Outlook

The description of DynAPs has provided a unifying cell biological locus for the action of DNAAFs and chaperones in the assembly of axonemal dyneins, but many key challenges and open questions remain.

First, we lack a comprehensive understanding of DynAPs across cell types and organisms. For example, though cytoplasmic pre-assembly of dyneins and the DNAAFs were first identified in Chlamydomonas (Fowkes and Mitchell, 1998; Omran et al., 2008), reports of foci or puncta similar to DynAPs are largely lacking in that species. That being said, Cfap298/Dnaaf16 is present in perinuclear foci in mammalian tracheal MCCs and similar foci are observed for the Chlamydomonas orthologue FBB18 (Austin-Tse et al., 2013; Wang et al., 2022). Likewise, a recent report demonstrates that Wdr92/Dnaaf10 is present in cytosolic foci in Chlamydomonas (Liu et al., 2019). Furthermore, Drosophila kl granules seem to be similar to DynAPs, but more in-depth comparative work will be required to determine the extent of that similarity. Even within vertebrate animals, DynAPs remain poorly defined, with most work focused on the Xenopus embryo epidermis or mammalian airway epithelia. DynAPs in the MCCs of the oviduct or brain have not been well-studied and whether equivalent organelles are present in cells that bear rotary beating cilia with “9+0” axonemal microtubule arrangement, for example in the left/right organizer, is unknown. More thorough comparative analysis of DynAPs will be welcome.

A complementary problem relates to changes in DynAP structure and/or function over time. DynAPs have been observed in relatively mature MCCs in Xenopus, fish and mammals, but MCC “maturity” remains a vague concept. In Xenopus, DynAPs were observed in MCCs bearing maximally elongated cilia, but recent work has revealed that the deployment of ciliary machinery is not yet complete at this stage (Huizar et al., 2018; Lee et al., 2023). Moreover, full polarization of ciliary beating is not achieved until even later (Mitchell et al., 2007). Likewise, mammalian MCCs in culture undergo a remarkably lengthy maturation process that continues many weeks after cilia achieve full length and begin to beat (Oltean et al., 2018). If DynAPs serve simply as sites for assembly of nascent ODA and IDAs, we might expect them to be reduced or eliminated once cells reach full maturity, since the basal rate of axonemal dynein turnover is low.

On the other hand, it is also possible that DynAPs still function in such mature cells. For example, time courses of MCC differentiation reveal that genes encoding axonemal dyneins remain undiminished long after cilia reach their full length (Bukowy-Bieryłło et al., 2022; Carson et al., 2002). This raises the possibility that axonemal dyneins are still being made and/or stored in homeostatic MCCs, which may relate to the findings of a recent study in Xenopus MCCs that showed very rapid regeneration of at least partial cilia even when translation is inhibited (Rao et al., 2025).

In sum, there is still much to be done, and further studies of these fascinating “organelles” are clearly warranted. We believe the breadth of DynAP studies to date is a strength, with work spanning from single-celled organisms to flies, frogs, fish, mice and humans. We hope that to this repertoire can be added methods for temporal control of DNAAF function, such as temperature-sensitive mutants or chemical or opto-genetic approaches. Indeed, a temperature-sensitive ODA mutant in a Tetrahymena ODA subunit recently proved useful in studies of Dnaaf9/Shulin (Mali et al., 2021). Finally, a new protein construct that when targeted to phase-separated organelles causes their disassembly, called a “killswitch” (Zhang et al., 2025) this tool may prove to be very useful indeed for understanding these curious structures we call DynAPs.

References:

  1. Austin-Tse C, Halbritter J, Zariwala MA, Gilberti RM, Gee HY, Hellman N, Pathak N, Liu Y, Panizzi JR, Patel-King RS, Tritschler D, Bower R, O’Toole E, Porath JD, Hurd TW, Chaki M, Diaz KA, Kohl S, Lovric S, Hwang DY, Braun DA, Schueler M, Airik R, Otto EA, Leigh MW, Noone PG, Carson JL, Davis SD, Pittman JE, Ferkol TW, Atkinson JJ, Olivier KN, Sagel SD, Dell SD, Rosenfeld M, Milla CE, Loges NT, Omran H, Porter ME, King SM, Knowles MR, Drummond IA, and Hildebrandt F. 2013. Zebrafish Ciliopathy Screen Plus Human Mutational Analysis Identifies C21orf59 and CCDC65 Defects as Causing Primary Ciliary Dyskinesia. Am J Hum Genet. 93:672–686. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Banani SF, Lee HO, Hyman AA, and Rosen MK. 2017. Biomolecular condensates: organizers of cellular biochemistry. Nat Rev Mol Cell Biol. 18:285–298. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bonnefoy S, Watson CM, Kernohan KD, Lemos M, Hutchinson S, Poulter JA, Crinnion LA, Berry I, Simmonds J, Vasudevan P, O’Callaghan C, Hirst RA, Rutman A, Huang L, Hartley T, Grynspan D, Moya E, Li C, Carr IM, Bonthron DT, Leroux M, Boycott KM, Bastin P, and Sheridan EG. 2018. Biallelic Mutations in LRRC56, Encoding a Protein Associated with Intraflagellar Transport, Cause Mucociliary Clearance and Laterality Defects. Am J Hum Genet. 103:727–739. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Braschi B, Omran H, Witman GB, Pazour GJ, Pfister KK, Bruford EA, and King SM. 2022. Consensus nomenclature for dyneins and associated assembly factors. J Cell Biol. 221. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Brody SL, Pan J, Huang T, Xu J, Xu H, Koenitizer JR, Brennan SK, Nanjundappa R, Saba TG, Rumman N, Berical A, Hawkins FJ, Wang X, Zhang R, Mahjoub MR, Horani A, and Dutcher SK. 2025. Undocking of an extensive ciliary network induces proteostasis and cell fate switching resulting in severe primary ciliary dyskinesia. Sci Transl Med. 17:eadp5173. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Brosseau GE 1960. Genetic Analysis of the Male Fertility Factors on the Y Chromosome of Drosophila Melanogaster. Genetics. 45:257–274. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Bukowy-Bieryłło Z, Daca-Roszak P, Jurczak J, Przystałowska-Macioła H, Jaksik R, Witt M, and Ziętkiewicz E. 2022. In vitro differentiation of ciliated cells in ALI-cultured human airway epithelium - The framework for functional studies on airway differentiation in ciliopathies. Eur J Cell Biol. 101:151189. [DOI] [PubMed] [Google Scholar]
  8. Carson JL, Reed W, Lucier T, Brighton L, Gambling TM, Huang CH, and Collier AM. 2002. Axonemal dynein expression in human fetal tracheal epithelium. Am J Physiol Lung Cell Mol Physiol. 282:L421–430. [DOI] [PubMed] [Google Scholar]
  9. Cho KJ, Noh SH, Han SM, Choi WI, Kim HY, Yu S, Lee JS, Rim JH, Lee MG, Hildebrandt F, and Gee HY. 2018. ZMYND10 stabilizes intermediate chain proteins in the cytoplasmic pre-assembly of dynein arms. PLoS Genet. 14:e1007316. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Cloutier P, Poitras C, Durand M, Hekmat O, Fiola-Masson É, Bouchard A, Faubert D, Chabot B, and Coulombe B. 2017. R2TP/Prefoldin-like component RUVBL1/RUVBL2 directly interacts with ZNHIT2 to regulate assembly of U5 small nuclear ribonucleoprotein. Nat Commun. 8:15615. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Dauden MI, López-Perrote A, and Llorca O. 2021. RUVBL1-RUVBL2 AAA-ATPase: a versatile scaffold for multiple complexes and functions. Curr Opin Struct Biol. 67:78–85. [DOI] [PubMed] [Google Scholar]
  12. Desai PB, Dean AB, and Mitchell DR. 2018. Cytoplasmic preassembly and trafficking of axonemal dyneins. In Dyneins: Structure, Biology, and Disease. Vol. 1. King SM, editor. Academic Press/Elsevier, London. [Google Scholar]
  13. Diener DR, Yang P, Geimer S, Cole DG, Sale WS, and Rosenbaum JL. 2011. Sequential assembly of flagellar radial spokes. Cytoskeleton (Hoboken). 68:389– 400. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Drew K, Lee C, Cox RM, Dang V, Devitt CC, McWhite CD, Papoulas O, Huizar RL, Marcotte EM, and Wallingford JB. 2020. A systematic, label-free method for identifying RNA-associated proteins in vivo provides insights into vertebrate ciliary beating machinery. Dev Biol. 467:108–117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Duquesnoy P, Escudier E, Vincensini L, Freshour J, Bridoux AM, Coste A, Deschildre A, de Blic J, Legendre M, Montantin G, Tenreiro H, Vojtek AM, Loussert C, Clément A, Escalier D, Bastin P, Mitchell DR, and Amselem S. 2009. Loss-of-function mutations in the human ortholog of Chlamydomonas reinhardtii ODA7 disrupt dynein arm assembly and cause primary ciliary dyskinesia. Am J Hum Genet. 85:890–896. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Dutcher SK 2020. Asymmetries in the cilia of Chlamydomonas. Philos Trans R Soc Lond B Biol Sci. 375:20190153. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Earwood R, Ninomiya H, Wang H, Shimada IS, Stroud M, Perez D, Uuganbayar U, Yamada C, Akiyama-Miyoshi T, Stefanovic B, and Kato Y. 2024. The binding of LARP6 and DNAAF6 in biomolecular condensates influences ciliogenesis of multiciliated cells. J Biol Chem. 300:107373. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Fabczak H, and Osinka A. 2019. Role of the Novel Hsp90 Co-Chaperones in Dynein Arms’ Preassembly. Int J Mol Sci. 20. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Fassad MR, Shoemark A, le Borgne P, Koll F, Patel M, Dixon M, Hayward J, Richardson C, Frost E, Jenkins L, Cullup T, Chung EMK, Lemullois M, Aubusson-Fleury A, Hogg C, Mitchell DR, Tassin AM, and Mitchison HM. 2018. C11orf70 Mutations Disrupting the Intraflagellar Transport-Dependent Assembly of Multiple Axonemal Dyneins Cause Primary Ciliary Dyskinesia. Am J Hum Genet. 102:956–972. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Feric M, Vaidya N, Harmon TS, Mitrea DM, Zhu L, Richardson TM, Kriwacki RW, Pappu RV, and Brangwynne CP. 2016. Coexisting Liquid Phases Underlie Nucleolar Subcompartments. Cell. 165:1686–1697. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Fingerhut JM, and Yamashita YM. 2020. mRNA localization mediates maturation of cytoplasmic cilia in Drosophila spermatogenesis. J Cell Biol. 219. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Fingerhut JM, Moran JV, and Yamashita YM. 2019. Satellite DNA-containing gigantic introns in a unique gene expression program during Drosophila spermatogenesis. PLoS Genet. 15:e1008028. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Fowkes ME, and Mitchell DR. 1998. The role of preassembled cytoplasmic complexes in assembly of flagellar dynein subunits. Mol Biol Cell. 9:2337–2347. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Glauninger H, Wong Hickernell CJ, Bard JAM, and Drummond DA. 2022. Stressful steps: Progress and challenges in understanding stress-induced mRNA condensation and accumulation in stress granules. Mol Cell. 82:2544–2556. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Grossman-Haham I 2023. Towards an atomic model of a beating ciliary axoneme. Curr Opin Struct Biol. 78:102516. [DOI] [PubMed] [Google Scholar]
  26. Haward F, Maslon MM, Yeyati PL, Bellora N, Hansen JN, Aitken S, Lawson J, von Kriegsheim A, Wachten D, Mill P, Adams IR, and Caceres JF. 2021. Nucleo-cytoplasmic shuttling of splicing factor SRSF1 is required for development and cilia function. Elife. 10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Hibino E, Ichiyama Y, Tsukamura A, Senju Y, Morimune T, Ohji M, Maruo Y, Nishimura M, and Mori M. 2022. Bex1 is essential for ciliogenesis and harbours biomolecular condensate-forming capacity. BMC Biol. 20:42. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Höben IM, Hjeij R, Olbrich H, Dougherty GW, Nöthe-Menchen T, Aprea I, Frank D, Pennekamp P, Dworniczak B, Wallmeier J, Raidt J, Nielsen KG, Philipsen MC, Santamaria F, Venditto L, Amirav I, Mussaffi H, Prenzel F, Wu K, Bakey Z, Schmidts M, Loges NT, and Omran H. 2018. Mutations in C11orf70 Cause Primary Ciliary Dyskinesia with Randomization of Left/Right Body Asymmetry Due to Defects of Outer and Inner Dynein Arms. Am J Hum Genet. 102:973–984. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Horani A, Ustione A, Huang T, Firth AL, Pan J, Gunsten SP, Haspel JA, Piston DW, and Brody SL. 2018. Establishment of the early cilia preassembly protein complex during motile ciliogenesis. Proc Natl Acad Sci U S A. 115:E1221– E1228. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Horani A, Druley TE, Zariwala MA, Patel AC, Levinson BT, Van Arendonk LG, Thornton KC, Giacalone JC, Albee AJ, Wilson KS, Turner EH, Nickerson DA, Shendure J, Bayly PV, Leigh MW, Knowles MR, Brody SL, Dutcher SK, and Ferkol TW. 2012. Whole-exome capture and sequencing identifies HEATR2 mutation as a cause of primary ciliary dyskinesia. Am J Hum Genet. 91:685–693. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Horani A, Ferkol TW, Shoseyov D, Wasserman MG, Oren YS, Kerem B, Amirav I, Cohen-Cymberknoh M, Dutcher SK, Brody SL, Elpeleg O, and Kerem E. 2013. LRRC6 mutation causes primary ciliary dyskinesia with dynein arm defects. PLoS One. 8:e59436. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Horani A, Ferkol TW, Dutcher SK, and Brody SL. 2016. Genetics and biology of primary ciliary dyskinesia. Paediatr Respir Rev. 18:18–24. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Huizar RL, Lee C, Boulgakov AA, Horani A, Tu F, Marcotte EM, Brody SL, and Wallingford JB. 2018. A liquid-like organelle at the root of motile ciliopathy. Elife. 7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Jain S, Wheeler JR, Walters RW, Agrawal A, Barsic A, and Parker R. 2016. ATPase-Modulated Stress Granules Contain a Diverse Proteome and Substructure. Cell. 164:487–498. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Kamiya R, and Yagi T. 2014. Functional diversity of axonemal dyneins as assessed by in vitro and in vivo motility assays of Chlamydomonas mutants. Zoological science. 31:633–645. [DOI] [PubMed] [Google Scholar]
  36. King SM 2018. Composition and assembly of axonemal dyneins. In Dyneins. Elsevier. 162–201. [Google Scholar]
  37. King SM 2021. Cytoplasmic factories for axonemal dynein assembly. J Cell Sci. 134. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Kishimoto N, Cao Y, Park A, and Sun Z. 2008. Cystic kidney gene seahorse regulates cilia-mediated processes and Wnt pathways. Dev Cell. 14:954–961. [DOI] [PubMed] [Google Scholar]
  39. Klingauf M, Stanek D, and Neugebauer KM. 2006. Enhancement of U4/U6 small nuclear ribonucleoprotein particle association in Cajal bodies predicted by mathematical modeling. Mol Biol Cell. 17:4972–4981. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Knowles MR, Ostrowski LE, Loges NT, Hurd T, Leigh MW, Huang L, Wolf WE, Carson JL, Hazucha MJ, Yin W, Davis SD, Dell SD, Ferkol TW, Sagel SD, Olivier KN, Jahnke C, Olbrich H, Werner C, Raidt J, Wallmeier J, Pennekamp P, Dougherty GW, Hjeij R, Gee HY, Otto EA, Halbritter J, Chaki M, Diaz KA, Braun DA, Porath JD, Schueler M, Baktai G, Griese M, Turner EH, Lewis AP, Bamshad MJ, Nickerson DA, Hildebrandt F, Shendure J, Omran H, and Zariwala MA. 2013. Mutations in SPAG1 cause primary ciliary dyskinesia associated with defective outer and inner dynein arms. Am J Hum Genet. 93:711–720. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Kott E, Legendre M, Copin B, Papon JF, Dastot-Le Moal F, Montantin G, Duquesnoy P, Piterboth W, Amram D, Bassinet L, Beucher J, Beydon N, Deneuville E, Houdouin V, Journel H, Just J, Nathan N, Tamalet A, Collot N, Jeanson L, Le Gouez M, Vallette B, Vojtek AM, Epaud R, Coste A, Clement A, Housset B, Louis B, Escudier E, and Amselem S. 2013. Loss-of-function mutations in RSPH1 cause primary ciliary dyskinesia with central-complex and radial-spoke defects. Am J Hum Genet. 93:561–570. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Kott E, Duquesnoy P, Copin B, Legendre M, Dastot-Le Moal F, Montantin G, Jeanson L, Tamalet A, Papon JF, Siffroi JP, Rives N, Mitchell V, de Blic J, Coste A, Clement A, Escalier D, Toure A, Escudier E, and Amselem S. 2012. Loss-of-function mutations in LRRC6, a gene essential for proper axonemal assembly of inner and outer dynein arms, cause primary ciliary dyskinesia. Am J Hum Genet. 91:958–964. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Lee C, Cox RM, Papoulas O, Horani A, Drew K, Devitt CC, Brody SL, Marcotte EM, and Wallingford JB. 2020. Functional partitioning of a liquid-like organelle during assembly of axonemal dyneins. Elife. 9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Lee C, Ma Y, Tu F, and Wallingford JB. 2023. Ordered deployment of distinct ciliary beating machines in growing axonemes of vertebrate multiciliated cells. Differentiation. 131:49–58. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Leslie JS, Hjeij R, Vivante A, Bearce EA, Dyer L, Wang J, Rawlins L, Kennedy J, Ubeyratna N, Fasham J, Irons ZH, Craig SB, Koenig J, George S, Pode- Shakked B, Bolkier Y, Barel O, Mane S, Frederiksen KK, Wenger O, Scott E, Cross HE, Lorentzen E, Norris DP, Anikster Y, Omran H, Grimes DT, Crosby AH, and Baple EL. 2022. Biallelic DAW1 variants cause a motile ciliopathy characterized by laterality defects and subtle ciliary beating abnormalities. Genet Med. 24:2249–2261. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Li Y, Zhao L, Yuan S, Zhang J, and Sun Z. 2017. Axonemal dynein assembly requires the R2TP complex component Pontin. Development. 144:4684–4693. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Li Y, Xu W, Cheng Y, Djenoune L, Zhuang C, Cox AL, Britto CJ, Yuan S, Wang S, and Sun Z. 2024. Cotranslational molecular condensation of cochaperones and assembly factors facilitates axonemal dynein biogenesis. Proc Natl Acad Sci U S A. 121:e2402818121. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Liu G, Wang L, and Pan J. 2019. Chlamydomonas WDR92 in association with R2TP-like complex and multiple DNAAFs to regulate ciliary dynein preassembly. J Mol Cell Biol. 11:770–780. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Liu H, Li H, Jiang Z, Jin S, Song R, Yang Y, Li J, Huang J, Zhang X, Dong X, Mori M, Fritzler MJ, He L, Cardoso WV, and Lu J. 2023. A local translation program regulates centriole amplification in the airway epithelium. Sci Rep. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Loges NT, Olbrich H, Becker-Heck A, Haffner K, Heer A, Reinhard C, Schmidts M, Kispert A, Zariwala MA, Leigh MW, Knowles MR, Zentgraf H, Seithe H, Nurnberg G, Nurnberg P, Reinhardt R, and Omran H. 2009. Deletions and point mutations of LRRC50 cause primary ciliary dyskinesia due to dynein arm defects. Am J Hum Genet. 85:883–889. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Lynham J, and Houry WA. 2022. The Role of Hsp90-R2TP in Macromolecular Complex Assembly and Stabilization. Biomolecules. 12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Mali GR, Ali FA, Lau CK, Begum F, Boulanger J, Howe JD, Chen ZA, Rappsilber J, Skehel M, and Carter AP. 2021. Shulin packages axonemal outer dynein arms for ciliary targeting. Science. 371:910–916. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Mali GR, Yeyati PL, Mizuno S, Dodd DO, Tennant PA, Keighren MA, Zur Lage P, Shoemark A, Garcia-Munoz A, Shimada A, Takeda H, Edlich F, Takahashi S, von Kreigsheim A, Jarman AP, and Mill P. 2018. ZMYND10 functions in a chaperone relay during axonemal dynein assembly. Elife. 7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Maurizy C, Quinternet M, Abel Y, Verheggen C, Santo PE, Bourguet M, C.F.P. A, Bragantini B, Chagot ME, Robert MC, Abeza C, Fabre P, Fort P, Vandermoere F, P MFS, Rain JC, Charpentier B, Cianférani S, Bandeiras TM, Pradet-Balade B, Manival X, and Bertrand E. 2018. The RPAP3-Cterminal domain identifies R2TP-like quaternary chaperones. Nat Commun. 9:2093. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. McSwiggen DT, Mir M, Darzacq X, and Tjian R. 2019. Evaluating phase separation in live cells: diagnosis, caveats, and functional consequences. Genes Dev. 33:1619–1634. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Mitchell B, Jacobs R, Li J, Chien S, and Kintner C. 2007. A positive feedback mechanism governs the polarity and motion of motile cilia. Nature. 447:97. [DOI] [PubMed] [Google Scholar]
  57. Mitchison HM, and Valente EM. 2017. Motile and non-motile cilia in human pathology: from function to phenotypes. J Pathol. 241:294–309. [DOI] [PubMed] [Google Scholar]
  58. Mitchison HM, Schmidts M, Loges NT, Freshour J, Dritsoula A, Hirst RA, O’Callaghan C, Blau H, Al Dabbagh M, Olbrich H, Beales PL, Yagi T, Mussaffi H, Chung EM, Omran H, and Mitchell DR. 2012. Mutations in axonemal dynein assembly factor DNAAF3 cause primary ciliary dyskinesia. Nat Genet. 44:381–389, S381–382. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Narita K, Nagatomo H, Kozuka-Hata H, Oyama M, and Takeda S. 2020. Discovery of a Vertebrate-Specific Factor that Processes Flagellar Glycolytic Enolase during Motile Ciliogenesis. iScience. 23:100992. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Olcese C, Patel MP, Shoemark A, Kiviluoto S, Legendre M, Williams HJ, Vaughan CK, Hayward J, Goldenberg A, Emes RD, Munye MM, Dyer L, Cahill T, Bevillard J, Gehrig C, Guipponi M, Chantot S, Duquesnoy P, Thomas L, Jeanson L, Copin B, Tamalet A, Thauvin-Robinet C, Papon JF, Garin A, Pin I, Vera G, Aurora P, Fassad MR, Jenkins L, Boustred C, Cullup T, Dixon M, Onoufriadis A, Bush A, Chung EM, Antonarakis SE, Loebinger MR, Wilson R, Armengot M, Escudier E, Hogg C, U.K.R. Group, Amselem S, Sun Z, Bartoloni L, Blouin JL, and Mitchison HM. 2017. X-linked primary ciliary dyskinesia due to mutations in the cytoplasmic axonemal dynein assembly factor PIH1D3. Nat Commun. 8:14279. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Oltean A, Schaffer AJ, Bayly PV, and Brody SL. 2018. Quantifying Ciliary Dynamics during Assembly Reveals Stepwise Waveform Maturation in Airway Cells. Am J Respir Cell Mol Biol. 59:511–522. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Omran H, Kobayashi D, Olbrich H, Tsukahara T, Loges NT, Hagiwara H, Zhang Q, Leblond G, O’Toole E, Hara C, Mizuno H, Kawano H, Fliegauf M, Yagi T, Koshida S, Miyawaki A, Zentgraf H, Seithe H, Reinhardt R, Watanabe Y, Kamiya R, Mitchell DR, and Takeda H. 2008. Ktu/PF13 is required for cytoplasmic pre-assembly of axonemal dyneins. Nature. 456:611–616. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Panizzi JR, Becker-Heck A, Castleman VH, Al-Mutairi DA, Liu Y, Loges NT, Pathak N, Austin-Tse C, Sheridan E, Schmidts M, Olbrich H, Werner C, Haffner K, Hellman N, Chodhari R, Gupta A, Kramer-Zucker A, Olale F, Burdine RD, Schier AF, O’Callaghan C, Chung EM, Reinhardt R, Mitchison HM, King SM, Omran H, and Drummond IA. 2012. CCDC103 mutations cause primary ciliary dyskinesia by disrupting assembly of ciliary dynein arms. Nat Genet. 44:714–719. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Philippe M, Salloum S, Slimani F, Chassé H, Robert M-C, Urbach S, Imbert J, Séveno M, Georges S, Boulon S, Verheggen C, and Bertrand E. 2025. Co- translational determination of quaternary structures in chaperone factories. bioRxiv:2025.2002.2012.637890. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Qiu T, and Roy S. 2022. Ciliary dynein arms: Cytoplasmic preassembly, intraflagellar transport, and axonemal docking. J Cell Physiol. [DOI] [PubMed] [Google Scholar]
  66. Rao VG, Subramanianbalachandar VA, Magaj MM, Redemann S, and Kulkarni SS. 2025. Mechanisms of cilia regeneration in Xenopus multiciliated epithelium in vivo. EMBO Rep. 26:2192–2220. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Serluca FC, Xu B, Okabe N, Baker K, Lin SY, Sullivan-Brown J, Konieczkowski DJ, Jaffe KM, Bradner JM, Fishman MC, and Burdine RD. 2009. Mutations in zebrafish leucine-rich repeat-containing six-like affect cilia motility and result in pronephric cysts, but have variable effects on left-right patterning. Development. 136:1621–1631. [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Smith AJ, Bustamante-Marin XM, Yin W, Sears PR, Herring LE, Dicheva NN, López-Giráldez F, Mane S, Tarran R, Leigh MW, Knowles MR, Zariwala MA, and Ostrowski LE. 2022. The role of SPAG1 in the assembly of axonemal dyneins in human airway epithelia. J Cell Sci. 135. [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Tarkar A, Loges NT, Slagle CE, Francis R, Dougherty GW, Tamayo JV, Shook B, Cantino M, Schwartz D, Jahnke C, Olbrich H, Werner C, Raidt J, Pennekamp P, Abouhamed M, Hjeij R, Kohler G, Griese M, Li Y, Lemke K, Klena N, Liu X, Gabriel G, Tobita K, Jaspers M, Morgan LC, Shapiro AJ, Letteboer SJ, Mans DA, Carson JL, Leigh MW, Wolf WE, Chen S, Lucas JS, Onoufriadis A, Plagnol V, Schmidts M, Boldt K, Roepman R, Zariwala MA, Lo CW, Mitchison HM, Knowles MR, Burdine RD, Loturco JJ, and Omran H. 2013. DYX1C1 is required for axonemal dynein assembly and ciliary motility. Nat Genet. 45:995–1003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Tu F, Sedzinski J, Ma Y, Marcotte EM, and Wallingford JB. 2018. Protein localization screening in vivo reveals novel regulators of multiciliated cell development and function. J Cell Sci. 131. [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Walentek P, and Quigley IK. 2017. What we can learn from a tadpole about ciliopathies and airway diseases: Using systems biology in Xenopus to study cilia and mucociliary epithelia. Genesis. 55. [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Wallmeier J, Nielsen KG, Kuehni CE, Lucas JS, Leigh MW, Zariwala MA, and Omran H. 2020. Motile ciliopathies. Nat Rev Dis Primers. 6:77. [DOI] [PubMed] [Google Scholar]
  73. Wang L, Li X, Liu G, and Pan J. 2022. FBB18 participates in preassembly of almost all axonemal dyneins independent of R2TP complex. PLoS Genet. 18:e1010374. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Wheeler JR, Matheny T, Jain S, Abrisch R, and Parker R. 2016. Distinct stages in stress granule assembly and disassembly. Elife. 5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Yamaguchi H, Oda T, Kikkawa M, and Takeda H. 2018. Systematic studies of all PIH proteins in zebrafish reveal their distinct roles in axonemal dynein assembly. eLife. 7:e36979. [DOI] [PMC free article] [PubMed] [Google Scholar]
  76. Zariwala MA, Gee HY, Kurkowiak M, Al-Mutairi DA, Leigh MW, Hurd TW, Hjeij R, Dell SD, Chaki M, Dougherty GW, Adan M, Spear PC, Esteve-Rudd J, Loges NT, Rosenfeld M, Diaz KA, Olbrich H, Wolf WE, Sheridan E, Batten TF, Halbritter J, Porath JD, Kohl S, Lovric S, Hwang DY, Pittman JE, Burns KA, Ferkol TW, Sagel SD, Olivier KN, Morgan LC, Werner C, Raidt J, Pennekamp P, Sun Z, Zhou W, Airik R, Natarajan S, Allen SJ, Amirav I, Wieczorek D, Landwehr K, Nielsen K, Schwerk N, Sertic J, Kohler G, Washburn J, Levy S, Fan S, Koerner-Rettberg C, Amselem S, Williams DS, Mitchell BJ, Drummond IA, Otto EA, Omran H, Knowles MR, and Hildebrandt F. 2013. ZMYND10 is mutated in primary ciliary dyskinesia and interacts with LRRC6. Am J Hum Genet. 93:336–345. [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Zhang Y, Stöppelkamp I, Fernandez-Pernas P, Allram M, Charman M, Magalhaes AP, Piedavent-Salomon M, Sommer G, Sung YC, Meyer K, Grams N, Halko E, Dongre S, Meierhofer D, Malszycki M, Ilik IA, Aktas T, Kraushar ML, Vastenhouw N, Weitzman MD, Grebien F, Niskanen H, and Hnisz D. 2025. Probing condensate microenvironments with a micropeptide killswitch. Nature. 643:1107–1116. [DOI] [PMC free article] [PubMed] [Google Scholar]
  78. Zhao L, Yuan S, Cao Y, Kallakuri S, Li Y, Kishimoto N, DiBella L, and Sun Z. 2013. Reptin/Ruvbl2 is a Lrrc6/Seahorse interactor essential for cilia motility. Proc Natl Acad Sci U S A. 110:12697–12702. [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Zur Lage P, Stefanopoulou P, Styczynska-Soczka K, Quinn N, Mali G, von Kriegsheim A, Mill P, and Jarman AP. 2018. Ciliary dynein motor preassembly is regulated by Wdr92 in association with HSP90 co-chaperone, R2TP. J Cell Biol. 217:2583–2598. [DOI] [PMC free article] [PubMed] [Google Scholar]

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