Zebrafish Motile Cilia: Comparison
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Please note this is a comparison between Version 3 by Conner Chen and Version 2 by Rita Xu.

Zebrafish is a vertebrate teleost widely used in many areas of research. As embryos, they develop quickly and provide unique opportunities for research studies owing to their transparency for at least 48 h post fertilization. Zebrafish have many ciliated organs that include primary cilia as well as motile cilia. Using zebrafish as an animal model helps to better understand human diseases such as Primary Ciliary Dyskinesia (PCD), an autosomal recessive disorder that affects cilia motility, currently associated with more than 50 genes. 

  • transmission electron microscopy
  • motile cilia
  • left–right organizer

1. Introduction

Motile cilia are centriole-derived organelles, surrounded by a membrane and containing microtubules formed by protofilaments that can be longer or shorter, depending on the number of tubulin molecules they contain, according to the function they perform [1][2][3]. This structure is called the ciliary axoneme or ciliary shaft [4], a transversal term throughout species, due to the well-conserved ciliary structure [3]. The axonemal ultrastructure in eukaryotic cells segregates into two significant patterns: 9 + 2, in which nine microtubule doublets organise around two microtubules in the centre known as the central pair (CP) complex; the 9 + 0 organisation, in which the CP is absent [5][6]. Moreover, there are two major types of motile ciliated cells: cells that produce from dozens to several hundred 9 + 2 motile cilia and cells that only generate one cilium.
Motile monocilia-lacking CPs were described in the node of mice [7][8][9]. These are very different from non-motile primary cilia (also with a 9 + 0 configuration) without dynein motor arms that can be found in almost every cell type having solo sensory functions and lacking the ability to generate movement [10][11]. On the other hand, Yu et al. in 2011 identified cilia with a 9 + 2 organisation but immotile, called kinocilia, in the hair cells of the inner ear that express Foxj1b [12] and in the ciliated receptor cells (primary sensory cells) of the zebrafish OP [13]. We can, thus, speculate that all combinations are present in nature, likely by adaptive evolution.
Regarding motile cilia dynamics, multiciliated cells that typically have cilia with a 9 + 2 configuration beat metachronically with a planar stroke to clear fluid or promote locomotion [14][15]. On the other hand, motile cilia lacking CP (9 + 0) have a specific movement pattern, described as planar rotation, circling or twisting. These 9 + 0 monocilia are present in the LRO in vertebrates such as mice or zebrafish [16][17][18], but reports on LRO cilia with 9 + 2 cilia and 9 + 4 also exist [7][18][19]. In cases of disease, multiciliated cells with cilia wholly or partially lacking the CP are frequently accounted as one of the phenotypes of Primary Ciliary Dyskinesia, an autosomal recessive disease of the motile cilia [20][21]. The rationale being that in the respiratory cilia, a rotational movement is not efficient in mucociliary clearance and is, therefore, causative of PCD symptoms.
Zebrafish became popular as an animal model in the 1980s, triggered by George Streisinger’s studies showing zebrafish as a genetically tractable organism, allowing a phenotypic characterisation of a large number of mutations that cause defects in a variety of organ systems [22]. As zebrafish are vertebrates, the translational interpretation of ciliary defects is very powerful, when compared, for instance, to Chlamydomonas reinhardtii, where the defects from cilia impairment are usually spotted as a lack of locomotion [23], although not exclusively. Zebrafish helped shed light on the role of specific genes in human diseases, as their genome has been widely studied, providing insight into their human orthologues [24]. Amongst the many advantages of zebrafish, we will stress the fact that zebrafish embryos contain cilia in nearly every cell type, and their organogenesis defects can be easily characterised using brightfield and fluorescent microscopy as the zebrafish embryo and larvae are mostly transparent until all major organ systems are formed. Many of the organs and tissues of zebrafish are similar to those of humans and 70% of genes are shared [24]. Motile cilia have been well described in the Kupffer’s vesicle, the zebrafish LRO, between 3 and 14 somite stages (ss) of development and in the OP at 48–72 hpf, amongst other locations [25].
Zebrafish, as almost all teleosts, have three types of olfactory receptor neurons (ORNs): ciliated ORNs, microvillous ORNs and crypt cells with both cilia and microvilli [26][27]. The bottom of the OP is coated with ORNs, each one with protruding non-motile primary cilia that contain olfactory receptors [28]. On the other hand, the rim is surrounded by a layer of cuboidal multiciliated cells protruding bundles of motile cilia [28]. These microvillous ORNs, and ciliated ORNs have similar morphological and molecular contents to the microvillous and ciliated ORNs of higher vertebrates [29]. Moreover, genetic engineering methodologies, including CRISPR/Cas9, have been successfully used in editing the zebrafish genome and have greatly facilitated the generation of zebrafish mutant models mimicking human ciliopathies [30][31][32].
PCD is a genetic disorder distinguished by recurrent infection in the lower and upper respiratory tract [33], reduced fertility and laterality problems (50% of PCD patients show situs inversus) [34]. Ultrastructural defects in motile cilia or a reduced cilia number are known to cause PCD [34][35]. Currently there are more than 50 genes identified that can cause PCD [36]. Almost all PCD genes show homologous genes in zebrafish, as shown in Table 1. Despite previous research on zebrafish cilia [13][18][26][37], no study fully characterised the cilia ultrastructure of the OP and LRO. With this work, we aim to determine the similarities and the significant differences between zebrafish and human motile cilia, comparing OP multiciliated cells and embryonic LRO monociliated cells. Using transmission electron microscopy (TEM), we confirmed the heterogenic configuration of the zebrafish OP cells. We also showed by electron tomography (ET) the variable presence of a CP in the LRO monocilia, a feature hypothesised in a previous publication by Tavares et al. [18].
Table 1. Human-Zebrafish homologue PCD genes as described in Ensemble.org [38].
PCD Gene Zebrafish Transcript Name Zebrafish Transcript ID
DNAH5 dnah5-201 ENSDART00000123150.4
dnah5-202 ENSDART00000191818.1
CCDC114 (ODAD1) ccdc114-201 ENSDART00000023745.8
ARMC4 (ODAD2) cr847789.1-201 ENSDART00000186851.1
armc4-201 ENSDART00000077453.5
armc4-204 ENSDART00000170018.2
armc4-203 ENSDART00000153115.2
armc4-202 ENSDART00000152887.2
TTC25 (ODAD4) ttc25-201 ENSDART00000080946.5
DNAH9 dnah9-201 ENSDART00000160926.2
DNAH11 dnah11-201 ENSDART00000148294.4
dnah11-202 ENSDART00000020821.10
dnah11-203 ENSDART00000138744.4
DNAI1 dnai1.2-201 ENSDART00000080431.5
dnai1.2-202 ENSDART00000142468.3
dnai1.1-205 ENSDART00000170205.2
dnai1.1-201 ENSDART00000160163.2
dnai1.1-204 ENSDART00000169676.2
dnai1.1-202 ENSDART00000163063.2
dnai1.1-203 ENSDART00000165798.2
DNAI2 dnai2a-201 ENSDART00000162579.2
dnai2a-202 ENSDART00000164199.2
dnai2b-203 ENSDART00000188726.1
dnai2b-201 ENSDART00000003339.9
dnai2b-202 ENSDART00000188648.1
DNAL1 dnal1-203 ENSDART00000188500.1
dnal1-202 ENSDART00000156182.2
dnal1-201 ENSDART00000043651.7
TXNDC3 (NME8) nme8-201 ENSDART00000163684.2
CCDC103 ccdc103-201 ENSDART00000075493.4
ccdc103-202 ENSDART00000132293.2
CFAP298 (C21orf59) cfap298-201 ENSDART00000051197.6
cfap298-202 ENSDART00000130093.3
cfap298-203 ENSDART00000181950.1
CFAP300 (c11orf70) cfap300-201 ENSDART00000151109.2
cfap300-202 ENSDART00000192737.1
DNAAF1 (LRRC50) dnaaf1-201 ENSDART00000145762.4
dnaaf1-203 ENSDART00000173909.2
dnaaf1-202 ENSDART00000173853.2
DNAAF2 (KTU) dnaaf2-201 ENSDART00000167840.2
DNAAF3 dnaaf3l-201 ENSDART00000079233.5
DNAAF4 (DYX1C1) dnaaf4-201 ENSDART00000165855.2
DNAAF5 (HEATR2) lo018183.1-201 ENSDART00000194031.1
DNAAF6 (PIH1D3) pih1d3-201 ENSDART00000056375.5
pih1d3-203 ENSDART00000145388.3
pih1d3-202 ENSDART00000136858.2
pih1d3-204 ENSDART00000183524.1
pih1d3-205 ENSDART00000191761.1
LRRC6 lrrc6-203 ENSDART00000188883.1
lrrc6-202 ENSDART00000132346.3
lrrc6-201 ENSDART00000075347.5
RPGR rpgrb-201 ENSDART00000088624.5
rpgrb-202 ENSDART00000124471.3
rpgrip-201 ENSDART00000138541.3
rpgrip-203 ENSDART00000190953.1
rpgrip-202 ENSDART00000179003.2
rpgrip1l-202 ENSDART00000185324.1
rpgrip1l-201 ENSDART00000126326.5
SPAG1 spag1b-201 ENSDART00000101207.5
spag1a-202 ENSDART00000185960.1
spag1a-201 ENSDART00000130537.3
ZMYND10 zmynd10-201 ENSDART00000017413.10
zmynd10-202 ENSDART00000189261.1
zmynd10-203 ENSDART00000183251.1
CCDC39 ccdc39-202 ENSDART00000190769.1
ccdc39-201 ENSDART00000169709.2
CCDC40 ccdc40-202 ENSDART00000169752.2
Ccdc40-201 ENSDART00000164275.2
Ccdc40-203 ENSDART00000182267.1
TTC12 ttc12-201 ENSDART00000156234.2
ttc12-202 ENSDART00000157380.2
CCDC65 (DRC2) ccdc65-201 ENSDART00000043946.8
ccdc65-202 ENSDART00000177219.2
CCDC164 (DRC1) drc1-201 ENSDART00000061829.5
GAS8 gas8-202 ENSDART00000170982.2
gas8-201 ENSDART00000165126.2
CFAP221 not found in ZF  
DNAJB13 dnajb13-204 ENSDART00000148093.3
dnajb13-201 ENSDART00000063365.6
dnajb13-203 ENSDART00000139097.2
dnajb13-202 ENSDART00000133505.2
HYDIN hydin-201 ENSDART00000143265.4
hydin-202 ENSDART00000145701.2
hydin-203 ENSDART00000169861.2
bx571975.1-201 ENSDART00000185269.1
NME5 nme5-201 ENSDART00000060998.6
RSPH1 rsph1-201 ENSDART00000160273.3
ct573248.3-201 ENSDART00000181186.1
RSPH3 rsph3-202 ENSDART00000128823.5
rsph3-201 ENSDART00000103394.3
RSPH4a rsph4a-201 ENSDART00000097340.5
RSPH9 rsph9-201 ENSDART00000010903.8
STK36 stk36-201 ENSDART00000086765.5
stk36-202 ENSDART00000139065.2
SPEF2 spef2-201 ENSDART00000159718.2
spef2-202 ENSDART00000168984.2
CFAP57 cfap57-201 ENSDART00000080900.6
cfap57-202 ENSDART00000149309.3
LRRC56 lrrc56-202 ENSDART00000161369.2
lrrc56-201 ENSDART00000150364.2
GAS2L2 gas2l2-201 ENSDART00000112744.4
NEK10 nek10-201 ENSDART00000155162.2
OFD1 ofd1-201 ENSDART00000000552.12
CCNO fq311924.1-201 ENSDART00000158096.2
FOXJ1 foxj1b-201 ENSDART00000126676.2
foxj1b-203 ENSDART00000181942.1
foxj1b-202 ENSDART00000153327.2
foxj1a-201 ENSDART00000157772.2
foxj1a-202 ENSDART00000168280.2
MCIDAS cu633857.1-201 ENSDART00000192716.1

2. Zebrafish Motile Cilia

Zebrafish have motile cilia in many of its organs, that are present since the early stages of development, as depicted in the diagram from Figure 1.
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Figure 1. Motile ciliated structures in zebrafish. Schematic representation of (A), a zebrafish embryo at 8 somite stages (ss) highlighting Kupffer’s vesicle (KV); (B) a 3-day post-fertilization (dpf) zebrafish larva indicating structures with motile cilia. Olfactory pit (OP) and KV cilia analysed in the present study are circled in orange.
Olstad et al. have previously shown the flow generated by the OP motile cilia [39]. Whereas Sampaio et al. (2014) and Tavares et al. (2017) have extensively described the rotational and wavy fashion beat of monocilia of the LRO [16][18]. These motile monocilia are essential to generate flow for the determination of left–right asymmetry [16][18][40]. We confirmed the localization and distribution of cilia in the wild-type (WT) LRO and OP of zebrafish by immunofluorescence (IF). To better understand how to orient the embryo for the subsequent TEM embedding and sectioning, we generated 3D blend projections and surfaces as shown in Figure 2 for the OP. We consider that this study greatly helped the TEM work. Confocal microscopy is useful but limited to 180 nm lateral and 500 nm axial resolution and is not appropriate for ultrastructural studies. Therefore, we next used TEM to access the ultrastructure of the respective organ cilia.
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Figure 2. Three-dimensional imaging analysis helps to orient and localize the OP of zebrafish for TEM studies. (A,B). Immunofluorescent labelling with anti-acetylated α-tubulin shows the distribution of multiciliated cells in the OP of 4 dpf larvae. Software Imaris (Bitplane) v.9.5.0 allowed 3D blend reconstructions of 2 different OPs from 2 different larvae. (C,D) 3D surface reconstructions from the respective OPs revealing the concave morphology of the organ when rotated. Anti-Acetylated α-tubulin immunofluorescence in magenta and DAPI in cyan. Scale bar 20 μm.
Next, we evaluated the cilia beat frequency for both organs, so that, in the future, researchers can compare it with zebrafish disease models for PCD. CBF in the LRO ranges from 15 to 50 Hz was evaluated by CiliarMove [41]. This software creates a heatmap for CBF that allowed us to unequivocally detect in a very visual way different cilia in the same focal plane beating differently as depicted by the blue, green and yellow ciliary colour codes (Figure 3A’). This raised an intriguing question as to what is the function of this spatial heterogeneity in the LRO CBF? Compared to the OP, where all cilia beat around 20 Hz, in the green colour code (Figure 3B’), it is tempting to speculate that monocilia in the LRO do not coordinate their CBF for a reason that may relate to the establishment of asymmetry. In addition to the intra-LRO cilia CBF variability, we also detected an inter-embryonic variability greater in the LRO than in the OP (Figure 3C).
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Figure 3. Cilia beat frequency (CBF) evaluation for zebrafish cilia. (A,A’) Full LRO heatmap showing several monocilia at one plane beating at different frequencies (20–50 Hz). (B,B’) OP heatmap showing multiciliated cells with cilia beating at more homogeneous frequencies (around 20 Hz). CBF was measured using the software CiliarMove [41] (C) Quantification of CBF from n = 61 LROs and n = 36 OPs from embryos at 10 ss and 4 dpf larvae, respectively.

3. Different Cilia in the Zebrafish Olfactory Pit

To characterise the ultrastructural pattern of cilia in the OP, a detailed investigation was conducted by means of examining cilia from five dpf zebrafish by TEM. The quantitative analysis of three different WT animals (N = 113 cilia; values show mean ± standard deviation) showed that 60% (± 1) of cilia had a 9 + 2 arrangement with dynein arms present, and 23% (± 2) of cilia presented absent or incomplete dynein arms (most notably in the outer dynein arm (ODA)) (Figure 4 and Table 2). We further showed the heterogenicity of cilia within specific regions of the OP. When determining the localisation of the different ciliary ultrastructural types, cilia observed in the peripheral OP had dynein arms reflecting their motile function, whereas cilia in the central OP had absence of ODA and inner dynein arms (IDA). A TEM analysis allowed the quantification of the heterogenicity of cilia in a specific region of the OP. Cilia observed in the OP had a 9 + 2 motile morphology in a more peripheral area and a 9 + 2 with an ODA and IDA absence towards the centre of the OP.
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Figure 4. The zebrafish OP has two different types of cilia. (A) Schematic representation of 5 dpf zebrafish head, structures of interest are marked with arrows—olfactory pits (OP). (B) TEM low magnification image showing a cross-section across an OP of a WT zebrafish, a bowl-shaped structure containing multiciliated cells and cilia in several orientations. In the periphery of this pit (dotted boxes a and inset), cilia containing classical motile structure 9 + 2 with dynein arms (black arrowheads). In the most internal region of the OP (dotted box b and inset) we detected cilia with 9 + 2 ultrastructural arrangement without dynein arms (red arrowheads); this pattern was visible in WT embryos (n = 3). (C) Snapshot from a movie of beating OP cilia and respective heatmap by software CiliarMove (C’), highlighting a region of immotile cilia (arrowhead) that was coincident with the region detected by TEM in B. n—nucleus; cb—cilia border; OP—olfactory pit. Thin bar 6 µm, thick bar 100 nm.
Table 2. Full TEM assessment and quantification of defects in cilia from the OP in wildtype zebrafish and human healthy controls. N = 3 zebrafish OPs and n = 113 cilia were examined, values shown are mean ± standard deviation.
Disarranged Cilia (%) Dynein Arms Assessment (%)
Counted > 50 Cilia   Both Arms Present ODA Missing IDA Missing Both Arms Missing
WT zebrafish (n = 3) 14 (±8) 62 (±1) 23 (±2) 14 (±2) 14 (±2)
Human control (n = 3) 3 (±1) 93 (±12) 1 (±1) 3 (±6) 2 (±4)
The percentages of missing ODA and IDA assessed by quantitative methods [42] (as shown in Table 2) were compared using Student’s t-test in both zebrafish and human samples for a significance analysis. The comparisons between WT zebrafish OP cilia and healthy control patients concerning the presence of ODAs and IDAs were significantly different (ODA *** p ≤ 0.001; IDA * p ≤ 0.05, Table 2 and Figure 5). In healthy humans, ODAs were rarely missing, contrary to the observed findings in the central region of the zebrafish OP. Therefore, for research purposes of modelling PCD using zebrafish OP cilia, one should consider cilia from the OP periphery.
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Figure 5. Ciliary ultrastructural comparison between the LRO, OP and human respiratory cilia. Variations of the axonemal arrangement of cilia in (A) the LRO of 10 somite stages WT zebrafish embryo and (B) the OP of 5 dpf WT zebrafish larvae. (C) Example of a human respiratory cilium cross-section from the airway. Arrowheads indicate missing ODA and the dotted circle shows missing CP. Scale bar 100 nm.
Next, cross-sections from the LRO cilia were also analysed by TEM (Figure 5). A limited number of cilia were observed due to the monociliated nature of the LRO cells. The ultrastructure of these cilia, as shown previously [18], had dynein arms in WT embryos (n > 100 cilia) and some cilia show an absent or partial configuration of the CP as shown in Figure 5.
To compare the structure of the microtubule doublets of zebrafish LRO cilia against human respiratory cilia, we performed ET to generate 3D reconstructions. After assessing the cilia from the LRO by ET, some concern regarding the size of the ODA was raised, as some variation in the size of the outer dynein arms was visible, suggesting it might be smaller. To clarify, we used the software Chimera [43] (USFC, California) to measure the volume of the ODA which was normalised to the total volume of the microtubule doublet (MTD). We analysed the volume of the MTD from the LRO of four different WT zebrafish and compared them to the MTD of three human control respiratory cilia (Figure 6). No significant differences were found between the two samples, indicating that LRO cilia and human airway cilia have similar ODA volumes (student’s test, p > 0.05).
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Figure 6. Ratio between ODA and MTD volume. Sub-tomographic averaging of the MTD of WT zebrafish cilia, showing ODA and IDA, tubules (A,B) and a portion of the radial spoke. The two sample groups were not significantly different (p > 0.05, Student’s t-test). N = 3 human patients and n = 4 zebrafish LROs.

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