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1.
Doris Gove 《Ethology : formerly Zeitschrift fur Tierpsychologie》1979,51(1):58-76
Many lizards and all snakes flick their tongues. It is known that this unique behavioral pattern serves to collect airborne and substrate chemicals which give the animal information via Jacobson's Organ about the location of food, conspecifics, and possibly other environmental factors. However, a comparative topographic analysis of tongue movements in squamate reptiles is lacking, and it might shed light on the evolution of this behavior. In this study, a survey was made of the lizards and snakes which tongue-flick. Observations and films were made of 25 lizard species representing 10 families and 30 snake species representing 5 families. The information from observations and film analyses of representative species was used to hypothesize the steps of the evolution of tongue-flicking from the simple downward extensions of primitive lizards to the complex multiple oscillations of snakes. 相似文献
2.
Shou-Hsian Mao Jaang-Jiun Wang Shu-Chuan Huang Chung-Faye Chao Cheng-Chen Chen 《Journal of morphology》1991,208(3):279-292
The general histology and ultrastructure of the tongue and anterior process of the sublingual plica of four Taiwanese venomous snakes, the Chinese cobra (Naja naja atra), banded krait (Bungarus multicinctus), Taiwan habu (Trimeresurus mucrosquamatus), and bamboo snake (Trimeresurus stejnegeri stejnegeri) are described. The tongue fork exhibits a mid-dorsal invagination that broadens gradually toward its base. No mid-ventral invagination is observed. The epithelial cells on both dorsal and ventral aspects of the tongue fork have large and small microfacets, micropores and microvilli. The cell size, distribution pattern of the large microfacets, and the number of small microfacets present on both sides of the fork are essentially the same within a species, but vary among species. The function of these ultrastructures on the cell surface might be for the capture of chemical substances. The large microfacets are raised areas of the cell membrane, each with a pale granule contained within. The chemical nature of the pale granule is not yet known. The small pores surrounding the large microfacets are shallow hollows left after the release of the pale granules from the microfacets. The basic histological pattern of the tongue fork of these species is similar, being composed of a mucosal layer outside and dense musculature inside. No taste buds are discernible. The anterior processes are concave-like expansions of the anteriormost portions of the sublingual plicae. The oblique folds and micropapillae of this organ might be helpful for receiving the chemicals collected on the tongue, when the tongue makes contact with the elevated processes. The elevated processes may penetrate the ducts of Jacobson's organs to effect the final transfer. 相似文献
3.
In almost all mammals a well developed, paired and blind ending vomeronasal Organ (VNO) situated within the basement of the nasal septum, communicates with the oral cavity. This contact is established by two nasopalatine ducts, which penetrate the rostral palate close to the incisors. These ducts open orally into the sulcus which moulds the palatine papilla. In several mammals taste buds were found in the epithelium of the patatine papilla located within the nasopalatine ducts or close to their oral openings. Presumably these taste buds interact with the vomeronasal olfaction. It is likely that they are leading to a chemosensory sensation comparable to the combination of normal taste and smell. As not all mammals with a functionable VNO possess taste buds in this position, an inspection of the rostral part of the tongue which touches the palatine papilla presented an interesting situation concerning the distribution of taste buds. This region of the tongue is almost completely free of taste buds in species like Tupaia glis and Didelphis marsupialis virginiana, which have taste buds in the epithelium of their palatine papilla. In Lemur catta however, where the palatine papilla is lacking taste buds, the respective tongue part is densely covered with them. In this case it appears likely that they in a way of substitution functionally are connected with vomeronasal olfaction. 相似文献
4.
To understand the mechanisms for introducing urine or vaginal secretions into the vomeronasal organ, we used 16 mm cinematography and a freeze frame/slow motion technique to analyze the mouth and tongue movements of Brahman bulls while they examined the vulvas of restrained, estrogen-primed cows. Prior to flehmen, the mouth slowly opened, the curled tip of the tongue compressed the hard palate and the body of the tongue protruded from the mouth. The tongue maintained this form and moved forward. Once the tip of the tongue reached the incisive papilla, the body of the tongue retracted and the tip of the tongue relaxed. This tongue compression stroke (TCS) of the hard palate occurred 2 to 6 times, lasting 1 4 to 1 2 sec/stroke. Pressure changes in the vomeronasal organ are assumed to occur during and following TCSs, resulting in aspiration of any liquid in the incisive pit into the incisive and vomeronasal ducts. Such aspiration probably does not occur during flehmen because the tongue is relaxed and on the floor of the mouth. 相似文献
5.
Most snakes ingest and transport their prey via a jaw ratchetingmechanism in which the left and right upper jaw arches are advancedover the prey in an alternating, unilateral fashion. This unilateraljaw ratcheting mechanism differs greatly from the hyolingualand inertial transport mechanisms used by lizards, both of whichare characterized by bilaterally synchronous jaw movements.Given the well-corroborated phylogenetic hypothesis that snakesare derived from lizards, this suggests that major changes occurredin both the morphology and motor control of the feeding apparatusduring the early evolution of snakes. However, most previousstudies of the evolution of unilateral feeding mechanisms insnakes have focused almost exclusively on the morphology ofthe jaw apparatus because there have been very few direct observationsof feeding behavior in basal snakes. In this paper I describethe prey transport mechanisms used by representatives of twofamilies of basal snakes, Leptotyphlopidae and Typhlopidae.In Leptotyphlopidae, a mandibular raking mechanism is used,in which bilaterally synchronous flexions of the lower jaw serveto ratchet prey into and through the mouth. In Typhlopidae,a maxillary raking mechanism is used, in which asynchronousratcheting movements of the highly mobile upper jaws are usedto drag prey through the oral cavity. These findings suggestthat the unilateral feeding mechanisms that characterize themajority of living snakes were not present primitively in Serpentes,but arose subsequently to the basal divergence between Scolecophidiaand Alethinophidia. 相似文献
6.
Iseki S Ishii-Suzuki M Tsunekawa N Yamada Y Eto K Obata K 《Birth defects research. Part A, Clinical and molecular teratology》2007,79(10):688-695
BACKGROUND: Gamma-aminobutyric acid is an inhibitory neurotransmitter, synthesized by two isoforms of glutamate decarboxylase (GAD), GAD65 and -67. Unexpectedly, inactivation of GAD67 induces cleft palate in mice. Reduction of spontaneous tongue movement resulting from decreased motor nerve activity has been related to the development of cleft palate in GAD67(-/-) fetuses. In the present study, development of cleft palate was examined histologically and manipulated with culture of the maxilla and partial resection of fetal tongue. METHODS: GAD67(-/-) mice and their littermates were used. Histological examination and immunohistochemistry were performed conventionally. Organ culture of the maxilla was carried out as reported previously. Fetuses were maintained alive under anesthesia and tips of their tongues were resected. RESULTS: Elevation of palatal shelves, the second step of palate formation, was not observed in GAD67(-/-) mice. In wild-type mice, GAD67 and gamma-aminobutyric acid were not expressed in the palatal shelves, except in the medial edge epithelium. During 2 days of culture of maxillae dissected from E13.5-E14.0 GAD67(-/-) fetuses, elevation and fusion of the palatal shelves were induced. When E13.5-15.5 mutant fetuses underwent partial tongue resection, the palatal shelves became elevated within 30 min. CONCLUSIONS: These results suggest that the potential for palate formation is maintained in the palatal shelves of GAD67(-/-) fetuses, but it is obstructed by other, probably neural, factors, resulting in cleft palate. 相似文献
7.
Cylindrophis ruffus ingests prey using two distinct mechanisms. During initial phases of prey transport, lateral movements of the rear of the braincase combine with small unilateral movements of the toothed bones of each side; prey is usually constricted during this phase to permit the snake to push its head over the prey. Once transport has carried the leading part of the prey into the anterior oesophagus, Cylindrophis begins to use bilaterally synchronized movements of the jaw apparatus combined with low-amplitude, short wave-length flexions of the anterior vertebral column. Transport of prey is many times faster during the bilateral phase than during the unilateral phase.
Radiographic and cinematographic evidence indicates that the mandibular tips of Cylindrophis do not separate more than 1.5–2.0 times the resting distance between the dentary tips. Although this limits potential gape size, the intramandibular joint is highly mobile, allowing the mandibles to conform to a variety of prey shapes. Manipulations of anaesthetized and fresh, dead specimens revealed that the palatomaxillary arches are tightly attached to the ventral bones of the snout, movements of each arch being reflected in equivalent movements of the ipsilateral elements of the snout.
Cylindrophis represents a functional stage intermediate between most lizards with limited palatomaxillary kinesis and advanced snakes with considerable palatomaxillary mobility. Contrary to previous hypotheses, however, upper jaw liberation in Cylindrophis is due to liberation of the ventral snout, not to reduction of attachments to the braincase and snout. This suggests that the nose played a crucial role in the evolution of the feeding apparatus in alethinophidian snakes. 相似文献
Radiographic and cinematographic evidence indicates that the mandibular tips of Cylindrophis do not separate more than 1.5–2.0 times the resting distance between the dentary tips. Although this limits potential gape size, the intramandibular joint is highly mobile, allowing the mandibles to conform to a variety of prey shapes. Manipulations of anaesthetized and fresh, dead specimens revealed that the palatomaxillary arches are tightly attached to the ventral bones of the snout, movements of each arch being reflected in equivalent movements of the ipsilateral elements of the snout.
Cylindrophis represents a functional stage intermediate between most lizards with limited palatomaxillary kinesis and advanced snakes with considerable palatomaxillary mobility. Contrary to previous hypotheses, however, upper jaw liberation in Cylindrophis is due to liberation of the ventral snout, not to reduction of attachments to the braincase and snout. This suggests that the nose played a crucial role in the evolution of the feeding apparatus in alethinophidian snakes. 相似文献
8.
Marie-Céline Buchy Eberhard Frey Steven W. Salisbury 《Lethaia: An International Journal of Palaeontology and Stratigraphy》2006,39(4):289-303
In the late 19th Century, the choanae (or internal nares) of the Plesiosauria were identified as a pair of palatal openings located rostral to the external nares, implying a rostrally directed respiratory duct and air path inside the rostrum. Despite obvious functional shortcomings, this idea was firmly established in the scientific literature by the first decade of the 20th Century. The functional consequences of this morphology were only re-examined by the end of the 20th Century, leading to the conclusion that the choanae were not involved in respiration but instead in underwater olfaction, the animals supposedly breathing with the mouth agape. Re-evaluation of the palatal and internal cranial anatomy of the Plesiosauria reveals that the traditional identification of the choanae as a pair of fenestrae situated rostral to the external nares appears erroneous. These openings more likely represent the bony apertures of ducts that lead to internal salt glands situated inside the maxillary rostrum. The 'real' functional choanae (or caudal interpterygoid vacuities), are situated at the caudal end of the bony palate between the sub-temporal fossae, as was suggested in the mid-19th Century. The existence of a functional secondary palate in the Plesiosauria is therefore strongly supported, and the anatomical, physiological, and evolutionary implications of such a structure are discussed. 相似文献
9.
Alejandro Rico-Guevara Tai-Hsi Fan Margaret A. Rubega 《Proceedings. Biological sciences / The Royal Society》2015,282(1813)
Pumping is a vital natural process, imitated by humans for thousands of years. We demonstrate that a hitherto undocumented mechanism of fluid transport pumps nectar onto the hummingbird tongue. Using high-speed cameras, we filmed the tongue–fluid interaction in 18 hummingbird species, from seven of the nine main hummingbird clades. During the offloading of the nectar inside the bill, hummingbirds compress their tongues upon extrusion; the compressed tongue remains flattened until it contacts the nectar. After contact with the nectar surface, the tongue reshapes filling entirely with nectar; we did not observe the formation of menisci required for the operation of capillarity during this process. We show that the tongue works as an elastic micropump; fluid at the tip is driven into the tongue''s grooves by forces resulting from re-expansion of a collapsed section. This work falsifies the long-standing idea that capillarity is an important force filling hummingbird tongue grooves during nectar feeding. The expansive filling mechanism we report in this paper recruits elastic recovery properties of the groove walls to load nectar into the tongue an order of magnitude faster than capillarity could. Such fast filling allows hummingbirds to extract nectar at higher rates than predicted by capillarity-based foraging models, in agreement with their fast licking rates. 相似文献
10.
C Rommel 《Gegenbaurs morphologisches Jahrbuch》1981,127(3):421-451
1. In Homo and the great apes (Pongidae) there occurs, besides the plica sublingualis a plica fimbriata at the ventral surface of the tongue. This duplicature of the mucosa does not occur in the Hylobytidae and in the other primates. 2. Some taste buds could be found in the epithelium of the plica sublingualis of the Pongidae. 3. There are many taste buds in the epithelium of the plica fimbriata of the Pongidae. On this sublingual structure there were counted 1776 taste buds in Pongo, 592 in Gorilla and 280 in Pan. A few taste buds could also be found on the plica fimbriata of a human newborn. 4. A glandula apicis linguae occurs in Homo, Pan, Gorilla and Pongo. 5. The fresh saliva of the glandula apicis linguae and the saliva on the floor of the mouth can be tested by the taste buds in the epithelium of the plica fimbriata, of papillae lenticulares and of areae gustatoriae at the ventral surface of the tongue. 6. It might be the function of the sublingual taste buds to taste the fresh saliva as a gradient for the central nervous comparison with the taste of the saliva on the dorsal surface of the tongue. 7. Because of the complete absence of a sublingua in the Platyrrhini and in the Cercopithecinae it is unlikely that the plica fimbriata of Homo and the great apes can be interpreted as a homalogon of the sublingua in the prosimians. 8. Because of the absence of a sublingua in other ordines of the Mammalia (Insectivora, Carnivora, Rodentia, Chiroptera, Ungulata) it is unlikely as well that the sublingua in the prosimians can be interpreted as a homologon of the tongues of the lower vertebrates. The sublingual structures occuring in the Marsupialia have to be investigated. 9. Because of these reasons the new development of the sublingua in the prosimians and the plica fimbriata in the Hominoidea, in complete independence from one another, seems to be a better explanation of the 2 structures and less contradictionary to anatomical and phylogenetic arguments. The different function of both structures in the recent primates gives a hint for the possible reason for their development during the process of evolution. 相似文献
11.
The fluid mechanics of bolus ejection from the oral cavity 总被引:1,自引:0,他引:1
The squeezing action of the tongue against the palate provides driving forces to propel swallowed material out of the mouth and through the pharynx. Transport in respose to these driving forces, however, is dependent on the material properties of the swallowed bolus. Given the complex geometry of the oral cavity and the unsteady nature of this process, the mechanics governing the oral phase of swallowing are not well understood. In the current work, the squeezing flow between two approaching parallel plates is used as a simplified mathematical model to study the fluid mechanics of bolus ejection from the oral cavity. Driving forces generated by the contraction of intrinsic and extrinsic lingual muscles are modeled as a spatially uniform pressure applied to the tongue. Approximating the tongue as a rigid body, the motion of tongue and fluid are then computed simultaneously as a function of time. Bolus ejection is parameterized by the time taken to clear half the bolus from the oral cavity, t1/2. We find that t1/2 increases with increased viscosity and density and decreases with increased applied pressure. In addition, for low viscosity boluses (μapproximately 1000 cP), viscosity dominates. A transition region between these two regimes is found in which both properties affect the solution characteristics. The relationship of these results to the assessment and treatment of swallowing disorders is discussed. 相似文献
12.
Water drinking in the mallard is accomplished by a fine-tuned set of movements of upper and lower jaw and of the tongue. During immersion of the tips of the bill, the oral cavity is formed into smaller volumes containing water and into connecting tubes. Two mechanisms serve the water transport: (1) lingual and jaw movements press water from the water-containing spaces into the tubes; (2) a quantitative simulation of the shape of the oral cavity during immersion shows that the two tubes are so narrow that capillary action also contributes to water transport. Thereafter, the tips of the bill are raised until they point upward. In this “tip-up” position, water flows into the esophagus because of gravity. We conclude that, in addition to normal tip-up drinking observed in almost all Passeriformes and Galliformes, a second type of tip-up drinking may be distinguished in Anseriformes. The integration of the drinking mechanism, keeping the water inside the mouth, and the straining mechanism, expelling the water along the beak rims, is effected by specific actions of the elaborate lingual apparatus. 相似文献
13.
This study is concerned with reconciling theoretical modelling of the fluid flow in the airway surface liquid with experimental
visualisation of tracer transport in human airway epithelial cultures. The airways are covered by a dense mat of cilia of
length ∼ 6 μm beating in a watery periciliary liquid (PCL). Above this there is a layer of viscoelastic mucus which traps
inhaled pathogens. Cilia propel mucus along the airway towards the trachea and mouth. Theoretical analyses of the beat cycle
smithd, fulb predict small transport of PCL compared with mucus, based on the assumption that the epithelium is impermeable
to fluid. However, an experimental study coord indicates nearly equal transport of PCL and mucus. Building on existing understanding
of steady advection-diffusion in the ASL (Blake and Gaffney, 2001; Mitran,2004) numerical simulation of an advection-diffusion
model of tracer transport is used to test several proposed flow profiles and to test the importance of oscillatory shearing
caused by the beating cilia. A mechanically derived oscillatory flow with very low mean transport of PCL results in relatively
little ‘smearing’ of the tracer pulses. Other effects such as mixing between the PCL and mucus, and significant transport
in the upper part of the PCL above the cilia tips are tested and result in still closer transport, with separation between
the tracer pulses in the two layers being less than 9%. Furthermore, experimental results may be replicated to a very high
degree of accuracy if mean transport of PCL is only 50% of mucus transport, significantly less than the mean PCL transport
first inferred on the basis of experimental results. 相似文献
14.
Invasion of black-tailed deer (Odocoileus hemionus columbianus) by larvae of the nose bot flies (Cephenemyia apicata and C. jellisoni) was investigated by obssrving their expulsion by larviparous females and their subsequent activity on the host. The drying uterine fluid encasing larvae ( = larval packet) delays desiccation, ensures adhesion to hairs, and immediately dissolves upon contact with saliva. Contrary to the widely accepted nasal mode of invasion, larvae placed on muzzles of deer crawl ventrally toward the upper lip, enter the mouth, and then crawl caudally along the hard palate or tongue toward the throat. Hair located between the muzzle and nostril prevents larvae from entering the nostrils. A natural per os mode of invasion, heretofore unrecognized, is proposed. This is initiated by: (1) females depositing larval packets on the muzzle of deer, or (2) deer licking larval packets from contaminated areas around the muzzle. The positive thermotropism of larvae is compatible with such a per os mode of entry into the host. 相似文献
15.
Bryan G. Fry Eivind A.B. Undheim Syed A. Ali Timothy N. W. Jackson Jordan Debono Holger Scheib Tim Ruder David Morgenstern Luke Cadwallader Darryl Whitehead Rob Nabuurs Louise van der Weerd Nicolas Vidal Kim Roelants Iwan Hendrikx Sandy Pineda Gonzalez Ivan Koludarov Alun Jones Glenn F. King Agostinho Antunes Kartik Sunagar 《Molecular & cellular proteomics : MCP》2013,12(7):1881-1899
16.
Summary The scincid lizardTiliqua rugosa possesses a large external nasal gland which is located intraconchally. Highly ramified tubules, imbedded primarily in the periphery of the gland, unite to form collecting ducts which empty into a short excretory canal. The diameter of the tubules increases progressively from 30. at the distal extremity of the gland to over 200 at the level of the collecting ducts. The intraglandular portion of the excretory canal is often dilated to form an ampulla. The thickness of the epithelium increases from 12 at the level of the tubules to 25–30 in the excretory canal.The excretory canal is lined with an epidermal epithelium close to the point where it enters the vestibule. In all the rest of the gland the tubules are lined with two cell types: large, typical muco-serous cells and striated cells. At the distal end of the tubules the striated cells are narrow and poorly differentiated and alternate more-or-less regularly with the muco-serous cells. The relative proportion of these striated cells increases progressively, as does their size, as one moves proximally down the tubule. In the gland as a whole the striated cells are approximately twice as numerous as the muco-serous cells but, due to their smaller size, they occupy less than one third of the tubular volume.Electron microscopy of the striated cells ofTiliqua rugosa revealed the presence of extensive lateral interdigitations and expansions of the basal cytoplasmic membrane, anatomical specialisations which are normally indicative of active salt transport. These modifications are less marked however than in the external nasal glands of the lizardsLacerta muralis andVaranus griseus, which do not appear to function as salt glands. In addition there are few mitochondria present, although they are of large size. The combination of these ultrastructural features, plus the fact that the striated cells are intermixed with muco-serous cells in the tubules, makes it most unlikely that the external nasal gland ofTiliqua rugosa is capable of elaborating an hyperosmotic fluid. What is more, this has never been conclusively demonstrated in this species in physiological studies.The progressive specialisation of the striated cells from the distal to the proximal section of the tubules poses the problem of the origin and differentiation of this cell type.A review of results obtained from the study ofTiliqua rugosa and other species of lizards shows that the nature of the relationship between structure and function of the external nasal gland is far from clear. The existence of salt glands, capable of excreting hyperosmotic solutions, is invariably linked with the presence in the gland of well-developed striated segments composed almost entirely of cells possessing extensive interdigitations of the lateral membranes. Amongst terrestrial lizards, nasal salt glands are usually found in herbivorous species and they are primarily adapted to the extrarenal excretion of potassium ions. The problem for carnivorous species is more often that of an excess of sodium rather than potassium ions and with the possible exceptionAcanthodactylus species, functional nasal salt glands have not been demonstrated in terrestrial carnivores, despite the presence in some cases of well-developed striated segments in the gland having a similar structure to those found in herbivores. In humid regions, carnivorous lizards probably never require extrarenal excretory mechanism and in arid regions their survival is assured by their capacity to tolerate hypernatraemia when confronted with excessive salt loads. Salt glands capable of eliminating sodium ions to any extent have only been described in two littoral species, an herbivorous iguanid and a carnivorous varanid. Unfortunately the structure of their respective nasal glands has not yet been described and their further study would be desirable. 相似文献
17.
Bryony Braschi Heymut Omran George B. Witman Gregory J. Pazour K. Kevin Pfister Elspeth A. Bruford Stephen M. King 《The Journal of cell biology》2022,221(2)
Dyneins are highly complex, multicomponent, microtubule-based molecular motors. These enzymes are responsible for numerous motile behaviors in cytoplasm, mediate retrograde intraflagellar transport (IFT), and power ciliary and flagellar motility. Variants in multiple genes encoding dyneins, outer dynein arm (ODA) docking complex subunits, and cytoplasmic factors involved in axonemal dynein preassembly (DNAAFs) are associated with human ciliopathies and are of clinical interest. Therefore, clear communication within this field is particularly important. Standardizing gene nomenclature, and basing it on orthology where possible, facilitates discussion and genetic comparison across species. Here, we discuss how the human gene nomenclature for dyneins, ODA docking complex subunits, and DNAAFs has been updated to be more functionally informative and consistent with that of the unicellular green alga Chlamydomonas reinhardtii, a key model organism for studying dyneins and ciliary function. We also detail additional nomenclature updates for vertebrate-specific genes that encode dynein chains and other proteins involved in dynein complex assembly.IntroductionDynein family motor proteins form multiple different dynein complexes in mammals, with important roles in a wide range of cellular functions (King, 2017; Osinka et al., 2019; Roberts, 2018). Dyneins can be broadly classified into two groups: cytoplasmic and axonemal. Dynein complexes “walk” toward the minus ends of microtubules; while doing so, they can transport a variety of cargoes within cells (Trokter et al., 2012). The motor activity of these complexes allows them to play key roles in enabling motility of whole cells, generating fluid flow across cell surfaces, and transporting organelles and other components within the cytoplasm.Dynein subunits are classified by mass into four categories: heavy (∼520 kD), intermediate (∼70 –140 kD), light intermediate (∼53–59 kD), and light (∼10–30 kD) chains (Pfister et al., 2006). The heavy and intermediate chains are specific to certain dynein complexes, while the light chains may be components of both cytoplasmic and axonemal dynein machinery, and in some cases, nondynein complexes. The light intermediate chains are present only in the cytoplasmic dynein class.Dynein-based movement is powered by the ATP-driven dynein heavy chain subunits (Schmidt and Carter, 2018). 15 genes in the human genome encode dynein heavy chains: 1 for each of the 2 cytoplasmic dynein complexes and 13 that encode heavy chain components of the various axonemal dynein complexes. A dynein-related gene, DNHD1 (dynein heavy chain domain 1) has been referred to as a “ghost gene”: it may be a remnant of an earlier duplication that has not decayed at a normal rate, as a truncated version might poison cytoplasmic dynein heavy chain dimerization and thus be lethal (Gibbons, 2018; Schmidt and Carter, 2018). DNHD1 is currently classified as an “orphan” dynein heavy chain-encoding gene (Kollmar, 2016) but may be in the process of becoming a pseudogene (Wickstead, 2018).Cytoplasmic dyneinsDynein 1 complexThe cytoplasmic dynein 1 complex (Table S1) is present throughout eukaryotes, with some notable exceptions such as green plants and red algae (Wickstead and Gull, 2007). It is involved in a wide variety of intracellular transport activities, transporting cargoes including chromosomes, mRNA, and protein complexes (Reck-Peterson et al., 2018). The dynein 1 complex also acts in cell division, helping to form and orient the mitotic spindle (Torisawa and Kimura, 2020), establish cell polarity (Lu and Gelfand, 2017), and position organelles (Allan, 2014; Oyarzún et al., 2019; Palmer et al., 2009).A dimer of DYNC1H1-encoded heavy chains forms the core of the cytoplasmic dynein 1 complex (Fig. 1 a) and acts as its ATPase motor (Palmer et al., 2009; Pfister et al., 2005). Each heavy chain contains six AAA+ domains, an antiparallel coiled-coil region with a microtubule-binding domain at its tip, and a C-terminal domain (Bhabha et al., 2016; Carter, 2013; Reck-Peterson et al., 2018; Roberts et al., 2013). Immediately N-terminal of the AAA1 domain is a linker that traverses the plane of the AAA ring and changes conformation during the ATPase cycle to drive motor activity. AAA1 exhibits ATP hydrolytic activity, acting as an ATPase and powering the dynein motor complex (Silvanovich et al., 2003), while nucleotide binding at several other AAA domains appears to modify how conformational change propagates through the AAA ring and affects microtubule-binding activity. The coordinated activity of both heavy chains within the dynein complex is required for processivity (Reck-Peterson et al., 2006).Open in a separate windowFigure 1.Cytoplasmic dynein complexes. (a) The cytoplasmic dynein 1 complex. The DYNC1H1 protein heavy chains have large globular heads at the C-termini that are composed of a ring of six AAA+ domains. The microtubule-binding domains are located at the tips of antiparallel coiled coils that derive from AAA4. The linker/N-terminal domains connect the AAA rings and the intermediate and light chains. (b) The cytoplasmic dynein 2 complex. The DYNC2H1 protein heavy chains power retrograde IFT and have the same general domain organization as DYNC1H1. However, the tails of the two heavy chains fold differently due to an asymmetry imposed by the two different intermediate chains: one is straight while the other forms a zigzag shape and interacts with the IFT-B train (Toropova et al., 2019). The linker/N-terminal domain connects the AAA ring and the intermediate and light chains. *It remains unknown whether the DYNLT2B protein forms a homodimer or a heterodimer with another Tctex-type light chain. (c) Schematic showing the interaction between the dynein 1 and dynactin complexes. The adapter molecule affects the type of cargo bound; in this figure, the hook microtubule tethering protein 3 (HOOK3)–encoded protein is acting as a cargo adapter.The intermediate chains of metazoan dynein 1 connect it to another multi-subunit complex known as dynactin (Loening et al., 2020). Dynactin is built around a filament of the protein encoded by ACTR1A (actin-related protein 1A). It activates dynein and regulates its binding to vesicles and organelles to be transported (Ketcham and Schroer, 2018). A coiled coil–containing cargo adaptor protein is required for dynein 1 activation (Fig. 1 c). A single adaptor protein sandwiches between dynactin and dynein, where it interacts with the dynein heavy chain tails and the light intermediate chain and along the length of the dynactin complex (Gonçalves et al., 2019; Reck-Peterson et al., 2018). There are currently ≥12 known cargo adaptor proteins, which are encoded by HOOK1, HOOK2, HOOK3, BICD2, BICDL1, BICDL2, RAB11FIP3, RASEF, CRACR2A, NIN, NINL, and SPDL1 (Barisic et al., 2010; Casenghi et al., 2005; Dona et al., 2015; Gonçalves et al., 2019; Horgan et al., 2010; Lee et al., 2018; Loening et al., 2020; Olenick et al., 2016; Vallee et al., 2021; Wang et al., 2019). Other protein cofactors may also be required for dynein recruitment to their cargoes. For example, the protein encoded by PAFAH1B1 (platelet activating factor acetylhydrolase 1b regulatory subunit 1; HUGO Gene Nomenclature Committee [HGNC] ID: 8574), also published using the alias LIS1 (lissencephaly 1) is required along with dynactin and BICD2 for dynein 1 to traffic many cargoes, such as nuclei, along microtubules (Faulkner et al., 2000; Splinter et al., 2012). LIS1 has most recently been suggested to stabilize the “open” conformation of cytoplasmic dynein 1 such that the heavy chains are able to undergo a mechanochemical cycle and cannot adopt the autoinhibited or “closed” state where movement of key mechanical elements is abrogated by interactions between heavy chains (Markus et al., 2020).The dynein light chains can be divided into three subfamilies: the t-complex associated (Tctex)–type family (encoded by DYNLT1, DYNLT2, DYNLT2B, DYNLT3, DYNLT4, and DYNLT5), the LC8-type family (encoded by DYNLL1, DYNLL2, and DNAL4), and the roadblock-type family (encoded by DYNLRB1 and DYNLRB2; Bowman et al., 1999; King et al., 1998; King et al., 1996; King and Patel-King, 1995). Most of these protein chains can be found in both cytoplasmic dynein complexes: the exceptions are DYNLT2, which is an axonemal dynein subunit; DYNLT2B, which is found in the dynein 2 complex and the I1/f inner dynein arm (IDA); DYNLT4 and DYNLT5, which are not well characterized; and DNAL4, which is present only in outer dynein arms (ODAs).Several proteins originally identified as dynein light chains are also found in numerous multimeric complexes unrelated to dyneins and appear to act as general dimerization engines or hubs (Williams et al., 2018). The LC8-type light chains (DYNLL1 and DYNLL2) are present in many enzymes including myosin V (Benashski et al., 1997; Espindola et al., 2000) and neuronal nitric oxide synthase (Jaffrey and Snyder, 1996). They also play a role in regulating apoptosis via an interaction with the BCL2 family protein encoded by BCL2L11 (Puthalakath et al., 1999). The DYNLT1 protein has reported roles in actin remodeling and neurite outgrowth (Chuang et al., 2005) and hypocretin signaling (Duguay et al., 2011). DYNLRB1 interacts with Rab6 family member proteins in the Golgi apparatus (Wanschers et al., 2008), and both roadblock-type dynein light chains are reportedly involved in a TGFβ signaling pathway (Jin et al., 2009).Dynein 2 complexCilia are highly complex microtubule-based organelles that extend from the cell surface and can be classified as either primary (or nonmotile) or motile (Satir and Christensen, 2008). Most eukaryotic cells, excluding blood cells and those actively dividing, have an associated primary or nonmotile cilium. These act as sensory organelles, detecting a broad range of signaling molecules (Kopinke et al., 2021; Mykytyn and Askwith, 2017; Saternos et al., 2020).The dynein 2 complex (also known as the intraflagellar transport [IFT] dynein or cytoplasmic dynein 1b in Chlamydomonas reinhardtii; Table S2) is found only in cells with associated cilia or flagella, where it locates within and around the base of these structures (Höök and Vallee, 2006). IFT trains are multiprotein complexes required for the assembly and function of cilia and flagella in eukaryotes (Dutcher, 2019; Wingfield et al., 2017). The anterograde IFT motor complex kinesin 2 moves IFT trains and associated cargoes plus the dynein 2 complex along microtubules, from the base to the tip of a cilium or flagellum (Toropova et al., 2019; Vuolo et al., 2020). The retrograde IFT motor complex dynein 2 transports IFT trains and associated factors from the tip back to the base (Hou and Witman, 2015; Pazour et al., 1998). The dynein 2 complex is required for the assembly of cilia and flagella (Pazour et al., 1999; Pfister et al., 2006) and also has key roles in ciliary signaling functions (Vuolo et al., 2020).The core of dynein 2 is composed of a dimer of two DYNC2H1-encoded heavy chains (Fig. 1 b). The tails of these identical heavy chains are directed into two different conformations by the other subunits in the complex (Toropova et al., 2019). Each heavy chain is stabilized by its interaction with a DYNC2LI1-encoded protein subunit. The C-terminal helix of one of these light intermediate subunits associates with a DYNC2I1 (previously WDR60)-encoded protein with a DYNLRB-encoded subunit, to enforce a distinct conformation on one heavy chain (Toropova et al., 2019; Vuolo et al., 2020).The DYNC2I1- and DYNC2I2-encoded intermediate chains bind the heavy chains via their C-terminal β-propeller domains. The N-terminal regions of these intermediate chains are dimerized by three DYNLL1/2 dimers and one of each of the other light chain dimers: DYNLT1/3, DYNLRB1/2, and DYNLT2B (Toropova et al., 2019). The DYNLT2B-encoded light chain is a unique accessory component of the dynein 2 complex. Whether it forms a homodimer or heterodimer with another light chain remains to be confirmed, although there is evidence to suggest that, unlike the other light chains, the DYNLT2B subunit may be monomeric (DiBella et al., 2001). Recent structural studies of Tetrahymena ODAs have revealed a Tctex-family heterodimer (Rao et al., 2021).Axonemal dyneinsMotile cilia (sometimes termed flagella when they occur singly or in small numbers on a cell) are more restricted to certain cell types. Their movement enables sperm to swim (Linck et al., 2016), respiratory cilia on epithelial cells to sweep away mucus containing trapped pathogens (Hansson, 2019), and oviduct epithelial cells to waft an ovum along a fallopian tube toward the uterus (Spassky and Meunier, 2017). Multiciliated cells in the brain help move the cerebrospinal fluid and also influence neuronal migration (Brooks and Wallingford, 2014). In the male reproductive tract, the epithelial cells of the efferent ducts are densely covered with multiple motile cilia necessary for the transport of sperm cells (Aprea et al., 2021a). Motility of nodal cilia in the embryonic left–right organizer is necessary for the determination of correct left–right body asymmetry (Nonaka et al., 1998).An axoneme is the microtubule superstructure core of the cilium and contains many tightly associated components. A motile cilium has a highly conserved “9 + 2” structure: 9 microtubule doublets that surround a central pair of 2 microtubule singlets (the “central apparatus”; Fig. 2). Axonemal dyneins are the motor complexes that drive a sliding motion between ciliary doublet microtubules, enabling movement. Motile cilia have IDAs and ODAs and radial spokes that are thought to be involved in signal transduction between the central pair and the outer microtubule ring (Ishikawa, 2017). Nonmotile cilia have only the outer doublet ring and have a 9 + 0 microtubule arrangement, although the number of outer doublets decreases and their arrangement changes beyond the proximal part of the cilium (Kiesel et al., 2020).Open in a separate windowFigure 2.Axonemal dynein complexes. (a) Axonemal ODA. The blue text denotes subunits found in ODA complexes in respiratory cilia, and red text denotes subunits found in ODA complexes in sperm flagella. (b) Axonemal inner arm I1/f complex subunits (IDA). (c) Monomeric IDAs. Each inner arm species is constructed around a distinct monomeric heavy chain associated with an actin monomer and either DNALI1 or centrin; species d contains two additional components. In most cases, the precise equivalence between the human and C. reinhardtii monomeric heavy chain species is uncertain.The ODAs (Table S3 and Fig. 2 a) and IDAs (Table S4 and Table S5, and Fig. 2, b and c) in motile cilia are arranged in two rows with a complex 96-nm repeat organization. They are permanently attached to the A-tubule of one outer doublet microtubule (see Fig. 3, a and b) and transiently interact in an ATP-dependent manner with the B-tubule of the adjacent doublet to generate a sliding force (King, 2017). IDAs with a single heavy chain are termed monomeric (Table S4), while the I1/f IDA (Table S5) is dimeric, with two nonidentical heavy chains. These different types of dyneins vary in terms of their enzymatic and motor properties, likely reflecting their precise roles in the generation of ciliary motility (King, 2017).Open in a separate windowFigure 3.Organization of a mammalian motile cilium. (a) The diagram illustrates the general 9 + 2 microtubule arrangement within the ciliary axoneme. The inner and outer rows of dynein arms generate the force required for ciliary beating. The N-DRC complex is a key regulatory structure that interconnects the doublet microtubules. The radial spokes regulate the beat of cilia by transducing signals between the doublets and the central microtubule pair. (b) Tomographic image of an averaged 96-nm repeat for a single human ciliary doublet microtubule, revealing the microtubule-associated dynein arms, N-DRC, and radial spoke. The scale bar represents 25 nm. This image was generated by Jason Schrad (Nicastro laboratory) using data from Lin et al. (2014). (c and d) Cross-section (c) and longitudinal (d) views of the 48-nm repeat organization of a bovine doublet microtubule. The components of the ODA-DC are individually colored and indicated. This ribbon diagram was generated with the PyMol molecular graphics system (Schrödinger) using Protein Data Bank accession no. 7RRO (Gui et al., 2021).ODA docking complex (ODA-DC)The correct functioning of cilia and flagella in most eukaryotes is dependent on the ODA chains attaching to the outer doublet microtubules at 24-nm intervals (Dean and Mitchell, 2015; King, 2017). The ODA-DC facilitates binding and may also play a role in regulating the activity of the ODAs (Takada et al., 2002). The ODA-DC in C. reinhardtii consists of three protein subunits, encoded by DCC1 (DC1), DCC2 (DC2), and DLE3 (DC3). In mammals, it consists of five protein subunits (Gui et al., 2021) encoded by five genes, now named ODAD1, ODAD2, ODAD3, ODAD4, and CLXN (calaxin; Fig. 2 a and Fig. 3, b and c). CLXN (previously EFCAB1) has been assigned the alias symbol ODAD5, and authors may refer to it as such in publications if they wish, referencing the approved gene symbol at least once to aid data retrieval. Only ODAD1 and ODAD3 have orthologues in C. reinhardtii (DCC2 and DCC1, respectively).Dynein axonemal assembly factors (DNAAFs)Genes encoding proteins that act as axonemal dynein assembly factors are named using the root symbol DNAAF. These proteins play an important role in the preassembly of IDAs and ODAs in the cytoplasm before their transport to cilia (Fabczak and Osinka, 2019; King, 2021).Historically, the DNAAF root has been used only for proteins directly involved in the preassembly of axonemal dynein arms in the cytoplasm. We wrote to authors who have published on the genes that we are reporting in this publication as newly updated DNAAFs (see their symbols in bold in Table S6) and discussed this issue with our specialist advisors for this gene group (https://www.genenames.org/data/genegroup/#!/group/1627). This effort resulted in an agreement to use the term “DNAAF” more broadly. Therefore, a DNAAF symbol can now also be assigned to genes encoding proteins that play a role in trafficking dynein arms from the cytoplasm to cilia.Association with human phenotypesHumans have four described cilia types, and defects in all types are associated with various diseases: motile 9 + 2 cilia (e.g., respiratory cilia, ependymal cilia, sperm flagella); motile 9 + 0 cilia (e.g., nodal cilia); nonmotile 9 + 2 cilia (e.g., the kinocilium of hair cells and the proximal region of olfactory cilia); and nonmotile 9 + 0 cilia (e.g., renal monocilia and the connecting cilia of photoreceptor cells). Cilia are located on almost all polarized cell types of the human body; therefore, cilia-related disorders (ciliopathies) affect many organ systems (Fliegauf et al., 2007). Genetic mutations that impair cilia and/or flagella beating cause a heterogeneous group of rare disorders referred to as motile ciliopathies (Wallmeier et al., 2020). The pathogenic mechanisms, clinical symptoms, and severity of the diseases depend on the specific affected genes and the tissues in which they are expressed. Defects in ependymal cilia can result in hydrocephalus. Reduced fertility can be due to defective cilia in the fallopian tubes or the efferent ducts as well as sperm flagella. The malfunction of motile monocilia on the left–right organizer during early embryonic development can lead to laterality defects such as situs inversus and heterotaxy. Severe impairment of mucociliary clearance in the respiratory tract leads to chronic bronchial problems. Primary ciliary dyskinesia (PCD), which can present with a variety of these features, is the most common motile ciliopathy.The genetic disorder PCD is heterogeneous and has been linked to variants in genes encoding dyneins, axonemal dynein assembly factors, and ODA-DC subunits (Wallmeier et al., 2020). PCD-associated phenotypes include chronic respiratory problems, recurrent middle ear infections, male infertility, and subfertility in females (Leigh et al., 2019). Roughly 50% of PCD patients are diagnosed with Kartagener syndrome, a subtype defined by a triad of symptoms: chronic sinusitis, bronchiectasis, and situs inversus, where the positions of major body organs are reversed (Zariwala et al., 2011). Situs inversus totalis is observed when all thoracic and abdominal viscera are reversed; individuals with situs inversus or situs ambiguus show more variable organ positioning (seen in ≥6% of PCD cases; Kennedy et al., 2007; Sempou and Khokha, 2019).Table 1.Human phenotypes associated with variants of genes encoding dyneins and dynein-associated proteins
Open in a separate windowaNote that for some of these phenotypes, there are several variants with varying degrees of severity, and different genes may be associated with different types of these genetic conditions.Mutations in DNAH5 encoding an axonemal ODA heavy chain are the most common genetic defect observed in PCD (Hornef et al., 2006). DNAH5 mutations result in dysmotility of respiratory as well as nodal cilia (Olbrich et al., 2002). Defective nodal cilia motility during early embryogenesis caused by mutations in genes encoding components essential for ciliary motility (e.g., due to DNAH5 mutations) result in situs inversus or situs ambiguus in approximately half of affected individuals due to the randomization of their left–right body asymmetry. Consistently, mice deficient for DNAH5 show immotility of respiratory cilia and embryonic nodal monocilia and exhibit ODA defects in both cilia types (Nöthe-Menchen et al., 2019). DNAH5 mutations also result in ODA defects and dysmotility of ependymal cilia (Ibañez-Tallon et al., 2004). DNAH5-deficient mice develop hydrocephalus during early postnatal life because the flow of cerebrospinal fluid around the brain is obstructed by the abnormal closure of the aqueduct of Sylvii connecting the third and fourth brain ventricles. Possibly due to the larger human brain size, the active propulsion of cerebrospinal fluid along the narrow passages of the ventricular system is not essential in most individuals with PCD; however, they still carry a slightly increased risk of developing hydrocephalus. This suggests that the non–motility-related functions of ependymal cilia might also be important (Wallmeier et al., 2020).All known motile cilia types with DNAH5 loss-of-function mutations display aberrant motility, with the exception of sperm flagella. This is because the paralogous protein DNAH8 is present in sperm and exhibits functional overlap. The male reproductive tracts of mice deficient for DNAH5 have immotile efferent duct cilia, which results in severe stasis of sperm cell transport; this is due to disruption of the ODA composition. In human individuals with loss-of-function DNAH5 mutations, reduced sperm count in the ejaculate (oligozoospermia) and dilatations of the epididymal head were observed, consistent with DNAH5 in efferent duct cilia having an important role in sperm cell transport (Aprea et al., 2021a).In females, the ODA composition of cilia in the Fallopian tube resembles that of respiratory cilia, with the ODA DNAH5 (dynein axonemal heavy chain 5) and DNAI1 (dynein axonemal intermediate chain 1) both being present (Raidt et al., 2015). The coordinated beating of the Fallopian tube ciliated cells produces a fluid flow from the distal site of the Fallopian tubes (ovaries), which transports the egg to the proximal end of the reproductive tract (uterine cavity; Lyons et al., 2006). Interestingly, some females with defective DNAH5 and DNAI1 are still able to conceive children. Thus, the motility of Fallopian tube cilia may not be essential for gamete transport, as Fallopian tube muscle contractions might aid in transporting the egg to the uterine cavity.Mutations in genes encoding DNAAFs cause variable degrees of absence of ODAs and IDAs in respiratory cilia and sperm flagella (Aprea et al., 2021b), indicating that the process of cytoplasmic assembly of dynein arms is critical in both cell types. DNAAF mutant individuals consistently exhibit severely hampered motility of both sperm flagella and respiratory cilia. The sperm flagella of some DNNAF mutant males have shortened flagella axonemes, indicating that their length is also influenced by DNAAF function during dynein arm assembly.Most defects of DNAAFs and axonemal dynein components affect motility of cilia and sperm flagella, contributing to motile ciliopathies (Leigh et al., 2019; Reiter and Leroux, 2017; Wallmeier et al., 2020). However, mutations in genes encoding cytoplasmic dynein subunits can affect the function of both motile and nonmotile cilia, as well other cellular processes. Thus, the clinical phenotype can vary enormously depending on the cell types that are affected. A variant of DYNC1H1 has been associated with a particular form of the ciliopathy SMALED (spinal muscular atrophy lower extremity dominant). This form of the condition mainly affects the lower limbs, causing progressive muscle weakness (Das et al., 2018). A different point mutation in DYNC1H1, also within the tail domain of the heavy chain protein, has been associated with the related neuropathy Charcot Marie Tooth disease. Dysfunction of the dynein heavy chains encoded by DYNC1H1 may also adversely affect maintenance of the morphology of mitochondria and may contribute to disease pathology (Eschbach et al., 2013).Variants of several genes encoding dynein 2 subunits (Dagoneau et al., 2009). If retrograde IFT trafficking of cargoes from the tip to the base of the cilium is compromised, then so is hedgehog (Hh) signaling in the developing embryo, and the resulting incorrect embryonic patterning can produce a range of phenotypes (Goetz and Anderson, 2010). Patients with these conditions have skeletal abnormalities including a narrow thorax, short ribs, and bony spurs in a three-pronged formation observed at the hip joint; they may also display polydactyly.Variants of some of the genes encoding dynein 2 subunits have also been linked to phenotypes affecting vision. The outer segment of photoreceptors is a modified cilium, and a constant turnover of outer segment constituents is required; IFT is key to this process. Four variants in DYNC2H1 in human are associated with nonsyndromic retinal degeneration (Vig et al., 2020). Some of these variants are suggested to affect the ciliary transport of the protein encoded by IFT88, an IFT component that is essential for the assembly and maintenance of vertebrate photoreceptors (Pazour et al., 2002).Standardizing gene nomenclatureThe HGNC (https://www.genenames.org) is the international authority assigning standardized nomenclature to human genes, and hence facilitating communication between researchers. We aim to assign unique, informative symbols and names to human genes that can be used in all domains, and across major biological and clinical databases and publications. Our sister project, the Vertebrate Gene Nomenclature Committee (VGNC; https://vertebrate.genenames.org), names genes across selected vertebrates in line with their human orthologues. VGNC species currently include chimp, macaque, cow, dog, horse, pig, and cat. We also work with other nomenclature committees responsible for naming genes in model vertebrates, such as mouse, rat, and Xenopus, to ensure consistency across species when possible (Tweedie et al., 2021).Every named human gene has a symbol report on the HGNC website listing key data, including the approved nomenclature, published aliases, and locus type. An HGNC symbol report also contains links to multiple relevant sequence databases and clinical resources. It may additionally contain a link to a gene group page (see below), links to VGNC pages for orthologues in selected vertebrate species, and links to key publications in Europe PMC and PubMed. All data including our nomenclature guidelines (Bruford et al., 2020) can be accessed via our website.The green alga C. reinhardtii is a key model organism for studying eukaryotic cilia and flagella and the dynein motor complexes that aid in their assembly and drive their movement. The alveolate Tetrahymena thermophila and sea urchins such as Strongylocentrotus purpuratus are also key model organisms for studying ciliary function. The nomenclature of human dyneins has been largely based on orthology with C. reinhardtii, but also partly based on sea urchin nomenclature. Unfortunately, there are inconsistencies in the naming of orthologues among these species due to historic numbering assignments based on protein migration in SDS/urea-polyacrylamide gels. We have brought mammalian dynein nomenclature more into line with that of C. reinhardtii where possible and have established a naming system for genes encoding dynein chains that are unique to vertebrate species.While the stability of gene symbols, particularly those associated with phenotypes, is now a priority for the HGNC, we are still willing to consider updates for genes approved with placeholder symbols or for genes with domain-based nomenclature that may not give a clue to the function of the encoded protein, for example, genes named based on whether their encoded proteins contain transmembrane domains or coiled-coil regions (CCDC). Symbol changes are made only if an approved symbol has not become entrenched in the literature and if the community working on the gene in question is supportive of change to something more functionally informative.In 2005, the nomenclature for the mammalian cytoplasmic dynein genes was revised (Pfister et al., 2005). The introduction of new DYNC1 and DYNC2 root symbols helped clarify whether genes encoded subunits that were components of the dynein 1 or dynein 2 complex. New root symbols were also introduced to subdivide the known human dynein light chains into three families: roadblock (DYNLRB), Tctex (DYNLT), and LC8 (DYNLL). A 2011 paper (Hom et al., 2011) reported updates made to C. reinhardtii dynein gene nomenclature based on the structural properties of their encoded protein products. This more systematic naming system helped to make the cross-species comparison of orthologues more straightforward and provided a framework for naming newly characterized dynein-encoding genes. Note that there are several human genes encoding dynein chains without orthologues in C. reinhardtii, as it lacks an equivalent of the cytoplasmic dynein 1/dynactin system, so some of the nomenclature is mammal specific.Here we discuss our recent nomenclature updates for genes encoding dynein complex subunits, ODA-DC subunits, and axonemal dynein assembly factors in the human genome (Tweedie et al., 2021), as well as in the model organisms that follow HGNC nomenclature such as mouse, rat, and Xenopus.Table 2.Summary table of nomenclature updates reported here
Open in a separate windowaInformation about C. reinhardtii ciliary proteins, including dynein components, is curated and available at http://chlamyfp.org/.bChlamydomonas encodes two paralogous proteins that both have the same human orthologue.cReserved symbol/alias symbol. This gene will either be updated as a DNAAF or a DNAAF symbol will be added as an alias if further future publications support this.Gene groupsHGNC gene groups are manually curated using data from publications and advice from our specialist advisors. The groups for genes encoding the subunits of human dynein complexes can be viewed here: https://www.genenames.org/data/genegroup/#!/group/537 and reflect the data shown in Table S1, Table S2, Table S3, Table S4, Table S5, and Table S6.Discussion of HGNC nomenclature updates for dyneins and their cytoplasmic assembly factors
Phenotype | Associated dynein or dynein-related gene variantsa | Selected associated publications (PubMed ID) | OMIM MIM number (phenotype subtype) |
---|---|---|---|
Primary ciliary dyskinesia (PCD): abnormal ciliary motility, respiratory distress, sinusitis, otitis media, bronchiectasis, laterality defects, infertility | DNAH1 | 11371505 20301301 24360805 | 617577 (CILD37) |
DNAH5 | 11062149 11788826 | 608644 (CILD3) | |
DNAH9 | 30471717 30471718 | 618300 (CILD40) | |
DNAH11 | 12142464 | 611884 (CILD7) | |
DNAI1 | 10577904 | 604366 (CILD1) | |
DNAI2 | 18950741 | 612444 (CILD9) | |
DNAL1 | 21496787 | 614017 (CILD16) | |
NME8 (alias DNAI8 and TXNDC3) | 17360648 | 610852 (CILD6) | |
ODAD1 | 23261302 23261303 23506398 30291279 32855706 | 615067 (CILD20) | |
ODAD2 | 23849778 24203976 25186273 | 615451 (CILD23) | |
ODAD3 | 24067530 25192045 25224326 30504913 31383820 | 616037 (CILD30) | |
ODAD4 | 27486780 | 617092 (CILD35) | |
DNAAF1 | 19944400 19944405 27261005 | 613193 (CILD13) | |
DNAAF2 | 31107948 32638265 34785929 | 612518 (CILD10) | |
DNAAF3 | 22387996 31186518 | 606763 (CILD2) | |
DNAAF4 | 23872636 | 615482 (CILD25) | |
DNAAF5 | 29358401 25232951 23040496 | 614874 (CILD18) | |
DNAAF6 | 32170493 | 300991 (CILD36) | |
ZMYND10 | 23604077 23891469 23891471 | 615444 (CILD22) | |
LRRC6 | 23122589 | 614935 (CILD19) | |
LRRC56 | 30388400 | 618254 (CILD39) | |
SPAG1 | 24055112 26228299 | 615505 (CILD28) | |
CFAP298 | 24094744 | 615500 (CILD26) | |
CFAP300 | 29727692 29727693 | 618063 (CILD38) | |
Spinal muscular atrophy (SMALED type 1): lower limb atrophy and weakness, mild to moderate cognitive impairment | DYNC1H1 | 24307404 25609763 32788638 | 158600 (SMALED) |
BICD2 | 26998597 29353221 32709491 | 615290 (SMALED2A) 618291 (SMALED2B) | |
Charcot-Marie-Tooth type 2: distal lower limb weakness, abnormal gait | DYNC1H1 | 24307404 20697106 22459677 22847149 33242470 | 614228 (CMT2O) |
DNAH10 | 26517670 | Not listed in OMIM | |
Asphyxiating thoracic dystrophies (including Jeune syndrome): skeletal abnormalities that may include short ribs and a chest wall deformity, shortened arm and leg bones, an unusually shaped pelvis, polydactyly, renal and hepatic disease (more rarely, retinal disease) | DYNC2H1 | 19442771 26874042 27925158 31935347 | 613091 (SRTD3) |
DYNC2I1 | 23910462 26874042 29271569 | 615503 (SRTD8) | |
DYNC2I2 | 24183449 24183451 | 615633 (SRTD11) | |
DYNC2LI1 | 26130459 | 617088 (SRTD15) | |
DYNLT2B | 25830415 26044572 28475963 | 617405 (SRTD17) | |
Retinal degeneration | DYNC2H1 | 32753734 | Not listed in OMIM |
Nonsyndromic rod-cone dystrophy | DYNC2I2 | 33124039 | Not listed in OMIM |
Neurodevelopmental disorder with microcephaly and structural brain anomalies | DYNC1I2 | 31079899 | 618492 (NEDMIBA) |
Mirror movements type 3: movements on one side of the body are involuntarily mirrored on the other side of the body | DNAL4 | 25098561 | 616059 (MRMV3) |
Mental retardation autosomal dominant 13 | DYNC1H1 | 23603762 22368300 | 614563 (MRD13) |
Spermatogenic failure | DNAH1 | 24360805 33989052 | 617576 (SPGF18) |
DNAH2 | 30811583 | 619094 (SPGF45) | |
DNAH8 | 32619401 | 619095 (SPGF46) | |
DNAH17 | 31178125 31658987 31841227 | 618643 (SPGF39) | |
Lissencephaly: developmental delay, myoclonic jerks and spasms, seizures, hypotonia, microcephaly, dysmorphic facies | PAFAH1B1 | 32692650 20301752 32341547 28886386 | 601545 (LIS) |
Seckel syndrome: growth retardation, microcephaly, developmental delay | NIN | 27053665 22933543 | 614851 (SCKL7) |
Approved HGNC Symbol | Name | Aliases (previously approved symbols in bold) | Chlamydomonas orthologuea (genes and proteins) | Protein present in |
---|---|---|---|---|
DYNLT2 | Dynein light chain Tctex-type 2 | TCTE3, TCTEX1D3, TCTEX2, Tctex4 | DLT2 (LC2) | Axonemal ODA complex |
ODAD1 | Outer dynein arm docking complex subunit 1 | CCDC114, FLJ32926, CILD20 | DCC2 (ODA1) and DCC3 (ODA5)b | Axonemal ODA complex |
ODAD2 | Outer dynein arm docking complex subunit 2 | ARMC4, FLJ10817, FLJ10376, DKFZP434P1735, CILD23, gudu | No orthologue | Axonemal ODA complex |
ODAD3 | Outer dynein arm docking complex subunit 3 | CCDC151, MGC20983, ODA10 | DCC1 (ODA3) and ODA10 (ODA10)b | Axonemal ODA complex |
ODAD4 | Outer dynein arm docking complex subunit 4 | TTC25, DKFZP434H0115 | No orthologue | Axonemal ODA complex |
DNAI3 | Dynein axonemal intermediate chain 3 | WDR63, DIC3, FLJ30067, NYD-SP29 | DIC3 (IC140) | Axonemal IDA I1/f complex |
DNAI4 | Dynein axonemal intermediate chain 4 | WDR78, DIC4, FLJ23129 | DIC4 (IC138) | Axonemal IDA I1/f complex |
DNAI7 | Dynein axonemal intermediate chain 7 | CFAP94, CASC1, LAS1, FLJ10921, PPP1R54, IC97 | DII6 (FAP94) | Axonemal IDA I1/f complex |
DYNLT2B | Dynein light chain Tctex-type 2B | TCTEX1D2, MGC33212 | DLT4 (Tctex2b) | Axonemal IDA I1/f complex |
Cytoplasmic dynein 2 complex | ||||
DYNC2I1 | Dynein 2 intermediate chain 1 | WDR60, FLJ10300, FAP163, CFAP163, DIC6 | DIC6 (FAP163) | Cytoplasmic dynein 2 complex |
DYNC2I2 | Dynein 2 intermediate chain 2 | WDR34, DIC5, MGC20486, bA216B9.3, FAP133, CFAP133 | DIC5 (FAP133) | Cytoplasmic dynein 2 complex |
DYNLT3 | Dynein light chain Tctex-type 3 | TCTE1L, TCTEX1L | DLT1 (LC9) | Cytoplasmic dynein 2 complex |
DNAAF8 | Dynein axonemal assembly factor 8 | C16orf71, FLJ43261, DKFZp686H2240 | Axonemal dynein assembly factor | |
DNAAF9 | Dynein axonemal assembly factor 9 | C20orf194, DKFZp434N061 | DNAAF9 | Axonemal dynein assembly factor |
DNAAF10 | Dynein axonemal assembly factor 10 | WDR92, FLJ31741, Monad | DNAAF10 | Axonemal dynein assembly factor |
DNAAF11 | Dynein axonemal assembly factor 11 | LRRC6, TSLRP, LRTP, CILD19, tilB | DNAAF11, MOT47, LRRC6, Seahorse | Axonemal dynein assembly factor |
LRRC56 | Leucine rich repeat containing 56 | DNAAF12, FLJ00101, DKFZp761L1518 | DLU2 (ODA8) | Axonemal dynein assembly factor |
SPAG1 | Sperm associated antigen 1 | DNAAF13, SP75, FLJ32920, HSD-3.8, TPIS, CT140, CILD28, | SPAG1 (SPAG1) | Axonemal dynein assembly factor |
PIH1D1 | PIH1 domain containing 1 | DNAAF14, FLJ20643, Pih1, MOT48, | DAP2 (MOT48) | Axonemal dynein assembly factor |
PIH1D2 | PIH1 domain containing 2 | DNAAF15 | Axonemal dynein assembly factor | |
CFAP298 | Cilia and flagella associated protein 298 | FLJ20467, DAB2, FBB18, CILD26, Kur, C21orf48, C21orf59, DNAAF16 | DAB2 | Axonemal dynein assembly factor |
CCDC103 | Coiled-coil domain containing 103 | FLJ13094, FLJ34211, PR46b, CILD17, DNAAF17c | CCDC103 | Axonemal dynein assembly factor |
DAW1 | Dynein assembly factor with WD repeats 1 | FLJ25955, ODA16, WDR69, DNAAF18 | DAW1 | Axonemal dynein assembly factor |
Dynein light chain nomenclature updates (dynein light chain Tctex-type [DYNLT])
Based on advice from experts in the field, we have updated the nomenclature of all the Tctex family genes to better reflect the function of their encoded proteins as dynein subunits. The six paralogs in this set now use the root symbol DYNLT in human.DYNLT1 and DYNLT3
The gene currently approved as DYNLT1 (HGNC ID: 11697) was first approved using the symbol TCTEL1 based on homology with the mouse gene Tcte1 (t-complex associated testis expressed 1; Watanabe et al., 1996), which was reported to be specifically expressed in murine testes (Lader et al., 1989; Sarvetnick et al., 1989). The t-complex is a region of the mouse genome that shows non-Mendelian segregation, and some of the genes in it are associated with spermatogenesis (Castaneda et al., 2020). The alias symbol Tctex1 was also used to publish on this gene; it was characterized as encoding a cytoplasmic dynein light chain (Dedesma et al., 2006; King et al., 1998) and later also identified in axonemal inner arm I1/f (Harrison et al., 1998); in C. reinhardtii, a closely related protein is present in the ODA (DiBella et al., 2005).The most closely related paralogous gene to DYNLT1, now approved as DYNLT3 (HGNC ID: 11694), was originally assigned the symbol TCTE1L (Tcte1-like) in human, again to reflect its homology to mouse Tcte1. It was also published as a candidate for the retinitis pigmentosa RP3 locus (Roux et al., 1994), although this link was later disproven (Meindl et al., 1996) when RPGR (retinitis pigmentosa GTPase regulator) was identified as the causative gene for this phenotype (Ferrari et al., 2011). DYNLT3 was reported to encode a cytoplasmic dynein light chain in 1998 (King et al., 1998) and was later published as also playing a role in regulating primary cilium length (Palmer et al., 2011). We have constructed a phylogenetic tree (Fig. 4) that shows there is no clear 1:1 orthology relationship for either human DYNLT1 or DYNLT3 with respect to invertebrate species.Open in a separate windowFigure 4.Maximum-likelihood phylogenetic tree to show the relationship of Tctex-type dynein light chains in selected species. This tree is shown with a midpoint rooting. The figures on the nodes show the Shimodaira–Hasegawa likelihood ratio test and the Ultrafast bootstrap support values for the branches (SH-aLRT %/UFBoot %). Bootstrap values of ≥70% only are shown. The scale bar represents the expected number of amino acid substitutions per site. M. musculus has multiple Dynlt1 and Dynlt2 paralogs, but as these are identical at the amino acid level, only one sequence has been included in each case. The colors highlight supported clades: green for DYNLT1 and DYNLT3 and their orthologues, blue for DYNLT2 and its orthologues, red for DYNLT2B and its orthologues, yellow for DYNLT4 and its orthologue, and purple for DYNLT5 and its orthologues.DYNLT2 and DYNLT2B
We have updated the nomenclature of the gene previously approved as TCTE3 (HGNC ID: 11695) to DYNLT2, and that of its closely related paralog previously approved as TCTEX1D2 (Tctex1 domain containing 2; HGNC ID: 28482) to DYNLT2B. These new symbols are more functionally informative, and this update brings the human nomenclature into line with that of C. reinhardtii, S. purpuratus, and T. thermophila (see Fig. 4). The phylogeny (Fig. 4) shows the paralogous relationship between DYNLT2 and DYNLT2B and that their 1:1 orthologues in the other species fall into two separate subclades.Although DYNLT2 and DYNLT2B are paralogs, their protein products are components of distinct dynein complexes. DYNLT2 encodes an axonemal dynein subunit, required for outer arm assembly (Patel-King et al., 1997), and has not been reported as being part of any cytoplasmic dynein complex. The DYNLT2B-encoded protein is part of the cytoplasmic dynein 2 complex (Hamada et al., 2018; Schmidts et al., 2015) and is also an axonemal inner arm I1/f complex subunit (DiBella et al., 2004).DYNLT4 and DYNLT5
We have updated the nomenclature of the gene previously approved as TCTEX1D4 (HGNC ID: 32315) to DYNLT4. This gene encodes a dynein light chain protein that belongs to the TCTEX1 family. Freitas et al. (2014) discussed its role in sperm motility and IFT.While discussing the update for TCTEX1D4 with experts, we also proposed a nomenclature update for TCTEX1D1 (HGNC ID: 26882). This gene could not be updated to DYNLT1 in line with the TCTEX1D1 numbering, as this symbol was already in use, so we proposed an update to DYNLT5. There is currently a single paper published on this human gene (Spitali et al., 2020), linking a variant of it with the phenotype Duchenne muscular dystrophy. Although it seems likely that, as a paralog of the other DYNLT genes, DYNLT5 will be found to encode a dynein light chain, we have included the term family member in its current gene name to indicate that although it is related to the other DYNLT genes, a shared function has not yet been established. The phylogeny (Fig. 4) reveals that S. purpuratus has a 1:1 orthologue of DYNLT5, while C. reinhardtii and T. thermophila do not.DNAI nomenclature updatesDNAI3 and DNAI4
We have updated the nomenclature of the human orthologues of C. reinhardtii DIC3, encoding IC140 (alias IDA7); and DIC4, encoding IC138 (alias BOP5), to DNAI3 (HGNC ID: 30711) and DNAI4 (HGNC ID: 26252), respectively. These genes were previously approved as WDR63 (WD repeat domain 63) and WDR78. In C. reinhardtii, IC140 and IC138 have been well characterized as intermediate chain subunits of an IDA complex (I1 dynein complex, also known as dynein-f; Hendrickson et al., 2004; Yang and Sale, 1998). Updating WDR63 and WDR78 using the DNAI root brings their nomenclature in line with the other human genes encoding axonemal dynein intermediate chains, DNAI1 and DNAI2. It also keeps the numbering system used equivalent to that of the C. reinhardtii orthologues.The DNAI3-encoded protein is not essential for fertility in male mice, as other intermediate chains of the IDA I1/f complex may compensate for this role in mouse sperm motility (Young et al., 2015). The mouse orthologue of DNAI4 encodes a dynein intermediate chain in vertebrates. The DNAI4 protein interacts with multiple subunits of the axonemal inner arm I1/f dynein complex and is essential for the ciliary assembly of this complex in vertebrates (Zhang et al., 2019).DNAI7 and NME8 (alias DNAI8)
We originally considered updating the nomenclature of the gene previously approved as CASC1 (cancer susceptibility 1; HGNC ID: 48939) to DNAI5. However, after discussion with experts, we realized this could be confusing, as it is not the orthologue of C. reinhardtii DIC5, and all the other human DNAI genes are numbered in line with their C. reinhardtii orthologues. There is also a DIC6 gene in C. reinhardtii, and its orthologue is the human gene now approved as DYNC2I1 (dynein 2 intermediate chain 1).We were also reluctant to reassign CASC1 as DII6, the symbol used for the C. reinhardtii orthologue of this gene (Hom et al., 2011). We do not have an established DII# (dynein inner arm interacting) root approved in human, and most of the orthologues of the DII# C. reinhardtii genes are already approved and published using alternative symbols. These genes include DNALI1 (dynein axonemal light intermediate chain 1), the orthologue of DII1; ACTG1 (actin γ1), the orthologue of DII4; and ANK2 (ankyrin 2), the orthologue of DII7. In addition, with the exception of DNALI1, it is possible that one or more of these genes may not necessarily encode proteins that are dynein-arm interacting in vertebrates. Therefore, we updated CASC1 as DNAI7, reflecting that its protein product is a dynein intermediate chain in human. The mouse orthologue of DNAI7 encodes an intermediate chain in vertebrates that forms part of the inner arm I1/f dynein complex required for ciliary beating (Zhang et al., 2019).This leaves NME8 (NME/NM23 family member 8) as the only remaining human gene known to encode a dynein intermediate chain but not named as such. This gene was previously approved as TXNDC3 (thioredoxin domain containing 3; Duriez et al., 2007) and has also been published using the alias symbol SPTRX2 (sperm-specific thioredoxin 2; Sadek et al., 2001).There are 10 genes in the human NME/NM23 family, at least five of which encode active nucleoside diphosphate kinases (Ćetković et al., 2015). NME8 (HGNC ID: 16473) is the human orthologue of the sea urchin IC1 gene (Duriez et al., 2007), which encodes a sea urchin ODA intermediate chain and, like its human orthologue, contains an N-terminal thioredoxin-like domain (Ogawa et al., 1996). In C. reinhardtii, the ODA contains two paralogous thioredoxin-like light chains (LC3 and LC5) but lacks a nucleoside diphosphate kinase (Patel-King et al., 1996).NME8 encodes a protein with a ciliary role, and its gene product is suggested to be bifunctional, with isoforms expressed at varying levels in different tissues (Duriez et al., 2007). The TXNDC3d7 protein isoform can bind microtubules, plays a role in ciliary function, and may be a component of ODAs (Duriez et al., 2007). As NME8 is already named as part of a gene group, is a functionally informative symbol, and has been used in the literature, we have decided to retain this nomenclature. However, this gene has been assigned the alias symbol DNAI8 and added to our dynein axonemal outer arm complex subunits gene group page (https://www.genenames.org/data/genegroup/#!/group/2031).DYNC2I1 and DYNC2I2
We have updated the nomenclature of the human orthologue of C. reinhardtii DIC6 encoding D1bIC1 (alias FAP163) from WDR60 to DYNC2I1 (HGNC ID: 28296). We have also updated the nomenclature of the human orthologue of C. reinhardtii DIC5, encoding D1bIC2 (alias FAP133) from WDR34 to DYNC2I2 (HGNC ID: 21862). The numbering was assigned in this way so that the human gene nomenclature corresponds to that of the C. reinhardtii proteins.DIC5/FAP133 in C. reinhardtii is associated with the IFT dynein motor (dynein 2, usually known as dynein 1b in C. reinhardtii) complex (Rompolas et al., 2007). DIC6/FAP163 encodes a C. reinhardtii intermediate chain that is closely related to DIC5/FAP133 and is also a component of the dynein 2 complex (Patel-King et al., 2013). Previous studies linked these two genes to ciliopathies including short rib polydactyly and Jeune syndrome (McInerney-Leo et al., 2013; Schmidts et al., 2013) and suggested that these orthologues of C. reinhardtii dynein intermediate chains may also encode components of the dynein 2 complex. Indeed, it was confirmed that both human genes encode dynein 2 intermediate chains (Asante et al., 2014).ODA-DC (ODAD) nomenclature updatesODAD1, ODAD2, ODAD3, ODAD4, and CLXN (ODAD5)
The ODA-DC has only recently been characterized in human (Hjeij et al., 2014; Onoufriadis et al., 2013; Wallmeier et al., 2016), and it became apparent that the nomenclature of the genes encoding the constituent proteins was not as functionally informative as it could be. The nomenclature of four of the ODA-DC subunits was initially based on the presence of structural domains in the encoded proteins: ARMC4 (armadillo repeat containing 4), CCDC114 and CCDC151 (coiled-coil domain containing 114 and 151, respectively), and TTC25 (tetratricopeptide repeat domain 25), as there was no functional information published when they were initially named.These four genes have now been reassigned using the root symbol ODAD (ODA-DC subunits). The ODAD genes have been assigned numbers in the order in which they were characterized as encoding ODA-DC subunits in human and in line with the ODA numbering in C. reinhardtii where possible. We could not use the DCC root in human for these genes, as it clashed with the approved symbol for an unrelated gene, DCC (DCC netrin 1 receptor; HGNC ID: 2701).ODAD1 is the orthologue of C. reinhardtii DCC2 (encoding DC2, alias ODA1), which encodes a docking complex subunit, and of its paralog DCC3, which encodes the ODA5 assembly factor (Takada et al., 2002). ODAD3 is the orthologue of DCC1, which encodes the protein DC1 (alias ODA3; Koutoulis et al., 1997), a docking complex subunit in C. reinhardtii, and of its paralog ODA10, which encodes a dynein assembly factor in C. reinhardtii (Dean and Mitchell, 2013). ODAD2 and ODAD4 have no known C. reinhardtii orthologues.A fifth gene has recently been published in a study examining mammalian tracheal cilia as encoding an ODA-DC subunit (Gui et al., 2021; Fig. 3, c and d). Its encoded protein, calaxin, is a member of a neuronal calcium sensor family and was originally identified in ODAs from the sea squirt Ciona intestinalis; subsequent studies revealed it is required for normal ciliary motility in mice (Mizuno et al., 2009; Mizuno et al., 2012; Sasaki et al., 2019). We have updated its approved nomenclature from the previously approved but less frequently used EFCAB1 (EF-hand calcium binding domain 1) to CLXN (calaxin), aliasing it as ODAD5 after discussion with authors.The symbol ODAD6 is reserved for the gene currently approved as CCDC63, a closely related paralog of ODAD1. We will continue to monitor the literature and may update the nomenclature of this gene, either approving ODAD6 or adding it as an alias if CCDC63 is shown to encode an ODA-DC subunit. The ODA-DC gene group page can be seen on our website (https://www.genenames.org/data/genegroup/#!/group/2019).DNAAFsWe have updated the nomenclature of four genes as DNAAFs, including two previously assigned using placeholder C#orf# symbols (see Table S6). There are now 18 genes included in our axonemal dynein assembly factor gene group set (https://www.genenames.org/data/genegroup/#!/group/1627).We have updated the nomenclature of the gene previously approved using the placeholder symbol C16orf71 (chromosome 16 open reading frame 71; HGNC ID: 25081) to DNAAF8. The Xenopus orthologue was recently published using the alias symbol Daap1 (dynein axonemal-associated protein 1; Lee et al., 2020), but following discussion, this gene has been approved as dnaaf8 in line with its human orthologue.We have also updated the nomenclature of the gene previously approved as C20orf194 (chromosome 20 open reading frame 194; HGNC ID: 17721) to DNAAF9. The Tetrahymena orthologue of this gene was published using the alias name “shulin” (Mali et al., 2021). Those authors’ work showed that the encoded protein has a role in keeping the axonemal ODAs in a nonfunctional state before delivery to cilia. With these authors, our experts, and all researchers who had previously published on this gene, we discussed assigning this gene as DNAAF9, and they were supportive of this update. A gene (Cre11.g467556) exhibiting some similarity to DNAAF9 is present in C. reinhardtii; this is in a potentially poorly assembled genomic region, and further characterization will be required to determine whether it is the true orthologue of this human gene.Two other genes, previously approved as WDR92 and LRRC6 (leucine rich repeat containing 6), have also been updated to DNAAF10 and DNAAF11, respectively. Both have been shown to encode proteins that are involved in axonemal dynein assembly (Patel-King and King, 2016; Fabczak and Osinka, 2019; Liu et al., 2019; Patel-King et al., 2019; Li et al., 2021; Zur Lage et al., 2018). The DNAAF10 protein product interacts with the protein encoded by SPAG1 (sperm associated antigen 1; see below) during dynein preassembly (Zur Lage et al., 2018). The DNAAF11 protein product interacts with the protein encoded by ZMYND10 (zinc finger MYND-type containing 10), which is aliased as DNAAF7 (Zariwala et al., 2013). ZMYND10 has been retained as the approved symbol because it has been well used in publications, and the current nomenclature reflects the fact that the encoded protein contains a MYND-type zinc finger domain.We also assigned four other genes (LRRC56, SPAG1, PIH1D1, and PIH1D2) with DNAAF aliases to reflect the roles of their encoded proteins in dynein assembly (Bonnefoy et al., 2018; Knowles et al., 2013; Yamaguchi et al., 2018). These were assigned the alias symbols DNAAF12, DNAAF13, DNAAF14, and DNAAF15, respectively. Although it seems very likely based on two publications (Bonnefoy et al. [2018] and Desai et al. [2015]) that LRRC56 encodes a DNAAF, we are continuing to monitor the literature and could consider updating the nomenclature of this gene to DNAAF12 if there is sufficient evidence published to support this.The SPAG1 and PIH1D1 symbols are already well established in the literature, and SPAG1, PIH1D1, and PIH1D2 all encode proteins that are subunits of complexes with many other functions as well as being involved in dynein assembly (von Morgen et al., 2015). The PIH1D2- and SPAG1-encoded proteins are part of the R2SP complex (Chagot et al., 2019), and the PIH1D1-encoded protein is part of the R2TP complex (Rodríguez and Llorca, 2020). Therefore, we have chosen to retain their currently approved symbols but have added them to our DNAAF gene group page. While we always ask that authors reference the approved gene symbols at least once in all publications, they can of course also use the DNAAF aliases.We also discussed a DNAAF symbol update for the orthologue of C. reinhardtii DAB2 with authors and our expert advisors. DAB2 accumulates in cilia, and their motility is impaired (Austin-Tse et al., 2013). Variants of the Danio rerio orthologue of this gene, Kurly, are found in zebrafish mutants that display abnormalities in their development and have dynein arm defects, suggesting that the Kurly protein plays a role in ciliary motility but is also involved in regulating planar cell polarity (Jaffe et al., 2016). The human orthologue, previously approved as C21orf59, encodes a protein that has been shown to interact with known DNAAFs, including proteins encoded by ZYMND10 and DNAAF11 (previously LRRC6; Cho et al., 2018), and has been associated with the human phenotype PCD (Bolkier et al., 2021). Discussion with authors and our specialist advisors for the DNAAFs and cilia- and flagella-associated proteins (CFAPs) revealed community support for assigning a more general CFAP symbol for this gene. Its association with cilia and flagella is clear, and it also has a wider function beyond its role as an axonemal dynein arm assembly factor. However, while we have updated this gene as CFAP298 (HGNC ID: 1301), we have also assigned it the alias symbol DNAAF16. We also updated another cilia-associated gene, the orthologue of C. reinhardtii FBB5, as CFAP300 (previously approved as C11orf70) and have assigned it the alias symbol DNAAF17. Phylogenetic analysis strongly suggests that this gene is specific to organisms with motile cilia (being part of the MotileCut grouping; Merchant et al., 2007), and our CFAP nomenclature specialist advisor supported this change. As more becomes known about the function of the CFAP300 protein, we can consider whether a further symbol change would be helpful for this gene.We are retaining the symbol DAW1 (dynein assembly factor with WD repeats 1), as it is the orthologue of C. reinhardtii DAW1 and its current nomenclature is functionally informative. However, we have aliased it as DNAAF18 and added it to the DNAAF gene group. We have also reserved the gene symbol DNAAF19 for the gene currently approved as CCDC103. The CCDC103 protein affects dynein assembly (King and Patel-King, 2020; Panizzi et al., 2012), but its exact role has still to be defined.ConclusionIn total, we have updated the nomenclature of nine genes encoding human dynein chains, four genes encoding proteins that form the ODA-DC, and four genes encoding axonemal dynein assembly factors. Several other genes have retained their current symbols but have been aliased as ODADs or DNAAFs and added to the appropriate HGNC gene group pages. All updates were made following consultation with experts from the community, and these changes were widely supported among the authors publishing in this field. While we aim to limit changes in gene nomenclature, especially when the genes are linked to a phenotype, these updates have largely replaced uninformative placeholder or domain-based symbols, and we view the new informative symbols as stable. As such, users should regard these new symbols as the permanent gene symbols for these human genes.We hope that all researchers will use the new nomenclature in their future publications to aid communication and data retrieval within the field. Approved symbols should be mentioned at least once in publications, along with the associated HGNC ID if possible.Materials and methodsDynein light chain phylogenetic treeAmino acid protein sequences for dynein light chains were obtained for each of the six selected species from NCBI. A multiple alignment was built using the MUSCLE online tool (https://www.ebi.ac.uk/Tools/msa/muscle/; Madeira et al., 2019) and edited using AliView 1.20 (Larsson, 2014). The ends of the alignment were trimmed, and all indels were removed. The IQ-TREE web server (http://iqtree.cibiv.univie.ac.at/) was used to construct a maximum-likelihood tree. The substitution model was autoselected with ultrafast bootstrapping and SH-aLRT branch test methods applied.Online supplemental materialThe supplementary tables show HGNC approved nomenclature for genes encoding subunits of dynein complexes alongside their known published alias symbols and their orthologs in C. reinhardtii. Table S1 shows cytoplasmic dynein 1 subunits. Table S2 shows cytoplasmic dynein 2 subunits. Table S3 shows axonemal ODA subunits. Table S4 shows monomeric dynein heavy chains and their accessory subunits. Table S5 shows axonemal inner arm dynein I1/f subunits. Table S6 shows axonemal dynein assembly factors (DNAAFs). 相似文献18.
Karen K. Siu Vitor Hugo B. Serro Ahmed Ziyyat Jeffrey E. Lee 《The Journal of cell biology》2021,220(10)
Fertilization is defined as the union of two gametes. During fertilization, sperm and egg fuse to form a diploid zygote to initiate prenatal development. In mammals, fertilization involves multiple ordered steps, including the acrosome reaction, zona pellucida penetration, sperm–egg attachment, and membrane fusion. Given the success of in vitro fertilization, one would think that the mechanisms of fertilization are understood; however, the precise details for many of the steps in fertilization remain a mystery. Recent studies using genetic knockout mouse models and structural biology are providing valuable insight into the molecular basis of sperm–egg attachment and fusion. Here, we review the cell biology of fertilization, specifically summarizing data from recent structural and functional studies that provide insights into the interactions involved in human gamete attachment and fusion.IntroductionDuring sexual reproduction, the oocyte and sperm fuse to generate a new and unique embryo. The journey of a sperm to an egg ends in the ampulla of the female oviduct. From there, the sperm must overcome a number of physical and biochemical barriers. After undergoing the acrosome reaction and binding the ova, the sperm penetrates through the cumulus oophorus cells and the zona pellucida (ZP) to reach the perivitelline space (PVS) and oocyte membrane. Upon fusion of the sperm and egg membranes, the sperm nucleus and organelles are incorporated into the egg cytoplasm.An understanding of the mechanisms of mammalian fertilization is crucial to treat infertility and develop new methods of birth control. Infertility affects 15% of couples globally, and in one third of these cases, the underlying cause is unknown (Gelbaya et al., 2014). Developments in assisted reproductive technologies have provided couples with new options to conceive but may have epigenetic side effects (Mani et al., 2020). Furthermore, only 40% of couples manage to have a child despite 2 yr of treatment. Safety, efficacy, and acceptability of contraceptives are also critically important, but many current female contraceptive methods have side effects that limit long-term use (Aitken et al., 2008), while male contraceptives are limited to condoms or vasectomy (Kanakis and Goulis, 2015). A better understanding of the molecular players involved in fertilization is necessary to drive innovation in both assisted reproductive technologies and contraception.In this review, we will first briefly review the events that prepare the gametes for fertilization. We will then discuss how recent studies of genetically altered mice and structural biology efforts have shed light on the molecular mechanisms of sperm–egg attachment and fusion. We will also discuss the gaps in current knowledge and suggest new perspectives and future directions in the search for other protein factors involved at the gamete fusion synapse.Cell biology of gametesFertilization requires proper gametogenesis (oogenesis in the female and spermatogenesis in the male), which produces haploid cells and introduces diversity. Primordial germ cells (PGC) are the embryonic precursors to spermatocytes and ova. The cells produced by the first few divisions of the fertilized egg are totipotent and capable of differentiating into any cell type, including germ cells. PGCs originate within the primary ectoderm of the embryo and then migrate into the yolk sac. Between weeks 4 and 6, the PGCs migrate back into the posterior body wall of the embryo, where they stimulate cells of the adjacent coelomic epithelium and mesonephros to form primitive sex cords and induce the formation of the genital ridges and gonads. The sex (gonadal) cords surround the PGCs and give rise to the tissue that will nourish and regulate the development of the maturing sex cells (ovarian follicles in the female and Sertoli cells in the male).EggOogenesis is a complex differentiation process by which mature functional ova develop from germ cells (Fig. 1 A; Edson et al., 2009). In humans, oogenesis begins in the ovary at 6–8 wk of fetus development, when PGCs differentiate into oogonia. By the 12th week, several million oogonia enter prophase, the first meiotic division and become dormant until shortly before ovulation (Hayashi et al., 2020). Due to their large and watery nuclei, these cells are referred to as germinal vesicles (Pan and Li, 2019). These primary oocytes become enclosed by follicle cells to form primordial follicles. The number of primordial follicles peaks at ∼7 million by the fifth month of fetal life, with ∼700,000 left at birth and 400,000 by puberty (Marcozzi et al., 2018). All of the egg cells that the ovaries will release are already present at birth.Open in a separate windowFigure 1.Gametogenesis and fertilization. (A–C) Illustration of oogenesis and follicle development (A), spermatogenesis (B), and the major steps in fertilization (C): (1) initial contact, (2) acrosome reaction, (3) ZP penetration, (4) sperm–egg fusion, (5) entry of sperm nucleus, (6) cortical reaction, and (7) fusion of the sperm and egg nuclei. The oocyte with its ZP measures 130 μm in diameter. Created with BioRender.During each menstrual cycle, hormones from the hypothalamic–pituitary–gonadal axis restart the division of the primary oocytes in meiosis I and follicular development (Atwood and Vadakkadath Meethal, 2016). Primary follicles develop into secondary follicles, containing each growing oocyte surrounded by two or more layers of proliferating follicle cells. ZP glycoproteins are secreted by the oocyte of the primary follicle and possibly the follicular cells (Törmälä et al., 2008). Although these glycoproteins form a physical barrier between the follicle cells and the oocyte, follicle cells and the oocyte remain connected through transzonal cytoplasmic projections from the follicle cells until fertilization (Makabe et al., 2006). A reciprocal dialog between the oocyte and its surrounding follicular cells coordinates the different phases of follicular development and the maintenance of meiotic arrest (Dalbies-Tran et al., 2020). Oocyte-derived microvilli control female fertility by optimizing ovarian follicle selection in mice (Zhang et al., 2021). The epithelium of 5–12 primary follicles proliferates to form a multilayered capsule around the oocyte. A few of these growing follicles continue to enlarge in response to follicle-stimulating hormone (FSH; Visser and Themmen, 2014). A single follicle becomes dominant, and the others degenerate by atresia (Atwood and Vadakkadath Meethal, 2016). Meiosis of the oocyte in the mature preovulatory follicle is blocked until a surge in levels of FSH and luteinizing hormone that occurs midway through the menstrual cycle. The membrane of the germinal vesicle nucleus breaks down, the chromosomes align in metaphase, and the oocyte expels its first polar body. The secondary oocyte then begins the second meiotic division, which is arrested at the meiotic metaphase II stage until ovulation (Gougeon, 1996). Ovulation depends on the breakdown of the follicle wall and occurs ∼38 h after the increase in levels of FSH and luteinizing hormone (Holesh et al., 2021). The disruption of the follicle wall expulses the oocyte, which is captured by the fimbriated mouth of the oviduct and moved into the ampulla. The oocyte retains its ability to be fertilized for ∼24 h and completes meiosis only if it is fertilized.SpermIn contrast to oogenesis, which is complete before birth, spermatogenesis is a continuous process that begins at puberty (Fig. 1 B). In humans, spermatogenesis takes 74 d to complete; thus, multiple spermatogenesis events occur simultaneously to allow for continual sperm production. Spermatogenesis occurs in the testis in a stepwise manner, beginning with diploid spermatogonia at the basal surface of seminiferous tubules and ending with mature elongated spermatozoa that are released in tubule lumens in a process called spermiation (Clermont, 1972; Yang and Oatley, 2014). During spermatogenesis, mitosis results in gene amplification, meiosis results in genome reduction, and finally maturation occurs (Hess and Renato de Franca, 2008). At this stage, sperm are not motile and are fertilization incompetent. Two additional sperm maturational processes are required outside the testis. First, sperm undergo a maturation process during epididymal transit (Bedford et al., 1973) involving posttranslational modifications of previously synthesized proteins and acquisition of proteins from the epididymal epithelium (James et al., 2020; see text box). After ejaculation into the female reproductive tract, dilution triggers additional changes in sperm, collectively termed capacitation (see text box), that prepare the sperm for the acrosome reaction.Epididymal maturationSperm exchange with the epididymal epithelium occurs by direct interaction with epithelial cells, by interaction with soluble proteins in the epididymal fluid or via extracellular exosome-like vesicles released by epithelial cells called epididymosomes (James et al., 2020). The purposes of this exchange are to redistribute sperm proteins and change the composition and lipid balance of the sperm membrane. These changes take place during the transit from the epididymis initial segment, through the caput and the corpus, to the cauda where sperm are stored (Cornwall, 2009). Epididymal transit lasts 10–12 d in mammals, but storage is dependent on sexual activity. Since fertilization is not immediate, fertilizing capacities of the spermatozoa are preserved by decapacitation factors that are active in the epididymis. An example of a decapacitation factor is SPINK3, which is secreted by seminal vesicles; it impairs sperm membrane hyperpolarization and calcium influx through CatSper (Zalazar et al., 2020). Epididymal plasma and sperm represent only a small fraction (5%) of semen in men (Batruch et al., 2011). Two thirds of the volume of semen comes from the seminal vesicles and the other third from the prostate. These secretions protect the sperm and prevent early maturation.Sperm capacitationMore than 70 yr ago, Austin and Chang described capacitation as the changes required for sperm to fertilize oocytes in vivo (Austin, 1952; Chang, 1951). Once sperm enter the female reproductive tract, they undergo capacitation. Capacitation results in hyperactivation of sperm movement and initiation of the acrosome reaction (Saling et al., 1979; Florman and First, 1988). During capacitation, stabilizing or decapacitation factors that are adsorbed on the sperm plasma membrane are removed (Bedford and Chang, 1962). These agents that initiate removal of decapacitation factors are electrolytes, energy substrates, and proteins such as seminal plasma protein or albumin. Removal of decapacitation factors increases sperm plasma membrane fluidity, allowing an increase in the permeability to calcium, chloride, and bicarbonate ions (Gangwar and Atreja, 2015). Sperm motility depends on the membrane potential, intracellular pH, and balance of intracellular ions (reviewed in Nowicka-Bauer and Szymczak-Cendlak, 2021). The most important ion for this function is Ca2+ (Hwang et al., 2019). This secondary messenger is an important signaling pathways activator that regulates sperm motility (Finkelstein et al., 2020). The activation of soluble adenyl cyclases generates cyclic adenosine monophosphate that in turn activates serine/threonine protein kinase A, which induces a cascade of protein phosphorylation initiating the induction of sperm motility (Chen et al., 2000). Protein phosphorylation, sperm hyperactivation, and the acrosome reaction are used in vitro to evaluate capacitation. Capacitation can be induced in vitro by incubation in medium containing calcium, bicarbonate ions, and serum albumin (Touré, 2019).Mammalian sperm capacitation occurs during sperm migration in the female tract. Mammalian males ejaculate millions of sperm cells into the female reproductive tract, but only a few hundred sperm at most reach the oocytes. This massive elimination process likely prevents polyspermy (reviewed in Kölle, 2015). Selection of human sperm during the journey begins in the acidic environment of the vagina. In the cervix, only morphologically normal sperm can migrate. Some sperm immediately pass into the cervical mucus, whereas the remaining sperm becomes a part of the coagulum. The next selection occurs at the uterus–tubal junction, the connection between the uterus and the oviduct that represents a major obstacle for sperm migration (Kölle, 2015). Experiments in mice indicate that sperm motility alone is insufficient for sperm migration through the uterus–tubal junction (Fujihara et al., 2018). Uterine contractions facilitate sperm transport as do molecular interactions. Several proteins, such as ADAM3 and other ADAM family members, are known to be involved in this step in mice (Yamaguchi et al., 2009; Xiong et al., 2019); most ADAM proteins have human orthologues.Spermatogenesis takes place in a species-specific cycle called the seminiferous epithelial cycle and is regulated in particular through the hypothalamic–pituitary–testicular axis. Indeed, at puberty, the testes (interstitial steroidogenic Leydig cells) secrete an increased amount of testosterone, which triggers growth of the testes, maturation of the seminiferous tubules, and the commencement of spermatogenesis. The Sertoli cells are the major somatic cells present in the seminiferous tubules and are considered to be the main regulators of spermatogenesis. They orchestrate spermatogenesis by supporting spermatogonial stem cells, determining the testis size, organizing meiotic and postmeiotic development and sperm output, supporting androgen production by maintaining the development and function of Leydig cells, and regulating other aspects of testis function like peritubular myoid cells, immune cells, and the vasculature, which participate in the maintenance of the spermatogonial stem cell niche.Acrosome reactionThe acrosome is a secretory vesicle located on the anterior region of sperm that originates from the spermatid Golgi apparatus. An acrosomal granule is formed by the fusion of proacrosomal vesicles in the vicinity of the nucleus. The region increases in size and spreads over the anterior part of the nucleus. The acrosome reaction is driven by SNARE complexes and results in the exocytosis of the contents of the acrosome upon fusion of the plasma membrane with the outer acrosomal membrane (Fig. 1 C; reviewed in Okabe, 2016; De Blas et al., 2005). The timing of the acrosome reaction is critical. Only sperm that have undergone this reaction are fertilization competent, but when a high proportion of sperm undergo the acrosome reaction prematurely, success of in vitro fertilization is low (Wiser et al., 2014). Several studies indicate that only a fraction of sperm is capable of undergoing spontaneous acrosome reaction. In human and mice sperm samples, 15–20% of cells undergo spontaneous acrosome reaction (Nakanishi et al., 2001), whereas only 20–30% undergo progesterone-induced acrosome reaction (Stival et al., 2016), suggesting physiological heterogeneity of sperm population. In addition, Inoue et al. demonstrated that acrosome-reacted mouse spermatozoa recovered from the PVS can fertilize other eggs (Inoue et al., 2011).Based on in vitro data, it was thought that the acrosome reaction occurs when the sperm contacts the ZP, particularly the ZP3 protein (Litscher and Wassarman, 1996). Using transgenic mice that express fluorescent markers in the acrosome (Nakanishi et al., 1999) and the midpiece mitochondria (Hasuwa et al., 2010), real-time observation of acrosomal exocytosis was possible. These experiments showed that most mouse spermatozoa capable of fertilization had undergone the acrosome reaction before contact with the oocyte ZP (Jin et al., 2011). Most spermatozoa begin to react in the isthmus of the oviduct before reaching the ampulla (Hino et al., 2016; La Spina et al., 2016). Contact with the ZP in vitro probably makes it possible to complete a partial acrosome reaction. The most important function of the acrosome reaction is to induce changes in the sperm membrane (Okabe, 2016). The relocations of IZUMO1 and SPACA6, proteins essential for sperm–egg fusion, that occur after the acrosome reaction are illustrative examples of these changes (Sosnik et al., 2009; Barbaux et al., 2020; Satouh et al., 2012). The presence of these proteins on the sperm membrane, in addition to the classic markers Pisum sativum agglutinin, Peanut agglutinin lectins, or CD46, can be used as markers for the acrosome reaction (Ito and Toshimori, 2016). The acrosome and its disruption are both crucial for effective fertilization, as low fertilization rates are observed upon intracytoplasmic sperm injection of acrosome-intact sperm (Morozumi and Yanagimachi, 2005) or round spermatozoa lacking acrosomes (Dávila Garza and Patrizio, 2013).ZP penetrationThe ZP is a physical barrier between the oocyte and the follicular cells that forms from glycoproteins secreted from the primary follicles (Fig. 1 C). The human ZP consists of four glycoproteins (hZP1–hZP4; Harris et al., 1994). Mice, which have been used for most of the ZP studies in mammals, express only three ZP glycoproteins (mZP1–mZP3; Litscher and Wassarman, 2007). Analysis of mouse lines expressing human ZP proteins demonstrated that only hZP2 is important in human sperm–egg binding (Gupta, 2021). Experiments using purified native or recombinant human ZP proteins have shown that hZP1, hZP3, and hZP4 bind to the capacitated human spermatozoa and induce the acrosome reaction (Gupta, 2021). ZP1 is required for the structural integrity of the ZP (Chakravarty et al., 2008). To better understand the roles of ZP glycoproteins, further studies, particularly on ZP protein glycosylation, are needed. The species-specific binding of the ZP to sperm is presumably related to these carbohydrate moieties (Clark, 2014). The sialyl-Lewis(x) sequence is the major carbohydrate ligand for human sperm–egg binding (Pang et al., 2011). The current hypothesis that hZP1, hZP3, and hZP4 bind to capacitated sperm and hZP2 binds to sperm with intact acrosomes will need to be revisited due to the recent demonstration that the acrosome reaction takes place before ZP contact. Regardless, the role of the ZP in preventing polyspermy is clear. Indeed, ZP hardening is due to ZP2 cleavage by ovastacin, a protease released into the PVS by cortical granules after the first sperm–egg fusion (Burkart et al., 2012).Sperm–egg attachment and membrane fusionAfter penetration of the ZP, the sperm enters the PVS and can attach and fuse with the egg plasma membrane. The development of genetic knockout animal models has proven critical in determining the importance of various sperm and egg proteins in sperm–egg attachment and fusion. Surprisingly, genetic knockout studies revealed that many factors originally thought to be important for fertilization were in fact not necessary (reviewed in Okabe, 2018, 2015). The proteins from sperm and egg that are essential for sperm–egg membrane interaction and fusion are listed in Protein Year identified Role in fertilization Structural features References CD9 1999 CD9 is expressed on the surface of the oocyte and accumulates during the attachment event; it may modulate the integrity of the oocyte membrane; its precise role in sperm–egg fusion remains unclear CD9 is a tetraspanin with four transmembrane domains and two extracellular loops (short and long) Miyado et al., 2000; Le Naour et al., 2000; Kaji et al., 2000; Chen et al., 1999; Umeda et al., 2020; Zimmerman et al., 2016; Zhang and Huang, 2012; Dahmane et al., 2019; Runge et al., 2007; Zhu et al., 2002; Chalbi et al., 2014; Rubinstein et al., 2006; Ziyyat et al., 2006 IZUMO1 2005 IZUMO1 relocates to the equatorial region of the sperm head after the acrosome reaction; high-affinity binding of IZUMO1 to JUNO results in initial attachment of sperm and egg in the PVS The protein has an N-terminal 4HB, followed by a β-hinge and an IgSF domain; the structure is stabilized by five disulfide bonds Inoue et al., 2005; Ellerman et al., 2009; Young et al., 2015; Satouh et al., 2012; Aydin et al., 2016; Ohto et al., 2016; Nishimura et al., 2016; Kato et al., 2016 JUNO 2014 JUNO is expressed on the surface of the oocyte membrane and serves as the receptor of IZUMO1 JUNO has structural similarity to folate receptors; it is a globular α/β protein composed of five α helices, three 310 helices, and four short β strands stabilized by eight disulfide bonds Bianchi et al., 2014; Kato et al., 2016; Han et al., 2016; Jean et al., 2019; Yamaguchi et al., 2007; Aydin et al., 2016; Ohto et al., 2016 SPACA6 2014 SPACA6 is expressed in sperm and localized to the equatorial segment after the acrosome reaction, but its specific role in sperm–egg fusion remains unknown The three-dimensional structure of SPACA6 is currently unknown; SPACA6 is similar in organization to IZUMO1 with a signal peptide, followed by an α-helical domain, an IgSF domain, a transmembrane helix, and a cytoplasmic tail Lorenzetti et al., 2014; Noda et al., 2020; Barbaux et al., 2020 TMEM95 2014 TMEM95 is localized to the equatorial segment of sperm and is essential for sperm–egg fusion and male fertility in mice, but its specific role in sperm–egg fusion is currently unknown The structure of TMEM95 is currently unknown; TMEM95 consists of a signal peptide, an N-terminal helix-rich region, a transmembrane helix, and a leucine-rich cytoplasmic domain Pausch et al., 2014; Zhang et al., 2016; Noda et al., 2020; Fernandez-Fuertes et al., 2017; Lamas-Toranzo et al., 2020 SOF1 2020 SOF1 is predicted to be a secreted factor essential for fusion; its role is still not fully understood No structural information to date; primary sequence analysis revealed the presence of conserved LLLL and CFNLAS motifs Noda et al., 2020 FIMP 2020 FIMP is involved in sperm–egg fusion; only the transmembrane form is important in fertilization, but its role is still not fully determined No structural information to date Fujihara et al., 2020 DCST1/DCST2 2021 DCST1 and DCST2 are involved in sperm–egg fusion; stability of SPACA6 is regulated by DCST1/2; DCST1/DCST2 are evolutionary conserved in vertebrates and invertebrates No structural information to date; contains six putative transmembrane helices Inoue et al., 2021