TY - JOUR
AU - Ikegami,, Koji
AB - Abstract The cilium is a tiny organelle, with a length of 1–10 μm and a diameter of ~200 nm, that projects from the surface of many cells and functions to generate fluid flow and/or sense extracellular signals from the environment. Abnormalities in cilia may cause a broad spectrum of disease, i.e. the so-called ciliopathies. Multiple imaging approaches have been implemented to understand the structure, motion and function of the tiny cilium. In this review, we focus on the microscopic observations and analyses of the dynamic behaviors of both motile cilia and primary cilium. Motile cilia repeat reciprocal motions at 15–25 Hz with a clear asymmetry of effective and recovery strokes. Observing the fast movement of motile cilia requires a high-speed camera with a frame rate of more than 100 fps. The labeling of cilia tips enables the detailed analysis of the asymmetric beating motion of motile cilia. The primary cilium, which is imagined to be ‘static,’ is also dynamic, i.e. it elongates, shrinks and disassembles, although this behavior is quite slower than that of motile cilia. The specific fluorescent labeling of primary cilium and time-lapse imaging are required to observe and analyze the slow behaviors of the primary cilium. We present some approaches, including some tips for successful procedures, in the successful imaging of the dynamic behaviors of motile cilia and primary cilium. ciliary beat frequency, time-lapse microscopy, ciliopathies, cilium, differential interference contrast microscopy, primary ciliary dyskinesia Introduction In 1675, Anton Van Leeuwenhoek first observed cilia in protozoa using a light microscope of his own design. He described the cilia as having ‘incredibly thin feet, or little legs, which moved very nimbly’ [1,2]. The cilium is a microtubule-based organelle that protrudes from the surface of the cell. There are many different types of cilia (Fig. 1a, d), but all share basic structural units: i.e. microtubule-based axoneme, a transition zone that controls which proteins can enter and leave the cilium [3,4], ciliary membrane, and a singlet zone that includes the ciliary tip. A basal body, i.e. nine triplet sets of microtubules, behaves as a ‘root’ of an axoneme [5,6], from which the axoneme extends through the ciliary membrane. Some cilia, mainly motile cilia, include a pair of microtubules in their center, which is called a central pair, that are responsible for the reciprocal linear ciliary motion. Fig. 1. View largeDownload slide Structure and functions of cilia. (a) Schematics of a cell with multiple motile cilia beating asymmetrically to generate fluid flow. (b) The major structures of the motile cilium. Motile cilium with a 9 + 2 arrangement of microtubule axoneme. The ciliary motion is generated through the coordinated action of inner and outer dynein arms. (c) The asymmetric beat pattern of cilium propels the fluid in the proper direction by using two distinctive strokes; effective stroke and recovery stroke. The effective stroke is faster than the recovery stroke. (d) Fluorescent microscopic images of a mature primary cilium. Acetylated tubulin (red) is used as a primary cilium marker. The nucleus is stained with DAPI (blue). The lower panel shows a DIC image of the same field of view as the upper panel. Note that is it hard to see the primary cilium without labeling. Scale bar = 10μm. (e) Sensory functions of the primary cilium. Left panel: simplified mechanism of chemosensation. The chemical stimulus is sensed through receptors present on the ciliary membrane. Right panel: simplified mechanism of mechanosensation. The fluid flowing in the external environment exerts a force on the primary cilium which opens ion channels on the primary cilium. (f) Steps of primary cilium assembly and disassembly. Ciliogenesis is induced at the G1/G0 phase of the cell cycle where mother centriole transitions to the basal body. The basal body then interacts with ciliary vesicle or ciliary membrane directly, and assembly of primary cilium starts by elongation of the axoneme. After maturation, the cilium begins to disassemble when a cell starts to divide. Fig. 1. View largeDownload slide Structure and functions of cilia. (a) Schematics of a cell with multiple motile cilia beating asymmetrically to generate fluid flow. (b) The major structures of the motile cilium. Motile cilium with a 9 + 2 arrangement of microtubule axoneme. The ciliary motion is generated through the coordinated action of inner and outer dynein arms. (c) The asymmetric beat pattern of cilium propels the fluid in the proper direction by using two distinctive strokes; effective stroke and recovery stroke. The effective stroke is faster than the recovery stroke. (d) Fluorescent microscopic images of a mature primary cilium. Acetylated tubulin (red) is used as a primary cilium marker. The nucleus is stained with DAPI (blue). The lower panel shows a DIC image of the same field of view as the upper panel. Note that is it hard to see the primary cilium without labeling. Scale bar = 10μm. (e) Sensory functions of the primary cilium. Left panel: simplified mechanism of chemosensation. The chemical stimulus is sensed through receptors present on the ciliary membrane. Right panel: simplified mechanism of mechanosensation. The fluid flowing in the external environment exerts a force on the primary cilium which opens ion channels on the primary cilium. (f) Steps of primary cilium assembly and disassembly. Ciliogenesis is induced at the G1/G0 phase of the cell cycle where mother centriole transitions to the basal body. The basal body then interacts with ciliary vesicle or ciliary membrane directly, and assembly of primary cilium starts by elongation of the axoneme. After maturation, the cilium begins to disassemble when a cell starts to divide. Cilia are present on a variety of cell types; therefore, they have a broad range of functions. Multiple motile cilia underlie the airway’s mucociliary function [7,8], generate cerebrospinal fluid flow in brain ventricles [9,10], and help transport oocytes in the fallopian tube [11] by propelling mucus or extracellular fluid (Fig. 1a). Motile monocilia in the embryonic node play a key role in the determination of the embryonic left–right axis by the generation of extraembryonic fluid flow [12]. Primary cilia have roles in sensing environmental signals, and expressing various receptors, ion channels and signaling molecules on the ciliary membrane [13]. The sensory modalities of cilium include mechanosensation (Fig. 1e; right) [14], chemosensation (Fig. 1e; left) [13], and photoreception in a specialized case [15]. Besides, primary cilia also manage key intracellular signaling processes during development and in tissue homeostasis, such as the sonic hedgehog signaling [16] or planar cell polarity pathways [17]. Defects in motile and/or non-motile cilia cause a wide range of diseases, i.e. the so-called ciliopathies. Dysfunction of ciliary motility results in a motile ciliopathy known as primary ciliary dyskinesia (PCD), which is caused in many cases by the compromised formation or function of the outer dynein arms, dynein regulatory complex or central pair, all of which are required for ciliary motility [18]. Sensory ciliopathies result specifically from defects in the sensory and/or signaling functions of cilia and include polycystic kidney disease, retinal degeneration, congenital heart defects, liver fibrosis, obesity, skeletal malformations and brain anomalies [19]. Motile cilia Motile cilia are 5–10 μm long and have a 9 + 2 axoneme arrangement containing nine interconnecting peripheral pairs of microtubules and a microtubule pair in the center of the axoneme (Fig. 1b) [20]. Each outer doublet is attached to the radial spokes through an A-tubule of outer doublet microtubules while the pair of inner singlet microtubules is connected by paired bridges, which are called the central apparatus [21]. The nine peripheral doublets of microtubules are attached through nexin links (interdoublet links), which keeps the outer doublets in a circular arrangement (Fig. 1b). The dynein motor proteins are arranged into two rows along the length of microtubule doublets: i.e. the inner and outer dynein arms [22]. The ciliary beat is generated by the sliding of the microtubule doublets against the adjacent microtubule doublets through the coordinated action of the inner and outer dynein arms [23–25]. The energy for this process is provided via the hydrolysis of adenosine triphosphate (ATP) by dynein ATPase [26]. The nexin–dynein complex converts the sliding into bending by limiting the sliding motion of the cilium [27,28]. Connections between the ciliary membrane, the axoneme and the radial spokes are also involved in the motion. Each cilium beats at a given tempo, which is known as the ciliary beat frequency (CBF) [29]. The CBF is 15–25 Hz in airway epithelia [30,31]. Cilia provide mucociliary clearance (MCC) through generating mucous flow by their asymmetric beating using two distinctive strokes: i.e. effective and recovery strokes (Fig. 1a, c) [32]. During an effective stroke, cilia engage with the mucous layer to propel the fluid forward; in contrast, during a recovery stroke, the cilia return to the original position in the underlying periciliary fluid against the flow. Effective strokes are faster than recovery strokes and produce a strong force to propel mucus forward (Fig. 1c). The speed of the effective strokes reduces toward the end of the stroke and ciliary motion steadily increases at the beginning of recovery strokes [33]. This asymmetric beating pattern is responsible for the net fluid flow in the direction of the effective stroke (Fig. 1a). There are 200–300 cilia on a ciliated epithelial cell. The cilia on these cells must beat in the same direction to drive the flow [34] and beat in coordinated metachronal waves to propel fluid properly [35]. Respiratory ciliary function abnormalities are associated with various diseases, such as cystic fibrosis, chronic obstructive pulmonary disease and sinusitis. Abnormal ciliary beating and disruption of cilia synchronicity lead to poor MCC as seen in Kartagener syndrome and PCD, respectively, which can result in chronic respiratory infections [36,37]. Although electron microscopic techniques have been useful to elucidate the structure of normal motile cilium and diagnosis of PCD [11,38], it is imperative to understand the ciliary motion for a complete understanding of ciliary function because abnormalities were present in several motile cilia even when the ultrastructure was normal. An example is DNAH11 mutations in PCD; i.e. PCD patients with DNAH11 mutations show irregular ciliary motion and dysfunction in MCC without overt ultrastructural abnormalities [39,40]. Thus, measurement of ciliary action can serve as an important indicator of upper respiratory health. Microscopic analyses of motile cilia motion in the respiratory system CBF is one of the simplest parameters to evaluate motile cilia function. Many microscopic methods have been developed to measure motile cilia motion. Classical approaches to measure CBF used photodiode/photomultiplier detectors [41–43] and high-speed cinematography [44]. Photodiode techniques indirectly estimate the CBF by detecting the changes in the light intensity passing through the cilia. Instead of the direct use of microscopic images, this method relies on reflected or transmitted light to measure CBF, which means the CBF measurements also include external variations [45]. High-speed cinematography techniques record ciliary images and CBF is calculated by analyzing the movies in slow motion to count cilia cycles. At present, CBF can be quantified by automated analysis of microscopic images captured with a high-speed charge-coupled device (CCD) camera because these tools are simpler to use and allow faster analysis immediately after recording [36,46,47]. The microscopic images of ciliary beats are usually processed using the average optical density (OD) for a region of interest versus time [48,49]. CBF is then calculated with either the autocorrelation of OD as a function of time or a fast Fourier transform. Although the measurement of CBF is vital for understanding MCC, CBF alone does not necessarily exhibit motile cilia functions. Indeed, PCD patients with DNAH11 mutations show larger CBFs than healthy subjects [39,40]. Another important parameter for evaluating motile cilia function is the ciliary beat pattern (CBP) or waveform. DNAH11 mutations seem to result in irregular CBP, i.e. a small beating amplitude [39,40]. A mutant of hydin, a component of the central apparatus, shows severe defects in tracheal ciliary motility, which was unable to bend normally [50]. The ciliary beat asymmetry is also an essential parameter of CBP. This asymmetry is crucial for ciliary function because the loss of beat asymmetry in murine tracheal cilia resulted in a reduction of cilia-generated fluid flow in trachea, which resulted in the accumulation of mucus in the nasal cavity [51]. Few studies have analyzed this asymmetry in detail because of the difficulty in tracing ciliary tips without labeling. Another reason hampering analyses of beat asymmetry might be economic; i.e. the fast motion of fluorescence requires an expensive high-speed camera with a high S/N ratio, such as an electron-multiplying CCD (EM-CCD), while fluorescent labeling of ciliary tips enables the automatic visualization of ciliary movement and detection of cilia beating trajectories [33]. Here, we present a simple procedure to record the motion of ciliary tips with a camera one-tenth cheaper than EM-CCD or with a civilian compact camera. Figure 2 shows the ex vivo analysis of ciliary beating in intact cilia on mouse tracheal epithelial cells. First, one-third of the ventral side of the dissected trachea was clipped out and dipped into Indian ink diluted at 1:200 with culture medium (Dulbecco’s modified Eagle medium) for 5–10 min to have carbon deposits attached to the cilia tips, which provide imaging contrast (Fig. 2a). Next, a piece of the trachea is placed in a chamber with the luminal face down in a small volume of culture medium deep enough to cover the tissue (Fig. 2a). However, an excess volume of medium results in the drift of trachea tissue during data acquisition, which hampers the tracking of ciliary motion in later analyses. Ciliary motility was then is recorded from the bottom of the chamber using an inverted microscope equipped with a high-speed camera. The optimal configuration of the lens includes 40× to 60× magnification, 0.5–0.7 of N.A., long working distance, and correction ring, to track ciliary strokes with beating amplitude of several μm on the tracheal lumen, which is 0.5–2 mm far from the chamber bottom. We do not recommend observing ciliary motion using a water dipping lens with an upright microscope by placing tracheal tissue with the luminal face up because the tissue piece drifts easily. Fig. 2. View largeDownload slide Live imaging of motile cilia. (a) A schema of how to prepare tracheal tissue sample, label ciliary tips with Indian ink and observe ciliary motility with a light microscope. (b) A screen capture image demonstrating the measurement of wave pattern from ciliary beating plane using the ImageJ software package. A raster line (yellow) drawn along the beating plane. Scale bar = 5 μm. (c) The extracted kymograph from panel b shows the wave pattern with the recovery phase, effective phase and rest phase of the ciliary beat cycle. (d) Analyses of the asymmetric beating of cilia by manual tracking of ciliary tips using ‘Manual Tracking’ plugin of FIJI and measurement of ciliary beating plane as the linear correlation function of the raw trajectory. Scale bar=5 μm. (e) The plot of the aligned trajectory measured by rotating the original trajectory from panel d. (f) The plot of the ciliary trajectory on the y-axis as a function of time. (g) The plot of the velocity in the y-direction against time. The upper positive values represent effective strokes and lower negative values represent recovery strokes. (h) Schematics of the isolation of tracheal cilia axonemes for in vitro imaging. The isolated axonemes reactivated with ATP and randomly attached to the glass surface. (i) Recorded images of motion of ATP-reactivated ciliary axoneme. (j) Kymograph of ATP-reactivated ciliary axoneme generated from images in panel i. The wave pattern corresponds to the asymmetric beating of tracheal cilia with fast effective strokes (upward) and slow recovery strokes (downward). The original data are obtained from [51]. Fig. 2. View largeDownload slide Live imaging of motile cilia. (a) A schema of how to prepare tracheal tissue sample, label ciliary tips with Indian ink and observe ciliary motility with a light microscope. (b) A screen capture image demonstrating the measurement of wave pattern from ciliary beating plane using the ImageJ software package. A raster line (yellow) drawn along the beating plane. Scale bar = 5 μm. (c) The extracted kymograph from panel b shows the wave pattern with the recovery phase, effective phase and rest phase of the ciliary beat cycle. (d) Analyses of the asymmetric beating of cilia by manual tracking of ciliary tips using ‘Manual Tracking’ plugin of FIJI and measurement of ciliary beating plane as the linear correlation function of the raw trajectory. Scale bar=5 μm. (e) The plot of the aligned trajectory measured by rotating the original trajectory from panel d. (f) The plot of the ciliary trajectory on the y-axis as a function of time. (g) The plot of the velocity in the y-direction against time. The upper positive values represent effective strokes and lower negative values represent recovery strokes. (h) Schematics of the isolation of tracheal cilia axonemes for in vitro imaging. The isolated axonemes reactivated with ATP and randomly attached to the glass surface. (i) Recorded images of motion of ATP-reactivated ciliary axoneme. (j) Kymograph of ATP-reactivated ciliary axoneme generated from images in panel i. The wave pattern corresponds to the asymmetric beating of tracheal cilia with fast effective strokes (upward) and slow recovery strokes (downward). The original data are obtained from [51]. Temperature considerations are important because ciliary activities are tightly correlated with temperature [49,52]. A study in the mouse respiratory epithelium showed that as the temperature declined from 37 to 25°C, the CBF and flow speed decreased, dropping from 18.2 ± 3.8 at 37°C to 12.6 ± 2.3 Hz at 22–24°C [53]. Although an ideal temperature to analyze ciliary motion is 37°C because it is the physiological body temperature; this temperature deteriorates the tissue immediately. Moreover, faster ciliary beating at 37°C demands a more expensive camera with higher frame rates. Therefore, ciliary motions are observed and analyzed at room temperature in several studies published in the literature [33,50]. For the quantitation of ciliary motility, differential interference contrast (DIC) imaged movies were loaded into the ImageJ software package (National Institutes of Health, Bethesda, MA, USA) and used the line tool to draw a raster line along the beating plane (Fig. 2b). A reslice of this line creates a kymograph extracted from a sequence of digital images, where the cilia movement along the line generates a wave pattern (Fig. 2c). The waveform shows the three phases of the beat cycle: i.e. recovery, effective and rest phases (Fig. 2c). To analyze the ciliary motion precisely, especially the beat asymmetry, Indian ink attached to ciliary tips was tracked manually using the ‘Manual Tracking’ plugin from FIJI (Fig. 2d). The beating plane was easily found as the linear correlation function of the raw trajectory (Fig. 2d) and the trajectory is aligned on the y-axis by rotating the original trajectory (Fig. 2e). The ciliary trajectory on the y-axis was plotted as a time function to analyze the beating asymmetry (Fig. 2f). The y-axial trajectory reproduces the kymograph well (compare Fig. 2c and f). The velocity in the y-direction was calculated from the trajectory on the y-axis and plotted against time (Fig. 2g). Figure 2g shows faster effective stokes (upper: positive values) than recovery strokes (lower: negative values). The analysis of ciliary beat asymmetry in the trachea semi-organ culture has a chicken-and-egg problem. That is, directional fluid flow can break the symmetry of ciliary beating hydrodynamically. Therefore, it is difficult to detect the cilia’s intrinsic asymmetric beating pattern. To purely analyze the asymmetric CBP, we developed an in vitro experimental paradigm where de-membraned tracheal cilia axonemes were isolated and reactivated with ATP in a chamber formed between two pieces of glass [51]. Axonemes tend to attach to the glass surface by one end, while the other end shows oscillatory movements when ATP is added (Fig. 2h). The motion of ATP-reactivated ciliary axoneme was recorded using oil or water immersion, high magnification, DIC lens (60× to 100×; N.A. 1.3–1.4), and a high-speed CCD camera (Fig. 2i). Next, kymograph of an ATP-reactivated ciliary axoneme was generated as shown in Fig. 2b (see Fig. 2j). The kymograph shows the clear beating asymmetry of isolated cilia with fast effective strokes and slow recovery strokes (Fig. 2j) as similarly observed in the intact cilia [51]. The ATP-reactivated isolated cilium provides evidence that the asymmetric beating pattern is truly intrinsic to the cilium. CBP analyses can be performed similarly to that done for intact cilia in ex vivo trachea cultures [51]. Primary cilium The primary cilium is a single organelle that protrudes from the surface of the most mammalian cell types during growth arrest [54–56] and serves as cellular antennae/towers to help detect and transduce extracellular signals [19]. The axoneme of primary cilium has almost the same configuration as motile cilia, but a clear difference is that it has the ‘9 + 0’ configuration consisting of a radial array of nine doublet microtubules with no central pair of singlet microtubules and lacks the dynein motors. Most primary cilia are non-motile except for nodal cilia present in the ventral nodes of vertebrates [57]. However, ‘non-motile’ does not mean that the primary cilia are static and do not move at all. The primary cilium grows, elongates, shrinks and disassembles depending upon the cell state (Fig. 1f). The primary cilium is formed as the cell enters the G0 phase and disassembled in cell division (Fig. 1f). Building blocks of the primary cilium, including tubulin, are delivered into the cilium by an intraflagellar transport (IFT) system driven by ATP-dependent molecular motors, kinesin and dynein. Defects in the primary cilia formation lead to more varied sensory, physiological and developmental anomalies than do defects in motile cilia, which cause polycystic kidney diseases, Joubert syndrome, Bardet–Biedl syndrome, cone-rod dystrophy, axial spondylometaphyseal dysplasia and spinocerebellar ataxia [19]. Mutations in the IFT particle protein gene Tg737/Ift88 have been shown to cause abnormal craniofacial and photoreceptor development, retinal degeneration, polycystic kidney disease and human cleft palate [58–60]. Mutations in the lipid-metabolizing enzyme, inositol polyphosphate 5-phosphatase E (INPP5E), creates abnormally shortened cilia, causes ciliary disassembly, and results in ciliopathies, such as Joubert syndrome and mental retardation, truncal obesity, retinal dystrophy and micropenis (MORM) syndrome [61–63]. In Joubert syndrome, point mutations in the catalytic domain of the INPP5E change the phosphatase activity of the enzyme while the MORM syndrome is caused by C-terminal truncating mutations of the INPP5E that affect ciliary localization [61–63]. According to these mutations, primary cilia become unstable in vivo and in vitro [58–60,62,64]. Serum stimulation, which is used to induce cilia disassembly in vitro, makes primary cilia of Inpp5e-KO cells more fragile [61,62,64]. Time-lapse imaging of primary cilium Live cell imaging is a powerful technique to observe the dynamic behavior of primary cilium directly. Neither high-speed nor video-rate recording is required to track the behavior of the primary cilium or trafficking of IFT particles, which contrasts with motile cilia imaging. Time-lapse imaging is suitable for this purpose; thus, time-lapse microscopy uses a method that extends live cell imaging from a single observation in time to the observation of cellular motion over long periods. Thus, time-lapse imaging studies consist of four consecutive steps: i.e. preparation of the cells of interest, observation and recording with an automated microscope with an environmental chamber, a camera integrated with a computer to acquire and store the images, and software to examine the time-lapse images, and measure and analyze the cells. Over the years, many studies have shown time-lapse imaging of cilia formation as sequential still images or video clips [65], IFT in primary cilium [66] and cilia resorption and disassembly upon serum stimulation [61,62,64]. More recently, some groups including us have demonstrated the excision of the tip of primary cilium through time-lapse imaging (Fig. 3a, b) [67–69]. Time-lapse imaging of primary cilium also reveals the mechanism and function of the ciliary tip excision: i.e. accumulation of phosphatidylinositol 4,5-bisphosphate (PI(4, 5)P2) in primary cilium triggers actin polymerization in the distal part of the primary cilium, which results in ciliary tip excision. Thus, the ciliary tip excision provides a cue for cilia disassembly and re-entry of cells from quiescent stage to into the cell cycle [69]. Fig. 3. View largeDownload slide Time-lapse imaging of primary cilium. (a, b) Time-lapse imaging of primary cilium tip excision in NIH3T3 cells expressing chimeras of Htr6 and mCherry florescent protein. Scale bar = 5 μm (a); 2 μm (b). (c) Various positions of the primary cilium. Left panel: cilium is protruding from the apical side of the cell, and projecting vertically into the medium. The excised tip is difficult to be tracked because it flows away into the medium. Middle panel: cilium sandwiched between the cover glass and the base of the cell. The excised tip can be easily tracked (data of panel a and b). Right panel: cilium projecting horizontally. A ciliary tip excised close to the surface of the cells sticks to the cell surface and can be easily tracked (data of MEFs in [69]). Pink: primary cilium; orange: excised ciliary tip; navy: centriole. Fig. 3. View largeDownload slide Time-lapse imaging of primary cilium. (a, b) Time-lapse imaging of primary cilium tip excision in NIH3T3 cells expressing chimeras of Htr6 and mCherry florescent protein. Scale bar = 5 μm (a); 2 μm (b). (c) Various positions of the primary cilium. Left panel: cilium is protruding from the apical side of the cell, and projecting vertically into the medium. The excised tip is difficult to be tracked because it flows away into the medium. Middle panel: cilium sandwiched between the cover glass and the base of the cell. The excised tip can be easily tracked (data of panel a and b). Right panel: cilium projecting horizontally. A ciliary tip excised close to the surface of the cells sticks to the cell surface and can be easily tracked (data of MEFs in [69]). Pink: primary cilium; orange: excised ciliary tip; navy: centriole. Using time-lapse imaging to analyze the dynamic behaviors of primary cilia requires the labeling of primary cilia due to the smallness and the low contrast of primary cilium. However, DIC microscopy hardly works for imaging primary cilium, although it is broadly used to image the motion of unstained organelles as well as living cells (Fig. 1d). The primary cilium is very short and thin at around 1–5 μm in length and 0.2 μm wide [70]. The size of the primary cilium quite smaller than the usual mammalian cell radius (10 μm); therefore, any field of view that contains the cilium also includes a portion of the cell body. The mammalian cell body is filled with refractive organelles that will scatter light and introduce wavefront distortions into the DIC light path, which makes it very hard to see the cilia of mammalian cultured cells using DIC (Fig. 1d). Another limitation of DIC microscopy is that it only shows the position of objects, but does not give information about the size of small objects and cannot be used to calculate the size of small molecular complexes moving in the cilium. Thus, it is necessary to image primary cilium using labels in which ciliary proteins are tagged with a fluorescent protein, which enables them to be detected and tracked over a cell background. Cilia-specific markers are required for the fluorescent time-lapse microscopy. Though fluorescent tubulin makes a primary cilium visible because tubulin occupies more than 50% of ciliary proteins, the ‘background’ fluorescence of tubulin in the cell body masks the signals from primary cilium. Signaling molecules and receptors highly enriched in primary cilia thus work as excellent ciliary markers. ADP-ribosylation factor-like protein 13 B (Arl13b) is a ciliary membrane small GTPase found in cilia [71] and is one of the most widely used markers for primary cilia [71]. Primary ciliary membrane proteins such as somatostatin receptor 3 (SsTR3) [72] and serotonin receptor 6 (5-HT6 or HTR6) [73] can also be used for primary cilium live imaging (Fig. 3a, b). Other markers include adenylyl cyclase type 3 (AC3) [74], inversin [75], polaris [76], Rab8 [77,78] and [79] smoothened. Many cells must have cilia with high-expression of fluorescent-tagged-protein to efficiently observe primary cilia and the ciliary tip excision. It is a lot easier to find cells for imaging when most of the cells in a population are ciliated. For mammalian cells, the key to obtaining a large fraction of ciliated cells is to control the cell cycle state, and most primary cilia can be formed in the G0 phase. [55,80]. For the induction of ciliogenesis, cells are either grown to confluence or serum-starved [81]. Another resolution is generating cell lines expressing stably fluorescent markers. In addition to a conventional approach of random insertion of the expression unit, genome editing technique can be used. One can insert fluorescent protein directly at gene loci of the endogenous ciliary protein with high efficiency by facilitating homology-dependent and -independent DNA repair (HIDR) through clustered regularly interspaced short palindromic repeat (CRISPR) and CRISPR-associated protein 9 (CRISPR/Cas9) system-based approaches. Site-specific double-strand breaks induced by Cas9/sgRNA are repaired through homologous recombination-mediated strand exchange or nonhomologous end joining in HIDR [82–86]. The other can have highly stable knock-in cell lines by inserting the expression unit into ROSA26 locus, which is highly resistant to gene silencing. ROSA26 locus is a safe area; exogenous expression units that inserted at this locus will not affect the expression of other genes [87] and allows locus-specific and copy number-controlled transgene expression. Rosa26 locus is amenable to nuclease-mediated knock-in including CRISPR/Cas9 gene-editing tools. Guide RNA targeting the ROSA26 locus can be co-introduced with Cas9 mRNA and donor DNA sequences of fluorescent proteins [88,89], which enables targeted integration of fluorescent protein transgene with highly stable expression. With these stable cell lines, fluorescent primary cilia can be readily found in the field of view. The positioning of the primary cilium is also a crucial factor to observe the behavior of the primary cilium, IFT trails, and the ciliary tip excision. It is quite hard to observe the whole of the primary cilium in a focal plane when the primary cilium stands vertically on the apical surface of the cell (left panel, Fig. 3c). Observing a primary cilium in a focal plane is essential for recording and analyzing IFT trails because the speed of IFT traffic is 1–2 μm/sec [90]. A preferable condition is a primary cilium protruding horizontally to the cell surface (right panel, Fig. 3c) or on the bottom, i.e. between the cell and glass surface (middle panel, Fig. 3c). Retinal pigment epithelial 1 (RPE1) cells often show primary cilium on the bottom of the cell, while it occasionally happens in NIH3T3 cells. IMCD-3 cells seem to protrude primary cilium on the bottom of the cell when cultured at low density. The excised ciliary tip is harder to be tracked when primary cilia project into the medium because the released cilia tip flows away into ‘free’ environment (left panel, Fig. 3c). Cilia tip severing can be easily observed when cilia are present on the base of the cell, i.e. between the cell and glass surface (middle panel, Fig. 3c). Figure 3a and b shows the time-lapse imaging of ciliary tip excision in NIH3T3 cells expressing chimeras of Htr6 and mCherry florescent protein, where both primary cilium and excised ciliary tip are observed between the cell and the glass. Alternatively, primary ciliary tip excision can be imaged if the cilia protruding on the top surface have the tip close to the cell surface. In the case, released cilia tip which seems to be ‘sticky’ [69], can be observed because it is attached on the cell surface (right panel, Fig. 3c). Several technical hurdles also need to be overcome for time-lapse imaging of primary cilium. Photobleaching and phototoxicity are significant problems in the time-lapse imaging. Excessive illumination bleaches fluorophores which reduces their emission period, and fluorescence excitation can trigger production of free oxygen radicals which damage the cells and affects cell growth. One robust solution to the problem of photobleaching and phototoxicity is to decrease the intensity of the excitation light by just turning down the laser during imaging and compensate by increasing the exposure time and the time interval between images. Using a spinning-disk confocal microscope or total internal reflection (TIRF) microscope can reduce photobleaching and phototoxicity [91–93]. Spinning-disk confocal microscopy has an additional advantage to recording fast events of the primary cilium, especially in those protruding vertically on the cell surface (left panel, Fig. 3c), though it requires a highly expensive high-spec camera with extremely high S/N ratio. TIRF microscopy also has an additional advantage to observing fast motion in primary cilium with little phototoxicity. A disadvantage of TIRF microscopy is that it forces the observation to be restricted to quite a thin plane near the coverslip, which can be used only in cases where primary cilia are present between the cell and the glass surface (middle panel, Fig. 3c) or pressed to cover glass with a specialized chamber [94]. Another major concern during time-lapse imaging is to keep the cells alive over an extended period ranging from minutes to hours. Cell viability can be preserved by achieving and maintaining a suitable environment with optimal and stable temperature [95], humidity and stably buffered cultured medium throughout the experiment [96]. Temperature is the most vital factor to control on the microscopic stage. Optimal temperature is critical for the survival and health of the cells. Mammalian cells are sensitive to temperature, will cease developing at low temperatures, and can die if exposed even briefly to high temperatures (e.g. 39°C–40°C). The primary cilium is sensitive to temperature and resorbs at a temperature of 42°C [97]. Decreased temperature also seems to disassembly or shortening of primary cilium [98]. The other issue is the optical drift in which the relative position of the plane of focus changes because of slight bending of the sample chamber and glass elements of the microscope due to temperature changes [93]. Stage-top incubator and lens heater are used to keep the temperature of both cells and microscope even, although commercially available systems have economic disadvantages because additional costly equipment is needed to maintain stable and optimal environmental conditions for cell growth, minimize changes in osmolarity and reduce exposure to light to decrease phototoxicity [99,100]. The expression level of fluorescent ciliary markers must be considered. A significant concern of overexpressing the tagged proteins is the effects of overexpression of protein on its localization, motion and subcellular trafficking. Mammalian cells overexpressing the ciliary membrane protein Arl13b showed a marked increase in ciliary length [101]. In another study, overexpression of Sstr3 in cilia affected the normal distribution of ciliary proteins and cilia morphology [102]. Also, overexpression of fluorescent-tagged proteins creates a high cellular background, which makes it difficult to differentiate primary cilium from cell body in wide-field fluorescence microscopy. The problem can be alleviated by taking care to study cells expressing low to medium levels of protein expression. The mild or weak promoter can be used [103]. Depending on the experiment under investigation, the staining of live cilia, such as with fluorescently tagged lipids [104], and lectin [105] staining is a good alternative [106]. Knock-in of fluorescent protein into genomic allele of ciliary markers is a state-of-art technique providing expression of the fluorescent ciliary marker at the endogenous level [69]. Prospects Imaging of ciliary dynamic behaviors is currently two-dimensional in most research studies. In our paradigm, the trajectories observed from the top, i.e. tracheal lumen, are just projected to the 2-D plane. It does not show the ‘true’ motion of 3-D moving motile cilia. Recently, a study has reported strategy of 3-D recording and analyses of motile tracheal cilia [107]. The time-lapse imaging of primary cilium is also limited in a focal plane. 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TI - Live cell imaging of dynamic behaviors of motile cilia and primary cilium
JF - Microscopy
DO - 10.1093/jmicro/dfy147
DA - 2019-04-01
UR - https://www.deepdyve.com/lp/oxford-university-press/live-cell-imaging-of-dynamic-behaviors-of-motile-cilia-and-primary-OkfU0GSTVD
SP - 99
VL - 68
IS - 2
DP - DeepDyve
ER -