Analysis of Ciliary Assembly and Function in Planaria

CHAPTER TWELVE

Analysis of Ciliary Assembly and Function in Planaria Panteleimon Rompolas*,1, Juliette Azimzadeh†, Wallace F. Marshall†, Stephen M. King‡

*Department of Genetics, Yale Stem Cell Center, Yale University School of Medicine, New Haven, Connecticut, USA † Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, California, USA ‡ Department of Molecular, Microbial and Structural Biology, University of Connecticut Health Center, Farmington, Connecticut, USA 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Maintaining Planarians in the Lab 2.1 Culture conditions 2.2 Feeding 2.3 Colony expansion 3. Genetic Manipulation of Planaria 3.1 RNAi via bacterial feeding 3.2 RNAi via microinjection of dsRNA 4. Imaging Planarian Cilia 4.1 Live video microscopy 4.2 Whole mount immunofluorescence 4.3 Transmission electron microscopy 4.4 Scanning electron microscopy 5. Planaria Gliding and Ciliary Motility Assays 5.1 Measuring planarian locomotion 5.2 Measuring ciliary beat frequency 6. Summary References

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Abstract Planarians are free-living invertebrates that employ motile cilia for locomotion. Specifically, cilia that populate the ventral epithelium of the planarian body are highly conserved, with a 9 þ 2 axoneme and a full complement of inner and outer arm dynein motors. The abundance of cilia on the planarian body, their unique accessibility, and high degree of conservation make this organism an attractive experimental model system for cilia biology. Moreover, planarians are genetically amenable and defects that compromise the function and structure of the cilia are not detrimental for their overall Methods in Enzymology, Volume 525 ISSN 0076-6879 http://dx.doi.org/10.1016/B978-0-12-397944-5.00012-2

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2013 Elsevier Inc. All rights reserved.

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health, making them an ideal system for cilia gene loss-of-function studies. In this chapter, we provide information for introducing and maintaining planarians for experimental purposes in the laboratory and describe protocols for RNAi-induced gene knockdown studies. Furthermore, we elaborate on different imaging techniques used to analyze cilia physiology and structure, including live video microscopy, immunofluorescence analysis, and electron microscopy. Last, we provide assays for evaluating physical parameters of ciliary motility, including quantification of planarian gliding locomotion and measurement of ciliary beat frequency.

1. INTRODUCTION Planarians are free-living, nonparasitic metazoans of the phylum Platyhelminthes (class: Turbellaria). A typical feature of planarians is their enormous capacity to regenerate large portions of their body after amputation or injury (Morgan, 1898; Sa´nchez Alvarado, 2003). This is due to the action of stem cells, or neoblasts, which are totipotent mesenchymal cells and the only mitotically active cells in asexual planarians (Wagner, Wang, & Reddien, 2011), ultimately giving rise to all the differentiated tissues and organs. Thus, neoblasts present a great potential for studying the genetic and molecular mechanisms of tissue regeneration. This has lead to the recent adoption of Schmidtea mediterranea as a major model organism for such studies (reviewed in Newmark & Sa´nchez Alvarado, 2002; Reddien & Sa´nchez Alvarado, 2004; Salo´, 2006) with the subsequent development of a sequenced genome and genetic tools for the identification and manipulation of planarian genes (Abril et al., 2010; Gonza´lez-Este´vez, Momose, Gehring, & Salo´, 2003; Robb, Ross, & Sa´nchez Alvarado, 2008; Sa´nchez Alvarado & Newmark, 1999). Planarians, like other invertebrates, have developed a characteristic mode of locomotion based on the concerted action of motile cilia that line the ventral epithelium of the planarian body and a muscular network (Reddien, Bermange, Murfitt, Jennings, & Sa´nchez Alvarado, 2005). Specifically, the planarian ventral epidermis consists of a monostratified multiciliated epithelium (Rompolas, Patel-King, & King, 2010; Smales & Blankespoor, 1978). These cilia are motile with typical dynein-powered 9 þ 2 microtubule doublet axonemes and characteristic synchronized beating, also found in other multiciliated epithelia, like the respiratory tract in mammals. These properties, in addition to established genetic and molecular tools, make planarians and S. mediterranea a particularly useful model organism for the study of cilia. In fact, in recent years an increasing number of investigators have made significant discoveries on cilia biology using

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planarians as a major experimental tool, uncovering the link between Hedgehog signaling genes and cilia biogenesis (Glazer et al., 2010; Rink, Gurley, Elliott, & Alvarado, 2009), the role of axonemal dynein subunits in synchronized ciliary beating (Rompolas et al., 2010), the role of Wnt signaling genes in the apical positioning of cilia (Almuedo-Castillo, Salo´, & Adell, 2011), and deciphering the molecular architecture and evolution of centrosomes and centrioles (Azimzadeh, Wong, Downhour, Sa´nchez Alvarado, & Marshall, 2012). In this chapter, we provide general protocols for using S. mediterranea as a model experimental organism in the laboratory to study the biology of cilia. These include methods for gene loss-of-function studies via RNAi, imaging techniques to visualize planarian cilia, and functional assays to analyze ciliary motility and cilia-based locomotion.

2. MAINTAINING PLANARIANS IN THE LAB Planarians are normally found living in freshwater ponds and springs. Using a pair of photoreceptors, planarians are able to detect a light source and normally swim away from it as a protective mechanism. In order to maintain a healthy population of planarians for experimental use in the laboratory, it is imperative to provide conditions that simulate best their normal habitat, including temperature, light conditions, culture water, etc.

2.1. Culture conditions 1. Planarians of the species S. mediterranea are maintained submerged in 1  solution of Montjuı¨ch salts (subsequently termed planaria medium: 1.6 mM NaCl, 1.0 mM CaCl2, 1.0 mM MgSO4, 0.1 mM MgCl2, 0.1 mM KCl, 1.2 mM NaHCO3) (Cebria´ & Newmark, 2005). 2. Prepare a large batch of 5  stock planaria medium and keep at 4  C for up to a year. Slight precipitation of the 5  stock is normal, however, if larger precipitates appear or the solution becomes cloudy make fresh. 3. Dilute 5 stock with deionized water to make 1  planaria medium. 4. Use plastic, food grade, wide containers with a flat bottom and removable lid to house planarians. 5. Use a mild detergent to wash the container and rinse thoroughly with deionized water to remove any soap residue. 6. Fill one-third the volume of the container with planaria medium. 7. Use plastic transfer pipettes to transfer planarians between containers. Planarians have flexible bodies and are able to squeeze through narrow

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spaces without injury; however, for larger animals, clip the tip of the pipette to widen the opening. Keep an appropriate number of animals per container by monitoring the population for stress (see below) due to overcrowding. Keep the lid of the container half ajar at all times to allow for exchange of gases and normal oxygenation of the medium. Put containers in a dark place and maintain a steady temperature between 18 and 20  C. Ideally, a temperature-controlled incubator with sufficient airflow is most appropriate to maintain stable culture conditions for a large population of planarians in the laboratory. Monitor the health of planaria frequently and observe for changes in behavior and culture conditions. Signs of stress include limited planarian mobility, firm attachment to the substrate, increased mucus secretion, loss of appetite, appearance of lesions in the planarian body, foul odor, increase of waste by-products, and cloudy water. If any of the above symptoms appear, quarantine the container, decrease the population density, and change the medium frequently until the population recovers. Do not feed the planarians during recovery. It is possible that some of the stress symptoms may be due to bacterial infection, which can be treated with the addition of gentamicin (50 mg/mL) in the medium.

2.2. Feeding Planarians are carnivores, feeding off dead animals when in the wild. As opposed to other organisms, planarians adapt to starvation by gradually decreasing the number of cells in their body and therefore their overall size. Conversely, when planarians are fed frequently, they can increase severalfold in size in a matter of a few weeks. 1. Feed planarians with good quality organic calf liver. 2. Remove the liver from the container and rinse thoroughly with tap water. 3. Use scissors to remove all the connective tissue. 4. Cut liver in small pieces and homogenize to a fine paste using a commercial grade blender. 5. Pass the liver homogenate through a fine mesh to remove any remaining fibers. 6. Aliquot liver homogenate in 15-mL Falcon tubes and store at 20  C for up to 3 months or 80  C for 1 year.

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7. Feed planarians once a week to maintain a steady population or every 2 days if the aim is to induce growth and expand the colony. 8. Thaw the liver homogenate on the bench until it reaches room temperature. 9. Use a plastic transfer pipette to aspirate a small volume of liver. 10. With the tip of the pipette submerged into the medium, place the liver into the container making sure the tip of the pipette remains always in contact with the bottom. This is to keep the liver homogenate in a concentrated single mass and avoid dispersion into the medium. 11. Cover the container to keep it dark and allow planarians to feed for 1 h. 12. Planarians that have finished feeding appear lighter in color due to the ingested liver and swim away from the food. 13. Remove big liver chunks with a transfer pipette and then carefully pour out the soiled medium. Planarians usually stay attached to the container and will not come off while removing the medium. 14. Use a transfer pipette to squirt fresh medium over the planarians to help them detach and gather them in one corner keeping the container in a slight angle. 15. Using a dry Kimwipe, clean the debri and mucus from the bottom of the container. 16. Rinse once and fill one-third of the container with fresh medium. 17. Repeat the cleaning regimen after every feeding or at least once a week.

2.3. Colony expansion There are two strains of the planarian S. mediterranea, which can reproduce sexually or asexually, respectively. The divergence of these two stains was the result of a chromosome translocation (De Vries, Baguna, & Ball, 1984; Salo´ & Baguna, 1985). Sexual reproduction involves the mating between two hermaphrodites and the subsequent deposition of fertilized eggs (Zayas et al., 2005). Eggs appear as small spheres of 1 mm diameter, having a hard pigmented shell and attached to the substrate through a thin thread. If left alone, eggs will hatch after approximately 3 weeks and young juvenile planarians will appear. Juveniles are almost colorless, and because of their small size, they can be easily missed and thrown away during cleaning. Therefore, it is advisable that unhatched eggs be carefully transferred to a smaller container separate from the adult population so that their progress can be monitored more carefully. Mating and laying of eggs in the sexual strain of S. mediterranea shows seasonal periodicity and occurs more

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frequently when the planarian population is under stress (Hoshi, Kobayashi, Arioka, Hase, & Matsumoto, 2003). The asexual strain of S. mediterranea will propagate by spontaneous fission, where their body separates close to the midline giving off a head and a tailpiece. Both body segments will create a blastema at the site of fission, eventually regenerating the missing part (Reddien & Sa´nchez Alvarado, 2004). The end result is two identical planarians which can then grow and repeat this cycle. Although both modes of planarian reproduction are possible in the laboratory, a more practical and efficient way to expand a colony is by utilizing the ability of planarians to fully regenerate after amputation. 1. Feed planarians every 2 days with liver homogenate until they reach a size of 1 in. 2. Stop the feeding 3 days before amputation to allow time for full digestion. 3. Using a plastic transfer pipette, place one planarian flatworm in a petri dish with fresh medium. 4. Use a scalpel and cut 1–2 clean crosscuts separating the body into 2–3 pieces. 5. Transfer the amputated planarian body segments into a new container with fresh medium and separate from the rest of the adult population. 6. Repeat steps 3–5 as required. 7. Place the container with amputated planarians in the dark incubator and allow 2–3 weeks to fully regenerate. 8. During this period, change the medium regularly to remove debris from the regeneration process. 9. Do not feed planarians until they have fully regenerated.

3. GENETIC MANIPULATION OF PLANARIA The S. mediterranea (taxid: 79327) genome has been recently sequenced and Blast searches of target genes are available through the National Center for Biotechnology Information Web site (http://blast.ncbi.nlm.nih.gov/) or alternatively through the S. mediterranea genome database (SmedGD; Robb et al., 2008). Name designation of S. mediterranea homologous genes follows the convention proposed by Reddien, Newmark, and Sa´nchez Alvarado (2008). Gene expression can be easily manipulated in S. mediterranea by means of dsRNA-mediated RNAi (Fire et al., 1998; Sa´nchez Alvarado & Newmark, 1999; Timmons, Court, & Fire, 2001). Here, we provide two

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alternative protocols for loss-of-function experiments in S. mediterranea that employ bacterial feeding or microinjection, respectively, for the delivery of dsRNAs (Fig. 12.1A and B).

A T7

cDNA

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cDNA

T7

L4440

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dsRNA

L4440 E. coli

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Figure 12.1 RNAi-mediated gene knockdown in planarians. (A) RNAi via bacterial feeding. A cDNA sequence of the gene of interest is inserted between the two opposing T7 RNA polymerase promoters in the pL4440 plasmid, which is then used to transform E. coli cells. The bacterial culture is induced in order for dsRNA synthesis to begin, and after induction, the bacteria are harvested and mixed with homogenized calf liver that is used to feed the flatworms. Food coloring is also added to the food mix to monitor the course of the feeding. As the bacteria in the food are digested, dsRNA is released and absorbed by cells throughout the body. This regimen is then repeated every 3 days or until a phenotype starts to appear. (B) RNAi via dsRNA microinjection. Scheme showing the position of the planarian gut. For dsRNA injection into the gut (1), the tip of the micropipette is positioned between the photoreceptors and the pharynx region, along the midline. Following injection, the flatworms are amputated pre- and postpharyngeally (2) and allowed to regenerate.

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3.1. RNAi via bacterial feeding 1. For a given gene target, a cDNA fragment of 200–600 bp is required. 2. Subclone the cDNA fragment into the pL4440 vector (Timmons & Fire, 1998) using the multiple cloning site between the two opposing T7 promoters. 3. Transform the resulting plasmid into HT115 (DE3) E. coli competent cells, which are deficient for RNaseIII activity preventing the degradation of the dsRNA (Timmons et al., 2001). 4. Select positive clones on LB plates supplemented with 50 mg/mL ampicillin and 15 mg/mL tetracycline. 5. Inoculate with freshly transformed HT115 cells a 4 mL starting culture of 2  YT medium (16 g tryptone, 10 g yeast extract, and 5 g NaCl per L) supplemented with 100 mg/mL ampicillin and 50 mg/mL tetracycline and grow in a shaker at 37  C for 16 h. 6. Dilute the starter culture 1:10 in a fresh prewarmed 2  YT medium supplemented with 100 mg/mL ampicillin and let the cells grow to OD595 ¼ 0.4. 7. Induce expression of dsRNA with 1 mM IPTG for 2–3 h. 8. Pellet bacteria corresponding to 2 mL of the induced culture using a tabletop microfuge. 9. Mix the bacterial pellet with 50 mL of liver homogenate (see Section 2.2) and add 1 mL of red food dye to monitor the uptake. 10. Fill a petri dish halfway with planaria medium and transfer a group of no more than 10 medium-sized (1 cm) flatworms. 11. Carefully place the food mix at the bottom of the petri dish using a 200 mL pipette tip, making sure that it touches the substrate so that it does not disperse. 12. Let planaria feed for at least 1 h and monitor uptake by looking for a reddish hue in their body color due to the food dye. 13. Repeat steps 5–13 every 3 days for a total of four feedings. Different genes may require additional feedings for effective knockdown. 14. Evaluate mRNA levels of the target genes after RNAi by qRT-PCR, northern blotting following standard protocols, or alternatively by in situ hybridization as described in Pearson et al. (2009).

3.2. RNAi via microinjection of dsRNA 1. Injection of dsRNA usually produces higher levels of gene inactivation than feeding. 2. For a given gene target, a cDNA fragment of at least 0.3 kb is required.

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3. Subclone the cDNA fragment into pL4440 (Timmons & Fire, 1998), pR244 (gateway), or pPR-T4P (LIC cloning) (Reddien et al., 2005). 4. Amplify by PCR both strands of the cDNA in two separate reactions including a single T7 promoter at the 50 -end of each strand. When using pR244 or pPR-T4P, the following primers can be used: Sense strand Forward: CCA CCG GTT CCA TGG CTA GC Reverse: GAG GCC CCA AGG GGT TAT GTG Antisense strand Forward: AAC CCC TCA AGA CCC GTT TAG A Reverse: GAA TTG GGT ACC GGG CCC Alternatively, a T7 promoter sequence can be added directly to the cDNA sequence: Sense strand Forward: TAA TAC GAC TCA CTA TAG G—16–20 cDNAspecific bp Reverse: 20–25 cDNA-specific bp Antisense strand Forward: 20–25 cDNA-specific bp Reverse: TAA TAC GAC TCA CTA TAG G—16–20 cDNAspecific bp 5. To remove contaminant RNase, add proteinase K and SDS to the PCRs to a final concentration of 0.5 mg/mL and 0.5%, respectively, and incubate 30 min at 37  C. 6. Extract with phenol:chloroform:isoamyl alcohol (25:24:1) and precipitate with ethanol. Dissolve the DNA pellet in RNase-free water. 7. Synthesize dsRNA using the T7 RiboMAX (Promega) or a similar kit. Use 1 mg of template DNA per reaction. Dissolve the RNA pellet in 50–100 mL RNase-free water. 8. Using a Nanoject Auto-Nanoliter II injector (Drummond Scientific) and a glass micropipette, inject 100–150 nL (three to five times 32 nL at slow speed) into the gut of 0.5- to 1-cm animals. Glass micropipettes are obtained using a micropipette puller (Sutter Instrument) and breaking the micropipette tip with fine tweezers to increase the tip diameter. 9. To inject into the gut, the micropipette tip is placed in the area between the photoreceptors and the pharynx along the midline (Fig. 12.1B). To monitor the injection, a food dye such as Allura red can be added to the dsRNA.

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10. Injection is repeated 3 consecutive days to obtain maximal levels of inactivation. 11. The day after the last injection, planarians are amputated pre- and postpharyngeally to trigger regeneration and thus accelerate the appearance of the phenotype. 12. Higher levels of inactivation can be obtained by injecting trunk fragments resulting from the first round of injection/amputation, 6 days after amputation, with 40–70 nL dsRNA (three to five times 13.6 nL at slow speed). 13. A second amputation is performed 7 days after the first one and phenotypes are recorded on the head and tail fragments resulting from the second amputation.

4. IMAGING PLANARIAN CILIA One of the great advantages that planaria present as experimental models, compared to other multicellular organisms, is the fact that the cilia line an external epithelium and are therefore easily accessible and can be visualized noninvasively throughout the planarian life cycle (Fig. 12.2).

4.1. Live video microscopy Planarians employ cilia to propel their body forward and muscle action to negotiate turns and make changes in the direction of movement (Nishimura et al., 2007; Rompolas et al., 2010). In order to visualize and study the beating of planaria cilia via live imaging techniques, it is crucial to immobilize the flatworms so that cilia remain on the same plane of view. Unfortunately, to date, available methods for inhibiting the gross movement of planarians, including cold temperature, genetic manipulation (Nogi, Zhang, Chan, & Marchant, 2009), or treatment with low concentration of ethanol (Stevenson & Beane, 2010), are insufficient or cause adverse effects to the cilia. Here, we present a protocol for live video microscopy of planarian cilia under physiological conditions. 1. Select flatworms that are relatively small in size (1–3 mm). The small size makes it easier to confine the animal and limit the effects of muscular activity. 2. Use a one-hole puncher available in office supply stores to punch a hole in a 1  1 cm square of parafilm. Place the parafilm spacer on a precleaned microscope slide and press on the borders to secure in place.

A 1 mm

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100 µm

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Ventral 10 µm

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Head

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Basal membrane 100 nm

Figure 12.2 Visualizing planarian cilia. (A) Picture of the planarian S. mediterranea, acquired with a digital camera fitted with a 60-mm macro lens. (B) Low and high magnification cross-section images of paraffin-embedded planarian tissue counterstained with hematoxylin and eosin. (C) Top and side views of planarian ventral epidermis. Cilia are stained with anti-a-tubulin antibody (clone B-5-1-2) and visualized using confocal microscopy. (D) Low and high magnification scanning electron micrographs showing the abundance of cilia in the planarian head epidermis. (E) The upper panel shows a low magnification transmission electron micrograph of planarian cilia in the pharyngeal epidermis prepared using the fixation procedure described here. The lower panel shows a higher magnification view of a single cilium prepared using an alternate fixation protocol (Rompolas et al., 2010) that also results in excellent preservation of the 9 þ 2 microtubular axoneme and associated substructures such as outer and inner dynein arms. Panels B, C, and D are adapted from Rompolas et al. (2010).

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3. Place a single drop of planaria medium along with the flatworm to be studied on the slide at the center of the parafilm hole. 4. Carefully place a coverslip on top and gradually press to remove the extra fluid and flatten the body of the animal. The width of the parafilm spacer will allow the immobilization of the planarian without causing injury. 5. Use an upright microscope (Olympus BX51 or equivalent) equipped with differential interference contrast (DIC) optics and coupled with a high frame-rate CMOS camera (X-PRI F1, AOS Technologies; or equivalent). 6. Under 60 magnification, focus on the lateral sides of the planarian head where the cilia are most easily visible. 7. Adjust the condenser, focus accordingly, and open the condenser and field diaphragms for Koehler illumination. 8. Orient the tissue so that the vertical axis of the cilia is parallel to the DIC shear axis for maximum contrast, by rotating the microscope stage. 9. Rotate the camera so that the epithelium is parallel and the cilia vertical, respectively, to the field of view in the preview monitor. 10. Select a frame rate of 250 frames/s. 11. Select the length of the video to be captured. We routinely acquire for 4 s. 12. Start acquisition. The software then allocates a frame buffer and starts collecting images. 13. Save file in an uncompressed AVI video format. 14. Preview the acquired video and verify that the cilia are intact and beating in synchrony. If the cilia look disorganized or deformed, it may be due to excessive force created by the coverslip. If the tissue is shifting, then not enough pressure is exercised. Repeat steps 1–6 with a different animal. 15. Use a video editing software application (VirtualDub, http://www. virtualdub.org; or equivalent) to edit the video file and crop a segment where the cilia beat consistently and in the same plane without tissue shifts. 16. Compress the final video file using any of the free available compression algorithms (H.264 or equivalent) for portability.

4.2. Whole mount immunofluorescence 1. Fix planarians by immersing them live in 1% HNO3, 0.85% formaldehyde, and 50 mM MgSO4 (Dawar, 1973) at room temperature. In all subsequent steps, planarians are kept suspended in solution.

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2. Keep in fixative for 24 h to fully dissolve the mucus and allow the fixation to reach completion. 3. Replace fixative with PBS, pH 7.2 and wash three times. 4. Postfix with freshly made 4% paraformaldehyde in PBS for 10 min on ice. 5. Wash three times with PBS. 6. Permeabilize with 1% Igepal CA-630 (Sigma, St. Louis, MO) in PBS for 10 min. 7. Block with 3% normal goat serum, 1% BSA, 1% cold-water-fish gelatin, 0.1% Igepal CA-630, and 0.05% Tween-20 in PBS. 8. Incubate the primary antibody diluted in 1% BSA, 0.1% cold-water-fish gelatin, and 0.05% Tween-20 in PBS for 16 h at 4  C. 9. Wash three times with PBS. 10. Incubate the fluorescently labeled secondary antibody diluted in 1% BSA, 0.1% cold-water-fish gelatin, and 0.05% Tween-20 in PBS for 1 h at room temperature. 11. Wash three times with PBS. 12. Mount planarians whole with their ventral surface facing upward on a glass slide using a glycerol-based mounting medium and silicone spacers between the slide and the coverslip. 13. Image cilia on the ventral epidermis using a laser scanning confocal microscope. After fixation (steps 1–5), planarians can also be embedded in paraffin or frozen in OCT medium and sectioned for immunohistochemistry or immunofluorescence analysis following standard protocols (Fig. 12.2B and C).

4.3. Transmission electron microscopy Ultrastructural analysis can be performed on cilia covering the ventral epidermis of planarians. In addition, motile cilia are present at a very high density at the surface of the pharynx, the feeding organ of planarians (Fig. 12.2E). 1. The presence of a thick mucus layer at the surface of the planarian epidermis affects the efficiency of both electron microscopy fixation and embedding. Mucus removal can be achieved using N-acetyl cysteine, but this treatment affects the ultrastructure of epidermal cells. We thus recommend the use of microwave processing as described here. 2. Cut the planarians in half or smaller fragments in fixation solution containing 2% glutaraldehyde and 4% paraformaldehyde in 100 mM

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sodium cacodylate, pH 7.2. Fix in a Pelco BioWave microwave tissue processor (Ted Pella, Inc.) at 150 W for 1 min with vacuum at 20 inches Hg, then 1 min with vacuum only. Repeat three times. Remove the fixative and exchange with 3% glutaraldehyde in 100 mM sodium cacodylate, pH 7.2. Incubate O/N at 4  C, then wash three times in 100 mM sodium cacodylate, pH 7.2, for 10 min each. Stain with 1% osmium tetroxide in 100 mM sodium cacodylate, pH 7.2, in the microwave tissue processor during 1 min with vacuum at 20 Hg, then 1 min with vacuum only. Repeat twice. Change to 350 W and fix during 30 s with vacuum at 20 inches Hg, then 30 s with vacuum only. Repeat three times. Protect from light and incubate at room temperature for 1 h. Wash the samples twice in 100 mM sodium cacodylate, pH 7.2 for 10 min each, then three times in water. En bloc stain the samples with 0.5% uranyl acetate in water overnight at 4  C. Filter the uranyl acetate solution through a 0.22-mm filter unit prior to use. Wash the samples twice in water during 10 min each. Perform low temperature dehydration with ethanol series. Incubate the samples in the following ethanol solution for 10 min each: 35% ethanol at 4  C, 50% ethanol on ice, 75% ethanol at 20  C, 80% ethanol at 20  C, 95% ethanol on dry ice, and 100% ethanol on dry ice—twice for the last step. Change to 100% acetone and incubate 10 min at room temperature. For embedding, we use an Epon/Araldite resin mixture. Resin infiltration is performed in the microwave tissue processor at 250 W during 3 min with vacuum at 20 inches Hg. Increasing concentrations of Epon/Araldite resin (without accelerator) diluted in acetone are used: 10%, 25%, 40%, 60% (2), 80% (2), and 100% (3 ) resin, respectively, and in sequence. Change to new 100% resin without accelerator and incubate overnight, rocking. Change to Epon/Araldite resin with accelerator and leave samples rocking for 4 h to infiltrate. Embed in fresh accelerated resin and cure at 60  C for at least 48 h.

4.4. Scanning electron microscopy 1. Planarians are first immersed live in relaxant fixative (1% HNO3, 0.85% formaldehyde, 50 mM MgSO4; Dawar, 1973) for 5 min at room temperature. 2. The fixative is replaced once with fresh and then the samples are left at room temperature for 16–24 h during which time the mucus that can hinder the visualization of the cilia should be completely removed.

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3. Replace fixative with two changes of 2.5% glutaraldehyde in 0.1 M sodium cacodylate, pH 7.4 (EM Sciences) and fix samples for 16–24 h at 4  C. 4. Wash samples with four changes of 0.1 M sodium cacodylate, pH 7.4. 5. Stain with 1% osmium tetroxide for 1 h in the dark, at room temperature. 6. Wash samples with four changes of 0.1 M sodium cacodylate, pH 7.4 and dehydrate with a series of 10%, 25%, 50%, 70%, 85%, 95%, and 100% ethanol solutions, respectively. 7. Dry dehydrated samples at the critical point (Autosamdri-815, Series A or equivalent) and mount with carbon tape and colloidal silver paint. 8. Sputter-coat samples (Cressington 208 HR Sputter Coater or equivalent). 9. Image planarian cilia with a scanning electron microscope (FEI Quanta 200 FEG SEM or equivalent) (Fig. 12.2D).

5. PLANARIA GLIDING AND CILIARY MOTILITY ASSAYS Planarians use their cilia to power their characteristic gliding motion (Glazer et al., 2010; Rink et al., 2009; Rompolas et al., 2010). Therefore, mutations that affect ciliary function and/or structure impact the overall motility of the organism. Thus, parameters of planarian locomotion, like gliding velocity, can be good measures of ciliary function.

5.1. Measuring planarian locomotion 1. Place a group of 10 medium-sized planarians (1 cm) in a petri dish with planaria medium. 2. Use a digital video camera fitted with a macro lens (MiniVID; LW Scientific or equivalent) and mounted on a camera copy stand so that the camera sits vertically on top of the petri dish. 3. Using the camera’s associated software, choose the “live preview” function. 4. Adjust the height of the camera on the copy stand so that the diameter of the petri dish fills exactly the field of view. Mark this height and keep constant for consistency among measurements. 5. Adjust the focus ring on the lens so that the planarians appear in focus on the preview. 6. Adjust the overhead lighting accordingly so that the petri dish is illuminated evenly.

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7. Allow the flatworms to acclimate in these conditions for 1 h. 8. Planarians will usually remain stationary in the absence of any stimulus; therefore, before acquisition use a plastic transfer pipette to gently stir the water in the petri dish until all the flatworms start to move. 9. Immediately commence video acquisition. 10. Acquire 60-s video segments in 15 frames/s in uncompressed AVI format (Fig. 12.3). 11. Open AVI video file in ImageJ (NIH Image). Check the “Convert to Greyscale” option from the popup menu and click OK. 12. With the image stack of the video frames open in ImageJ, go to the main menu and choose “Image > Stacks > Z Project. . ..” 13. In the popup window, choose “Min Intensity” in the pull-down menu and press OK. This allows visualizing the tracks of every flatworm in the petri dish during the 60-s video time window (Fig. 12.3, right panels). t = 60 s

t-projection

Smed-ift88(RNAi)

Wild type

t=0

Figure 12.3 Assay for measuring planarian locomotion. A group of planarians are placed in a petri dish and imaged live, continuously for 60 s. The left and middle panels show the first and last frames of such recordings from wild type (top panel) and planaria who lack cilia (Smedift88(RNAi), bottom panel; Rompolas et al., 2010). Projection of all the time frames on a single image reveals the tracks of individual planaria during the 60-s movie, which allows the precise measurement of their respective gliding velocity. In this example, the velocity of wild-type animals was  1.47 mm/s compared to Smed-ift88(RNAi) planarians which moved considerably slower ( 0.46 mm/s) due to the lack of normal cilia (see Rompolas et al., 2010).

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14. From the main menu go to “Image > Properties” and enter the appropriate values for “Unit of Length” (e.g., millimeters), “Pixel Width,” and “Pixel Height.” To determine these values, use a ruler in place of the petri dish to acquire a test video and divide the actual dimensions of the field of view by the pixel resolution in the x (width) and y (height) axes. 15. Using the “freehand line drawing tool” from the ImageJ Toolbox window, trace the distance that each flatworm traveled in the projected image (Fig. 12.3, right panels). 16. From the main menu go to “Analyze > Measure.” A popup table will appear with the length of the measurement. Repeat steps 15 and 16 for each flatworm. 17. To calculate the actual distance traveled, the length of the body of each flatworm needs to be subtracted from the values measured in step 16. To do that, reopen the AVI video file (step 11) but now chose only the first frame from the popup menu. Repeat steps 14–16 to measure the body length of each flatworm. Subtract the measured values of step 17 from those of step 16, this is the velocity in millimeters per second.

5.2. Measuring ciliary beat frequency Beat frequency of planarian cilia can be measured by kymographs produced from live video microscopy described in Section 4 and similar to the method that has been described for Chlamydomonas flagella (Dentler, Vanderwaal, & Porter, 2009). 1. Follow the protocol in Section 4.1 to acquire a 1-s video, at 250 frames/s, of live planarian cilia beating freely and consistently. 2. Open the uncompressed AVI video file in ImageJ. 3. Check the “Convert to Greyscale” option from the popup menu and click OK. 4. By moving the slider at the bottom of the image stack window left and right make sure that the cilia are moving freely and there are no tissue movements. 5. The epithelium should be oriented horizontally and the cilia vertically to the image window, respectively. 6. If needed adjust the orientation of the tissue by choosing from ImageJ main menu “Image > Transform > Rotate. . ..” 7. Enter the appropriate rotation angle in the popup window, choose “Bicubic” interpolation in the pull-down menu and press OK.

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A

B 20 µm

C y

250 frames = 1 s

Figure 12.4 Assay for measuring ciliary beat frequency in planarians. (A) A single frame from a 60-s movie showing the ventral ciliated epithelium of a planarian imaged live by highspeed video microscopy, at 250 frames/s, using DIC optics. (B) A rectangular area at the interface between cilia and epidermis is cropped and aligned so that the epithelium is oriented horizontally and the cilia vertically to the image window, respectively. (C) A kymograph is produced from the edited decompiled video, illustrating  24 successive ciliary beat cycles.

8. Use the rectangular selection tool from ImageJ toolbox window and select a small area containing the entire length of a small group of cilia, cropping as close as possible to the epithelium (Fig. 12.4A and B). 9. Adjust the contrast appropriately by choosing “Image > Adjust > Brightness/Contrast. . .” and clicking the “Auto,” to make the cilia better stand out from background. 10. From the main menu go to “Image > Stacks > Reslice [/]. . ..” 11. In the popup window, go to “Start at” and choose “Left” from the pulldown menu and check the “Rotate 90 degrees” checkbox. Press OK. 12. Move the slider at the bottom of the new kymograph window to move the line scan along the X axis of the original cilia movie until clear peaks appear (Fig. 12.4C). 13. From the main menu choose “Edit > Copy” and then “File > New > Internal Clipboard.” Save the final kymograph in the appropriate format by choosing “File> Save as.” 14. Count the number of peaks in the kymograph to measure ciliary beat frequency in Hz (beats per second) (Fig. 12.4C).

6. SUMMARY Planaria are an emerging and attractive model organism to study cilia. This is due to an easily accessible multiciliated epithelium, with motile cilia that contain structurally and genetically conserved 9 þ 2 axonemes.

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These cilia beat collectively and in synchrony to propel a layer of mucus and provide the organism with its characteristic gliding locomotion. In addition, planarians are genetically amenable with established robust tools for gene loss-of-function experiments by RNAi and a fully sequenced genome. Finally, planarians are cheap and easy to maintain and propagate in large numbers in a laboratory setting to serve multiple experimental needs.

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