Analysis of primary cilia in the developing mouse brain

CHAPTER

Analysis of primary cilia in the developing mouse brain

6

Judith T. M. L. Paridaen1, Wieland B. Huttner, Michaela Wilsch-Bra¨uninger1 Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany 1

Corresponding authors: E-mail: [email protected]; [email protected]

CHAPTER OUTLINE Introduction .............................................................................................................. 94 1. Neurogenesis in the Developing Neocortex ............................................................ 94 2. Primary Cilia in the Developing Neocortex ............................................................. 96 3. Material .............................................................................................................. 97 3.1 Visualization of Neural Progenitor Primary Cilia by Immunofluorescence .... 97 3.2 Visualization of Neural Progenitor Primary Cilia by Live Imaging................ 98 3.2.1 In utero electroporation....................................................................... 98 3.2.2 Live imaging of primary cilia in cultured dissociated dorsal telencephalon........................................................................... 99 3.2.3 Live imaging of organotypic slices of dorsolateral telencephalon ........... 99 3.3 Electron Microscopy ............................................................................ 100 3.3.1 General preparations ........................................................................ 100 3.3.2 Conventional TEM and SBF-SEM analysis ......................................... 101 3.3.3 Sample preparation for CLEM ........................................................... 102 4. Methods ............................................................................................................ 103 4.1 Visualization of Neural Progenitor Primary Cilia by Immunofluorescence .. 103 4.2 Visualization of Neural Progenitor Primary Cilia by Live Imaging.............. 104 4.2.1 In utero electroporation to introduce plasmids encoding primary ciliary and membrane markers ................................................................... 105 4.2.2 Live imaging of primary cilia in cultured dissociated dorsal telencephalon......................................................................... 107 4.2.3 Live imaging of organotypic slices of dorsolateral telencephalon ......... 109 4.2.4 Analysis of results............................................................................. 110 4.3 Visualization of Neural Progenitor Primary Cilia by Conventional EM ........ 111 4.3.1 General sample preparation for EM ................................................... 111 4.3.2 Conventional plastic embedding and serial sectioning........................ 115

Methods in Cell Biology, Volume 127, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.12.012 © 2015 Elsevier Inc. All rights reserved.

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4.4 Visualization of Neural Progenitor Primary Cilia by SBF-SEM .................. 117 4.4.1 SBF-SEM contrasting........................................................................ 118 4.4.2 SBF-SEM embedding ....................................................................... 118 4.4.3 Checking and mounting of SBF-SEM samples ................................... 119 4.4.4 SBF-SEM investigation...................................................................... 119 4.4.5 SBF-SEM image analysis .................................................................. 120 4.5 Visualization of Neural Progenitor Primary Cilia by Tokuyasu Cryosectioning and CLEM .......................................................................................... 120 4.5.1 Preparation of Tokuyasu samples...................................................... 121 4.5.2 Cryosectioning.................................................................................. 123 4.5.3 Immunolabeling................................................................................ 124 4.5.4 CLEM investigation ........................................................................... 125 Conclusion ............................................................................................................. 126 Acknowledgments ................................................................................................... 126 References ............................................................................................................. 126

Abstract Stem and progenitor cells in the developing mammalian brain are highly polarized cells that carry a primary cilium protruding into the brain ventricles. Here, cilia detect signals present in the cerebrospinal fluid that fills the ventricles. Recently, striking observations have been made regarding the dynamics of primary cilia in mitosis and cilium reformation after cell division. In neural progenitors, primary cilia are not completely disassembled during cell division, and some ciliary membrane remnant can be inherited by one daughter cell that tends to maintain a progenitor fate. Furthermore, newborn differentiating cells grow a primary cilium on their basolateral plasma membrane, in spite of them possessing apical membrane and adherens junctions, and thus change the environment to which the primary cilium is exposed. These phenomena are proposed to be involved in cell fate determination and delamination of daughter cells in conjunction with the production of neurons. Here, we describe several methods that can be used to study the structure, localization, and dynamics of primary cilia in the developing mouse brain; these include time-lapse imaging of live mouse embryonic brain tissues, and analysis of primary cilia structure and localization using correlative light- and electron- and serial-block-face scanning electron microscopy.

INTRODUCTION 1. NEUROGENESIS IN THE DEVELOPING NEOCORTEX Embryonic neural stem and progenitor cells generate a diverse array of neuronal and glial cell types during development. In the developing mammalian dorsolateral telencephalon that generates the neocortex, these neural stem and progenitor cells show diverse morphologies and behavior, which are linked to the expansion of the neocortex during evolution (Florio & Huttner, 2014). Early in embryonic development, the developing brain is organized as a pseudo-stratified epithelium

1. Neurogenesis in the Developing Neocortex

consisting of highly polarized neural stem cells with apical and basal processes that span from the ventricle to the pia (basal lamina) (Farkas & Huttner, 2008) (Figure 1). Two main types of neural progenitors exist, namely the apical progenitors (APs) and basal progenitors (BPs). APs include neuroepithelial cells and apical radial glial cells that localize to the ventricular germinal zone (VZ) and divide mainly at the ventricular surface. In contrast, BPs comprise basal radial glia cells and intermediate progenitors that reside and divide in a more basally positioned germinal zone named the subventricular zone (SVZ; Figure 1). In the developing mouse dorsolateral telencephalon that develops into the neocortex, neurons are produced from embryonic day (E)10 onwards. Prior to the onset of neurogenesis, APs undergo symmetric

ventricle

ciliary membrane (CM) mother centriole (MC) daughter centriole signals (CSF)

BP

VZ AP

AP

AP

BP

signals (VZ/SVZ) adherens junctions signal transduction AP apical progenitor

SVZ

interphase

BP basal progenitor

daughter daughter cell 1 cell 2 CM retention CM+ CM by MC mitosis

daughter daughter cell 1 cell 2 apical basolateral cilium cilium

daughter daughter cell 1 cell 2

FIGURE 1 Primary cilia in progenitors in the developing dorsal telencephalon. Apical progenitors (AP; first cell on the left) display an apical primary cilium (magenta (dark gray in print versions)), originating from the mother centriole (yellow (light gray in print versions)) that protrudes into the brain ventricle. In the ventricle, the primary cilium detects or releases signals (light blue (very dark gray in print versions) dots) in(to) the cerebrospinal fluid that are transmitted to the cell/nucleus (light blue jagged arrows). In dividing APs (second cell from the left), the mother centriole retains a ciliary membrane remnant (CM, magenta (light gray in print versions)) throughout mitosis. The mother centriole (yellow (white in print versions)) and CM are inherited by a daughter cell that quickly reestablishes an apical primary cilium and tends to remain an apical progenitor (third cell from the left). This cell usually also inherits the basal process. The other daughter cell inherits the daughter centriole (blue (light gray in print versions)) that docks to the basolateral membrane, where it initiates basolateral ciliogenesis (middle right). The basolateral cilium can receive or send signals (gray dots) from/to the basal extracellular space and transmit these to the cell body and nucleus (gray arrows). Ultimately, this cell shows apical constriction (double-headed arrow) and retracts the apical process from the adherens junction belt (dotted arrow). The cell delaminates from the ventricular surface (cell to the right) and migrates basally as a BP or a neuron (multipolar cell shown as light gray background).

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CHAPTER 6 Analysis of primary cilia in the developing mouse brain

proliferative divisions leading to an increase in their numbers. During neurogenesis, APs predominantly undergo asymmetric divisions, generating one AP daughter and one neurogenic daughter cell (neuron or BP). BPs delaminate from the VZ and migrate to the SVZ. In the mouse developing neocortex, BPs divide mainly symmetrically to yield two neurons, thus doubling the neuron output. The neurons, generated either directly from APs or indirectly from BPs, migrate to the cortical plate at the basal side of the cortical wall. The cortical plate will further develop into the six-layered cortex typical for the mammalian brain (Florio & Huttner, 2014).

2. PRIMARY CILIA IN THE DEVELOPING NEOCORTEX APs display apico-basal polarity and have slender apical and basal processes that contact both the ventricular fluid and the basal lamina, respectively, which are hundreds of microns apart. Importantly, APs adhere to neighboring cells through the adherens junction belt. In interphase APs, the centrosome is docked at the apical plasma membrane (Farkas & Huttner, 2008) (Figure 1, left). The mother centriole within the centrosome functions as the basal body to anchor a primary cilium. This 0.5e2-mm-long cilium extends into the cerebrospinal fluid (CSF)filled lateral ventricle (Louvi & Grove, 2011; Sotelo & Trujillo-Cenoz, 1958). Here, the primary cilium is able to detect signals present in the CSF, such as Shh, IGF, and EGF (Higginbotham et al., 2012; Lehtinen et al., 2011; Yeh et al., 2013). In dividing APs, a remnant of the primary cilium membrane (CM) can be internalized together with the mother centriole and is asymmetrically inherited together with the mother centriole by one of the daughter cells (Figure 1) (Paridaen, Wilsch-Bra¨uninger, & Huttner, 2013). Interestingly, in asymmetric AP divisions, the mother centriole and CM are preferentially inherited by daughter cells that remain APs (Paridaen et al., 2013; Wang et al., 2009). These daughter cells are able to reestablish a primary cilium earlier than their sister cells. In contrast, the daughter cells without inherited CM need to undergo de novo ciliogenesis that involves docking of the new mother centriole to the membrane and recruitment of Golgi-derived ciliary membrane prior to axoneme extension (Figure 1) (Follit, Tuft, Fogarty, & Pazour, 2006; Pedersen, Veland, Schroder, & Christensen, 2008; Sorokin, 1962). The asynchronous ciliogenesis is likely to differentially expose pairs of daughter cells to proliferative signals from the CSF, thus introducing asymmetric signaling between pairs of daughter cells. Therefore, the daughter cells receiving the mother centriole and CM may be destined to remain proliferative, whereas the daughter cells without CM need to undergo de novo ciliogenesis and may be destined to differentiate. Interestingly, upon the onset of neurogenesis, cells appear that extend a primary cilium docked to the basolateral rather than apical plasma membrane (Figure 1, middle) (Wilsch-Bra¨uninger, Peters, Paridaen, & Huttner, 2012). These

3. Material

basolateral (BL) cilium-bearing cells contact the ventricle, are integrated into the adherens junction belt, and display full apico-basal polarity (Figure 1, middle). The BL cilia are likely to receive (or send) signals from (to) the basal extracellular space rather than from the CSF, with possible consequences for their downstream fate. Because most BL cilia-bearing cells express a BP-specific transcription factor, they constitute newborn BPs. Upon inducing increased numbers of BPs by overexpression of a specific transcription factor, an increase in the number of BL cilia was observed (Wilsch-Bra¨uninger et al., 2012). Constriction of the adherens junction belt of some BL cilia-bearing cells implies that they are about to delaminate from the adherens junction belt and migrate to the SVZ (Figure 1, right). The presence of duplicated centrioles (characterizing late G1/S/G2 stages of the cell cycle (Nigg & Stearns, 2011)) in BL cilia-bearing cells indicates that the basolateral plasma membrane is the final destination of the cilia and not merely an intermediate position in reestablishing the primary cilium on the apical plasma membrane after mitosis. Clearly, ciliogenesis plays an important role in embryonic neurogenesis. Here, we will describe methods that can be used to study ciliogenesis in the developing brain, focusing on the analysis of apical and BL cilia, and the CM remnant in dividing and interphase APs, respectively, within the developing mouse neocortex. Techniques described include live imaging of APs in cultures of dissociated embryonic dorsolateral telencephalon and organotypic slice cultures, and electron microscopy (EM) analyses such as conventional transmission EM (TEM), serial block face-scanning EM (SBF-SEM), and correlative light and electron microscopy (CLEM). A combination of these techniques will lead to a comprehensive overview of the dynamics and composition of primary cilia in the developing brain.

3. MATERIAL 3.1 VISUALIZATION OF NEURAL PROGENITOR PRIMARY CILIA BY IMMUNOFLUORESCENCE • • • • • • • • • • • •

4% paraformaldehydedharmful! phosphate-buffered saline (PBS) sucrose Tissue-Tek (OCT, Sakura) or equivalent freezing medium dry ice embedding peeling molds cryostat (Microm, HM560, or equivalent) vibratome (Leica, VT1200S, or equivalent) low-melting-point agarose glycine Triton X-100 2% filtered gelatine in PBS, stored at
20  C

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CHAPTER 6 Analysis of primary cilia in the developing mouse brain

• • • • • • •

NaCl rabbit anti-Arl13b antibody (Proteintech, 17711-1-AP; 1:500 dilution) mouse monoclonal anti-gamma tubulin antibody (Sigma, T6557, 1:300 dilution) secondary antibodies (e.g., donkey anti-rabbit Alexa-488 and donkey anti-mouse Alexa-555, Invitrogen, 1:500 dilution) 40 ,6-diamidino-2-phenylindole (DAPI; Roche) Mowiol 4-88 (Sigma, 81,381) 0.17-mm-thick coverslips.

3.2 VISUALIZATION OF NEURAL PROGENITOR PRIMARY CILIA BY LIVE IMAGING 3.2.1 In utero electroporation •

• •

• • • • • • • •

• • • • • • • • •

borosilicate glass capillaries with filament, 1.2 mm outer diameter  0.69 mm inner diameter (Sutter Instruments, BF 120-69-10; determine temperature with ramp test of your capillary puller machine, pull 100, velocity 100, time 100) microcapillary puller (Sutter Instruments) endotoxin-free maxi-kit (Qiagen) prepped DNA in sterile PBS (in 3e5 mg/mL concentration) ciliary marker: CMV-promoter-driven fluorescent protein (FP)-tagged Arl13b (e.g., EGFP-Arl13b, Arl13b-mKate2 (Paridaen et al., 2013)) membrane marker: CMV-promoter-driven N-terminal 20 AA palmitoylation motif of GAP43 fused to EGFP, mKate2, or tdTomato microloader tips (Eppendorf) Fast Green, 2.5% solution in sterile PBS two bottles of prewarmed sterile PBS (penicillin/streptomycin can be added) electroporator (Harvard Apparatus, BTX ECM830) þ footswitch platinum tweezer electrode (Nepagene, CUY650P3) picoliter microinjector þ micromanipulator (alternative: mouth-controlled pipette system) heat plate set to 37  C table-top anesthesia system consisting of isoflurane vaporizer þ nose cone þ clear Perspex box þ activated charcoal filter to capture excess isoflurane (Vetequip and Vaporguard) analgesics, syringe and needle (27 3/4 G) sterile gauze 70% ethanol instrument hot-bead sterilizer Dumont forceps, curved forceps (2), small vannas scissor, small spatula (Fine Science Tools) vicryl suture (5-0, P-3, 45 cm, Ethicon) autoclip surgical wound clips 9 mm and clip applier heat pad clean mouse cage.

3. Material

3.2.2 Live imaging of primary cilia in cultured dissociated dorsal telencephalon • • • •

• • • • • • • • • • • • • • •

high-precision 0.17-mm-thick glass-bottomed Petri dish (Mattek Corporation, P35G-0.17-14-C) for DIC imaging: glass top cover (Mattek Corporation, P35GTOP-0-20-C) cell culture-grade sterile poly-D-lysine hydrobromide, Mr 30,000e70,000 (Sigma, P7280) Tyrode solution, prewarmed at 37  C: dissolve Tyrode’s salt (Sigma, T214510x1L) with 1 g of NaHCO3 and 13 mM HEPES in 1 L of distilled H2O. Adjust the pH of the solution to 7.4. Filter sterilize the solution and store at 4  C for up to 1 month. 35  10 mm Petri dishes silicone-coated Petri dish (custom made with Sylgard 184 Kit Silicone Elastomer) dissection Dumont forceps 55 (2) 20 G ophthalmic V-lance knife (Alcon, 8065912001) Pasteur pipettes, plastic pipettes and tips dissociation kit (e.g., neural tissue dissociation kit (papain-based), Miltenyi Biotech 130-092-628) prewarmed DMEM medium 4.5 g/L glucose, glutamax with phenol red penicillin/streptomycin cell culture-grade fetal calf serum basic fibroblast growth factor (bFGF, Fgf2) epidermal growth factor (EGF) Trypan blue hemocytometer wide-field microscope system with environmental control, CO2 supply, high-precision motorized stage, and high-quality CCD camera.

3.2.3 Live imaging of organotypic slices of dorsolateral telencephalon • • • • • • • • • • •

Cellmatrix type 1-A (Kyowa chemical products, 631-00651) 5 DMEM/F12 (Sigma, D8900), filter-sterilized reconstitution buffer (262 mM NaHCO3, 0.05 N NaOH, 200 mM HEPES), filter-sterilized, aliquotted, and stored at 4  C prewarmed Tyrode’s solution 35  10 mm Petri dishes high-precision 0.17-mm glass-bottomed Petri dish (Mattek Corporation, P35G-0.17-14-C) silicone-coated Petri dish (custom made with Sylgard 184 Kit Silicone Elastomer) dissection forceps Dumont 55 (2) 20 G ophthalmic V-lance knife (Alcon, 8065912001) surgical 4.0 cutting edge needle blade micro-knife (Sharpoint, 78-6810) Pasteur pipettes, plastic

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CHAPTER 6 Analysis of primary cilia in the developing mouse brain

• •

• • • • • • • • •

pipettes and tips slice culture medium: Add 10 mL of immediately centrifuged rat serum (Charles River Laboratories, Japan), 1 mL of 2 mM L-glutamine, 1 mL of penicillinestreptomycin (100), 1 mL of 100  N-2 supplement (Invitrogen, 17504-044), 2 mL of 50  B-27 supplement (Invitrogen, 17502-048), and 1 mL of HEPES buffer to 84 mL of Neurobasal medium (Invitrogen, 21103-049). Store 5-mL aliquots at
20  C until use. basic fibroblast growth factor (bFGF/Fgf2) epidermal growth factor (EGF) gas-permeable membranes, circle sut, Fep-foil 0.17 (Helmut Saur Laborbedarf, 0727.017) silicone grease (f.i. Bayer, Baysilone silicone grease) whole embryo incubator (Ikemoto scientific technology, RKI-10-0310) O2 (40%), CO2 (5%), N2 (55%) mixture, 50 L (Air Liquide) multiphoton microscope with long-distance high numerical aperture (NA) 40 objective Bachofer chamber with tubing and humidifier for humified O2 supply objective heater.

3.3 ELECTRON MICROSCOPY 3.3.1 General preparations 3.3.1.1 Buffers and fixatives • • • •

•

PBS containing 0.9 mM CaCl2/0.5 mM MgCl2 0.1 M PIPES pH 7.4 (Merck, 1.10220) 0.1 M phosphate buffer pH 7.4 16% paraformaldehyde, EM grade (EMS, Cat. 15710), 4% final concentration for immunolabeling, 1e2% final concentration in combination with glutaraldehyde for morphologydharmful! 25% glutaraldehyde, EM grade (EMS, Cat. 16220), 1% final concentration for morphology, maybe addition of 0.05% to fixative for immunolabelingdharmful!

3.3.1.2 Vibratome sectioning • • • • • • • •

Dumont fine dissecting forceps #5 (or #55) agarose, low melting point (EMS, Cat. 10207), 4% in PBS þ CaCl2/MgCl2 razor blades (EMS, 72000)dbreak the blades in half, while still wrapped in the covering paperdsharp! Roticoll1 glue (Roth, 0258.1) disposable embedding molds, 8 mm bottom (Polysciences, 18985) painting brush, size 1 small Petri dish (Nunc, 153066) vibratome (Leica, VT1200)

3. Material

3.3.2 Conventional TEM and SBF-SEM analysis Most of the chemicals used for fixation, contrasting, and embedding are harmful, toxic, or cancerogenic (indicated in italic). These steps should be performed in a well-ventilated fume hood.

3.3.2.1 Heavy metal contrasting • • • • •

• • •

osmium tetroxide, 4% aqueous solution, EM grade (EMS, 19190)dtoxic! potassium hexacyanoferrate (II) trihydrate (Sigma, P9387), 3% stock solution (kept at 4  C in the dark)dirritating! tannic acid (EMS, 21710), 0.2% in water (freshly prepared) thiocarbohydrazide (TCH) (Aldrich, 223220), 1% in water, dissolve at 60  C before usedtoxic! uranyl acetate (Polysciences, 21447), 3% stock solution in water (kept at 4  C in the dark), used at 0.5% in 25% methanol for en bloc staining, at 0.3% in 1.8% methylcellulose for contrasting of immunolabeled sectionsdtoxic and radioactive! L-aspartic acid (Aldrich, A9310-0), 0.03 M stock, pH 5.5 adjusted with 1 N KOH (kept at 4  C) lead nitrate (EMS, 10099-74-8)dtoxic! prepare lead aspartate by freshly dissolving lead nitrate (0.02 M final) in 0.03 M aspartate solution, adjusting to pH 5.5 with KOH, and preincubating the solution for 30 min at 60  C prior to use (Walton, 1979).

3.3.2.2 Dehydration and embedding • • • • • • • •

•

absolute ethanol, dilution series at 30%, 50%, 70%, 80%, 90%, 96%, and 100% molecular sieve 0.3 mm (Merck, 1.05704) microscope slides Teflon solution (MS-143v PTFE Release agentddry lubricant, Millere Stephenson) Aclar foil, 7.8 mil (199 mm thick) (EMS, 50425) silicone embedding mold (EMS, 70901) plastic Pasteur pipettes, fine and wide tip Epon replacement (called “EPON” in the text): glycidether 100 (Roth, 8619.2); hardener MNA (Roth, 8639.2); glycidether hardener DBA (Roth, 8623.2); DMP-30 (EMS)dall resin monomers are irritating, cancerogenic, or toxic! “EPON” resin, hard: • 37 mL glycidether (e.g., Roth or Embed 812, EMS) • 25 mL DBA (DDSA) • 20 mL MNA • 1.3 mL DMP-30 Durcupan, ACM, four components (Fluka)dsee above! Durcupan resin, hard: mix in this order, stir thoroughly after each component

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• • • •

11.4 g component A 10 g component B 0.05e0.1 g component D 0.3 g component C

resins are aliquotted in 10-mL plastic syringes, sealed with parafilm, and stored at
20  C. They are warmed up to room temperature prior to use.

3.3.2.3 Mounting and cutting • • • • • • • • • • • • • • • • • •

cyanoacrylate glue (superglue, e.g., UHU) CW2400 conductive epoxy (Circuitworks) Gatan 3View sample pins (Gatan or workshop) glass knives (Leica) diamond knife (Diatome) eyelash (mounted with nailpolish to toothpick) Toluidine Blue O (EMS, F22050), 0.5% in 1% disodium tetraborate mini hotplate (Agar Scientific) 60  C oven (Memmert, UFB500 with ventilation) dissection microscope (Leica, MZ95) light microscope (Leica, DM E) EM slot grids 2  1 mm (Science Services, G2010-Cu) Morgagni EM (FEI) at 80 kV Magellan 400 SEM (FEI) 3View2XP SBF-SEM microtome (Gatan) Digital Micrograph software, Version 2.30 (Gatan) Fiji software (Schindelin et al., 2012) with TrakEM2 plugin (Cardona et al., 2012) 3View Import List plugin for Fiji (downloadable from www.fiji.sc)

3.3.3 Sample preparation for CLEM 3.3.3.1 Tokuyasu cryosectioning • • • • • • • • • • •

37  C incubator large Petri dish for wet chamber 12% gelatine (baker’s type) in PBS (test batch for precipitation with phosphate), aliquots stored at
20  C 2.3 M sucrose in 0.1 M phosphate buffer, aliquots stored at
20  C methylcellulose 25 cpi (Sigma, M-6385), 2% in water, stored at 4  C polyvinylpyrrolidone (PVP) (MW 10 000) (Aldrich, 856452): use 15% PVP, 0.03 M Na2CO3, 1.7 M sucrose, aliquots stored at
20  C parafilm filter paper (Whatman, 1001) scalpel blades #11 (EMS, 72049-11)dsharp! snap-on lids (Roth, LC86.1) cryo-pins (Leica or W þ O Niettechnik: rivet Din 661, 2  10 mm, #8040397 (aluminum) or #8540314 (copper))

4. Methods

• • • • • • • • • • • • •

pick-up loop (Perfect Loop, Leica, or homemade from copper wire) ice bucket (or styrofoam box) cryo-forceps (Leica) Formvar (15/95 polyvinyl formal resin) (EMS, 15800), 1% in chloroform 100 mesh copper grids (Science Services, G100H-Cu), rinsed in acetone self-adhesive paper (address) sticker, 63  33 mm (Herma) microscope slides coverglass 24  60 mm coverglass round, 12 mm carbon coating device (Baltec/Leica, MED 020) glow discharger (Baltec/Leica, MED 020) ultramicrotome with cryochamber (Leica, UCT6 with FCS) liquid nitrogen (and liquid nitrogen sample storage tank)dmay cause burns and suffocation!

3.3.3.2 Immunolabeling • • • • •

• • • • • • • • • • • •

BSA: serum albumin fraction V (Sigma), 0.5% final gelatine (baker’s type), 0.2% final PBS blocking buffer: PBS þ 0.2% gelatine þ 0.5% BSA, freshly prepared primary antibodies (10 higher concentrated than for fluorescent labeling of cryo- or vibratome LM sections): rabbit anti-Arl13b (ProteinTech, 1711-1-AP) used at 1:50 in blocking buffer Protein Ae10 nm (Department of Cell Biology, Utrecht University) fluorescently labeled secondary antibodies (donkey anti-rabbit-Alexa 564) DAPI 0.1 mg/mL in water (0.01 mg/mL final) cold metal plate (ca. 10  10 cm) ice bucket contrasting loops (homemade: copper wire wrapped around a 3.3-mm drill with a densely coiled stalk, mounted with Epon into the tip of blue (1 mL) pipette tips) filter paper (Whatman) Axioplan 2 epifluorescence light microscope (Zeiss) RT monochrome Spot camera (Diagnostic Systems) HBO100 halogen lamp 63 Acroplan objective, NA 0.8, oil-free (Zeiss) 50% glycerol in PBS.

4. METHODS 4.1 VISUALIZATION OF NEURAL PROGENITOR PRIMARY CILIA BY IMMUNOFLUORESCENCE The primary cilia of neural progenitors and neurons in the developing mouse brain can be visualized by Arl13b immunofluorescence (Caspary, Larkins, & Anderson,

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2007). We recommend combining Arl13b with gamma-tubulin immunolabeling to also visualize the centrosomes. Another ciliary protein that is present in neural progenitor cells is type III adenylyl cyclase (ACIII; (Higginbotham et al., 2012)). Due to dense staining of filaments in neural progenitors, acetylated tubulin is not suitable for use as a ciliary marker. Sacrifice the pregnant dam and collect the uterus in PBS. Quickly dissect the whole heads (embryonic day (E) 9.5eE12.5) or brains (E13.5 and older) from the embryos using dissection forceps. Transfer the heads or brains quickly into 4% paraformaldehyde in phosphate buffer and fix for 4 h or overnight at 4  C. Wash the samples three times with PBS. For preparing cryosections, incubate samples in 30% sucrose in PBS at 4  C until they sink to the bottom of the tube. Next, embed the samples in embedding molds in Tissue-Tek. Take care to orient the samples properly. Freeze on dry ice and store at
20/
80  C. Cut coronal 14e20-mm sections using a cryostat. After drying, sections can be stored at
20  C until use. Alternatively, 50e100-mm brain sections can be cut using a vibratome. To this end, embed the brains in 4% low-melting-point agarose in PBS prior to cutting. Collect the slices in a 24-well plate filled with PBS and process for immunofluorescence within 1e2 weeks. For immunolabeling, permeabilize the sections by incubation in 0.3% Triton X-100 in PBS for 30 min, followed by quenching in 0.1 M glycine pH 7.4 for 30 min. Wash sections three times with PBS with 0.3% Triton X-100, 300 mM NaCl, and 0.2% gelatine. Incubate sections overnight at 4  C in a humified box with primary antibody diluted in PBS containing 0.3% Triton X-100, 300 mM NaCl, and 0.2% gelatine. For cryosections, apply 100 mL of antibody solution and cover the slide with parafilm. For vibratome sections, use 100e200 mL of antibody solution depending on the number of sections. Apply PBS to all empty wells and wrap the plate in plastic foil. Following washes with PBS containing 0.3% Triton X-100, 300 mM NaCl, and 0.2% gelatine, incubate the sections with the appropriate secondary antibodies (anti-rabbit and anti-mouse) containing 0.5 mg/mL DAPI for 1 h at room temperature for cryosections, and overnight at 4  C for vibratome sections. After washing at least three times with PBS, mount the sections in Mowiol using 0.17-mm-thick coverslips. The sections can be imaged using standard confocal microscope setups. We use 40 or 60 objectives and image at the maximum optical resolution (200 nm) to visualize the ciliary membrane remnants in dividing cells.

4.2 VISUALIZATION OF NEURAL PROGENITOR PRIMARY CILIA BY LIVE IMAGING To follow the dynamics of primary cilia in neural progenitors, one can transiently express a ciliary marker in combination with a marker outlining the cell by electroporation into the embryonic brain. For marking the cilia, FP-tagged versions of Arl13b, somatostatin receptor 3 (Sstr3) (Ha¨ndel et al., 1999), serotonin receptor 5HT6 (Berbari, Johnson, Lewis, Askwith, & Mykytyn, 2008), or the Shh component

4. Methods

1. In utero electroporation DNA

2. Isolate telencephalon n=2-4 (for orientation)

3. Remove meninges and separate hemispheres

+ – dl v dm v dl GE

GE

dissection forceps

E12.5-E16.5 mouse embryo

4. Dissect hemispheres (dissociated cell culture) Max. 1 hour

(A)

5. Dissociation 6. Plate cells n=4-8 15 min + 5 min 37 °C enzyme solutions

+ bFGF (+ EGF)

Max. 1 hour

widefield microscope

5. Slicing

6. Embed slices 7. Culture

5-7 slices + collagen 40 min 37 °C

micro surgical knife

8. Live image

>3 hours 37 ºC

ophthalmic knife

4. Dissect hemispheres (organotypic slice culture)

(B)

7. Culture

~2 x 10

8. Live image

+ SCM + bFGF (+ EGF) 1-2 hours 37 °C (40% O ) multiphoton microscope

FIGURE 2 Schematic drawing of procedure used for live imaging of cultured dissociated dorsolateral telencephalon (A) and organotypic slices (B) using wide-field and multiphoton microscopy, respectively. Steps 1e3 describe the electroporation, and initial brain dissections that are shared by both methods. Crucial information on timing, tools, number of brains/ hemispheres/cells/slices as well as medium supplements are indicated in italics. See main text for further details. bFGF, basic fibroblast growth factor; dl, dorsolateral telencephalon; dm, dorsomedial telencephalon; EGF, epidermal growth factor; GE, ganglionic eminence; SCM, slice culture medium; v, lateral ventricle.

Smoothened (Rohatgi, Milenkovic, & Scott, 2007) can be used. An important issue is that overexpression of ciliary proteins may induce increased ciliary length, which may affect ciliary function, as has been observed for Sstr3 and 5HT6 (Guadiana et al., 2013; our unpublished observations). In our experience, expression of CMV-promoter-driven Arl13b has only minimal effects on cilium length. To follow ciliary dynamics in relation to neural progenitor morphology, we combine expression of FP-tagged Arl13b with a membrane-targeted FP. After electroporation, live brain tissue is processed into either a single cell suspension or organotypic slices (Figure 2).

4.2.1 In utero electroporation to introduce plasmids encoding primary ciliary and membrane markers In utero electroporation of mouse embryos is routinely used to introduce transient expression of DNA constructs (Saito & Nakatsuji, 2001; Tabata & Nakajima, 2008). It consists of microinjection of DNA plasmid(s) into the brain ventricle, followed by

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application of an electric field that transiently creates pores in the plasma membrane, allowing entry of DNA into the cytoplasm of cells with a membrane bordering the ventricle (Figure 2, top panel). Therefore, immediately after electroporation, the vast majority of electroporated cells constitute APs. When performed properly, embryo survival is close to 100%, and depending on the constructs used, the plasmids are efficiently expressed in 50e80% of the electroporated embryos. Electroporation efficiency decreases with increasing size of the constructs, with plasmids >10 kilobases showing very low transfection efficiency. Visualization and injection into the lateral ventricles is easiest at the E12/E13 stages. Therefore, we recommend practicing the procedure in embryos at these stages before attempting other stages.

4.2.1.1 Procedure Prepare pulled microcapillaries. Dilute endotoxin-free prepped DNA constructs to 0.5e2 mg/mL in sterile PBS with 0.25% Fast Green to allow easy visualization of brain ventricles. We use pCMV-EGFP-Arl13b of pCMV-Arl13b-mKate2 (Paridaen et al., 2013) at final concentrations of 0.75 and 1.5 mg/mL, respectively. Prewarm PBS and the heat plate to 37  C. Sterilize instruments by ethanol disinfection/sterilizer. Prepare a syringe with analgesics according to the manufacturer’s instructions, taking care to remove air bubbles. Connect a platinum electrode to the electroporator and place the electrode ends in a bottle with PBS. Program the electroporator according to the following parameters: voltage 30e40 V (E12: 30V; E13: 35V; E14: 40V), pulse length 50 ms, 5e6 pulses, pulse interval 1 s, unipolar pulse. Place the pregnant dam in the Perspex box and open isoflurane flow to 4% with O2 flow at 0.5 liters per minute to induce anesthesia. Carefully monitor the heart rate, which gets slower with proper anesthesia. When the mouse is fully anesthetized, move the mouse to the heat plate covered with sterile gauze with its nose inside the nose cone and lower the isoflurane flow to 2e2.5%. Monitor the heart rate throughout the surgery. Gently fix the limbs to the heat plate with paper tape and shave the belly using an electric razor or razor blade. Disinfect the skin with iodide solution and ethanol. Administer the analgesics by subcutaneous injection. Lift the skin up with curved forceps and make a 1e2-cm incision in the belly midline with the vannas scissor. Loosen skin from the peritoneum and make a small incision in the peritoneum at the midline along the linea alba. Avoid damaging any blood vessels. Cover the belly with sterile gauze. Carefully pull out uteri horns by grabbing between embryos. Make sure to keep the surgery area and uterus wet with PBS throughout the surgery. Backfill a capillary with DNA solution and cut off the tip with dissecting forceps. Visualize the lateral ventricles and inject DNA solution until the ventricles are visibly green. Apply some fresh PBS to the uterus. Orient the electrode so that the anode is directly dorsal to the injected ventricle and apply the electric pulses. Avoid placing any electrode end directly on the placenta. To increase the number of electroporated cells, both hemispheres can be electroporated. Take care not to place the electrodes on the same area twice. To avoid abortion, the embryos closest to the uterus midline are usually not electroporated.

4. Methods

For live imaging of dissociated brain tissue, the dam can be sacrificed immediately and the uterus collected in prewarmed Tyrode solution. Proceed immediately with dissection (see section 4.2.2 below). For experiments using organotypic slices, return the uterus horns to the peritoneal cavity and fill the peritoneal cavity with PBS. Suture the peritoneum and close the skin using surgical clips. The total time of surgery and anesthesia should not exceed 1 h. Return the dam to a clean cage on a heat pad and monitor closely for recovery. To start imaging as soon as plasmids are expressed, sacrifice the animal after 6 h and dissect embryos immediately (see section 4.2.3 below). Usually, live imaging is started 8e12 h after electroporation.

4.2.2 Live imaging of primary cilia in cultured dissociated dorsal telencephalon 4.2.2.1 Dissociation of dorsolateral telencephalon For an overview of the procedure, see Figure 2(A). Prior to dissection, coat glassbottomed Mattek dishes with 100 mg/mL poly-D-lysine for at least 30 min. Wash three times with sterile PBS before use. For electroporation followed by dissociation into single cells, the pregnant dam is sacrificed immediately after finishing the surgery. The uterus is collected in prewarmed Tyrode salt solution. Electroporated embryos are dissected from the uterus and the heads collected into a fresh Petri dish with Tyrode salt solution. Brains are dissected one by one, the meninges are removed, and the telencephalon is split into the separate hemispheres using dissection forceps (Figure 2, top panel). For embryos
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medium containing 10% FCS and penicillin/streptomycin. Filter the resulting cell suspension through a 40-mm filter, and wash the tube and filter with an additional 0.5 mL of medium. Dilute 10 mL of the cell suspension with Trypan blue (1:10) in 100 mL medium and count the cells using a hemocytometer. The percentage of dead cells as determined by Trypan blue dye exclusion usually is 3e7%. Expected cell counts are 0.8e2.0  106/embryo (2 hemispheres) at E12.5, 4.0e6.0  106/embryo (2 hemispheres) at E14.5, and 8.0e10.0  106/embryo (2 hemispheres) at E15.5. Dilute cell suspension in medium to a final concentration of 1.3  106/mL for E14.5 and E15.5, and 0.9  106/mL for E12.5 cells. Add 10 ng/mL (final concentration) bFGF (R&D Systems) for E12 embryos, and 10 ng/mL bFGF plus 20 ng/mL EGF (Sigma) for E14e15 embryos. Plate cells by adding 2 mL of the diluted cell suspension to poly-D-lysine-coated glass-bottomed dishes. Culture the dishes at 37  C at 5% CO2 for at least 3 h to allow the cells to adhere before the start of live imaging.

4.2.2.2 Live wide-field imaging Expression of CMV-promoter-driven plasmids starts from 4 h postelectroporation. Therefore, live imaging can usually be started within 6e12 h after electroporation. Since the electroporation targets only cells with a ventricular contact, the vast majority of electroporated cells constitute apical radial glial cells. The average cell cycle of apical radial glial cells at E14.5 is w19 h (Arai, Pulvers, Haffner, Schilling, Nusslein, Calegari et al., 2011). For this reason, single electroporated cells are expected to divide within the first 24 h of imaging. For live imaging, a wide-field microscope setup with a high-quality cooled CCD camera, a high-precision motorized stage, a high NA water objective with a coverslip correction collar, and an environmental chamber with supply of humified 5% CO2 is advised. This will allow acquisition of images of single radial glial cells at intervals of 10e15 min at multiple positions. Acquisition of differential interference contrast (DIC) images aids with identification of mitotic stages, although it is not essential for the analysis. Our setup consists of a Deltavision Core wide-field microscope (Applied Precision) equipped with an Olympus UPlanApo 60 NA 1.20 water objective, a Photometrics Cool Snap HQ2 camera, a high-precision motorized stage, and an environmental chamber set to 37  C with custom-built CO2 control. Prior to the start of live imaging, let the objective, environmental chamber, and sample equilibrate at 37  C for at least an hour. In our experience, addition of phenol red to the medium does not lead to noticeable background fluorescence, but this may depend on the wide-field system used. DIC imaging may help with identifying dividing cells and enables observing the general morphology of the cells. For DIC imaging, the Petri dish should either be uncovered or covered with a glass top. For setting up DIC, close the field diaphragm and adjust to center the diaphragm edges in the field of view through the eyepiece. Open the diaphragm so that the edges are just outside the field of view. Place the polarizing filter and prism in the light path. Adjust the prism while using the eyepiece to check for the optimal contrast.

4. Methods

Select 30e40 stage positions of single labeled cells using the eyepiece. Use of a membrane-targeted FP makes it easier to assess cell health. Ensure selecting only healthy-looking cells without any obvious membrane blebbing. Next, optimize the exposure time and light power intensity using the acquisition software. To avoid overexposure of the image, select cells with similar fluorescence signals. The aim is to use the lowest light intensity and exposure times as possible to avoid phototoxicity. Use a gray value histogram in the acquisition software to ensure that all the observed signal intensities are within the scale of gray values. Usually, we adjust exposure parameters so that the maximum observed signal intensity is about 2/3 of the maximum gray value. For imaging primary cilia, we usually acquire z-stacks of 12e15 mm at 0.3-mm z-steps, with a pixel size of 0.23 mm, at intervals of 10e 15 min for a total time of 8e16 h. Importantly, we use 2  2 pixel binning to increase signal-to-noise ratios. For longer live imaging (>16 h), we recommend lowering the amount of acquired time points by increasing the interval times as phototoxicity can emerge from 12 to 16 h onwards.

4.2.3 Live imaging of organotypic slices of dorsolateral telencephalon 4.2.3.1 Preparation of organotypic brain slices For an overview of the procedure, see Figure 2(B). For live imaging of organotypic slices, sacrifice the pregnant dams about 6e24 h after EP, and collect the uterus in a 50-mL tube with prewarmed Tyrode solution. Prior to dissection, prepare the collagen mixture on ice. Add 250 mL of sterile water and 500 mL of 5 DMEM/ F12 to a 50-mL tube and keep on ice. Add 1.5 mL of Collagen 1-A cell matrix and mix. Add 250 mL reconstitution buffer and mix. Keep the mixture on ice until use. Dissect the embryo heads and transfer the heads to a fresh Petri dish with Tyrode solution. Dissect the brains and transfer to a fresh dish with Tyrode solution. Remove the meninges and separate the hemispheres. Transfer the hemispheres to a siliconcoated Petri dish with Tyrode solution. Using an ophthalmic V-lance knife, remove most of the dorsomedial telencephalon by cutting along the dorsomedial curve, and remove most of the ventral telencephalon (ganglionic eminence) by cutting slightly below the pallialesubpallial boundary, leaving a small strip of ganglionic eminence and dorsomedial telencephalon (Figure 2(B)). Turn the dorsolateral telencephalon so that the ventricular surface is up. Using a microsurgical needle blade, remove the anterior and posterior part of the dorsolateral cortex. Cut the remaining middle section into five to eight sections (150e400 mm thick). The best way to cut the slices is by pushing the needle blade down through the tissue onto the silicon. As an alternative, vibratome sectioning at 250 mm of brains embedded in low-melting agarose in PBS could be performed instead of manual slicing, although manual slicing is faster and results in better tissue health. Immediately after slicing, a fluorescence stereomicroscope can be used to select slices with electroporated cells. Alternatively, this can be done after the collagen embedding prior to imaging. Transfer 10e18 slices to the ice-cold collagen solution using a plastic Pasteur pipette with a cut-off tip. Keep the tube on ice. Transfer the slices with some extra collagen onto 35-mm glass-bottomed Mattek dishes kept on ice. Orient the slices in

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a lateral position so that one cutting edge faces the glass coverslip. Remove excess collagen by slow aspiration using a P200 pipette. Avoid introducing any bubbles. There should be a thin layer of collagen on top of the slices left. Incubate the dish for 5 min at 37  C before transferring to the incubator. After solidification of the collagen (40 min at 37  C), slowly add 2 mL of slice culture medium containing 10 ng/mL bFGF and 20 ng/mL EGF. Be especially careful when adding the medium directly on top of the slices as they may loosen from the collagen matrix. Culture the slices for at least 1 h before the start of live imaging in an incubator with supply of humified 40% O2. We use a custom-built Perspex box within a whole-embryo culture incubator (Ikemoto). Movies are started within 8e12 h after electroporation.

4.2.3.2 Live multiphoton imaging of organotypic slices To be able to image deeper into the slice, use a long-distance water objective with high NA for microscopy. We utilize a Zeiss LD C-Apochromat 40 1.1 NA longdistance water objective. Prior to live imaging, prewarm the Bachofer chamber and the objective using an objective heater to 37  C for at least 30 min. Select a dish containing electroporated cells and place it in the Bachofer chamber. The dish containing the slices is sealed with a gas-permeable membrane and silicone grease. Close the chamber and supply with 40% O2 and 5% CO2. Equilibrate the system for 30 min. Using the eyepiece, select slices that contain electroporated radial glial cells with their processes aligned more or less parallel to the z-plane. Avoid selecting cells that are close to either cutting edge as these cells usually are damaged. For the live imaging of organotypic slices, we use a Zeiss 710 Multiphoton Laser Scanning Microscope equipped with a tunable pulsed Ti-sapphire laser (Chameleon Vision II). Our system contains highly sensitive, non-descanned detectors to image the samples. Therefore, darken the room and switch off or blind any light source in the room. We additionally cover the entire microscope with thick black cloth. To image EGFP and tdTomato simultaneously, tune the laser to 940 nm. Set up the gain and offset in the acquisition software so that the laser power is as low as possible and no pixels are over- or underexposed. Use 2 averaging to improve signal-tonoise ratio. Set up z-stacks encompassing the apical endfoot of the cells of interest in the middle of the z-stack. We usually generate z-stacks of 50e100 mm. To image primary cilia, one needs to image at the maximum optical resolution. On our system, we employ a z-step of 0.58 mm and a pixel size of 0.18 mm. Depending on the microscope setup, two to three positions can be imaged at 12e15-min intervals for up to 15e20 h without apparent phototoxicity or photobleaching. To avoid phototoxicity in longer live imaging experiments, we recommend lowering the amount of acquired time points by increasing the interval times.

4.2.4 Analysis of results Example result images of Arl13b and g-tubulin immunofluorescence at the VZ for E10.5 and E14.5 are shown in Figure 3(A). Example still images of a time-lapse movie of a single dividing AP retaining a CM remnant throughout mitosis are shown

4. Methods

in Figure 3(B). For a typical example of a dividing electroporated AP and CM inheritance within an organotypic slice, see the still images in Figure 3(C). We use the freely available software package Fiji (based on ImageJ (NIH)) (Schindelin et al., 2012) to analyze the time-lapse movies. As time-lapse movie data files are large, one needs a computer system with high RAM and appropriate data storage capacities. If necessary, change the memory setup in the Fiji options menu. Maximum projection of the z-stack helps in identifying single apical radial glial cells and their primary cilium. However, for detailed analysis, one needs to manually analyze single z-planes within the stack. Analysis of single radial glial cells in organotypic slices is often hampered by the fact that many neighboring cells will be electroporated and expressing the plasmids as well. To circumvent this, one may use conditionally expressed constructs (Cre-lox based (e.g., Shitamukai, Konno, & Matsuzaki, 2011)) or low-titer retroviral constructs (Yu, Bultje, Wang, & Shi, 2009).

4.3 VISUALIZATION OF NEURAL PROGENITOR PRIMARY CILIA BY CONVENTIONAL EM Light microscopic immuno- and time-lapse imaging can give detailed information about ciliary distribution and dynamics as judged by the distribution of the fluorescent (ciliary) proteins used for the visualization. However, only labeled tissue structures will be visible, not the overall tissue context. The precise localization of cilia in respect to the plasma membrane or fine details of the ciliary structure (e.g., number of appendages or microtubule arrangement) is missed in fluorescent imaging. To fill this gap, an electron microscopic study of cilia in the developing mouse brain can be added to the analysis at the light microscope (LM). This will lead to a comprehensive view on the cilia in mouse brain development by utilizing the advantages of each method for different aspects.

4.3.1 General sample preparation for EM 4.3.1.1 Dissection and fixation for EM studies Good immobilization of cellular components at their native position within the cell is essential for studies on the ultrastructure and subcellular localization of proteins by transmission and scanning EM (TEM and SEM). Rapid dissection and fixation of the tissue reduces the timespan for proteases to act on the tissue and for membranes to disintegrate. To this end, mouse embryos taken from the mother (between E10.5 and 14.5) are best directly transferred into weak fixative (1/5 dilution, but work fast to avoid inhaling the aldehyde fumes) at room temperature. In the diluted fixative, the embryos are dissected out of the uterus and extraembryonic tissues with sharp forceps and are gently transferred (with a cut plastic Pasteur pipette or a spatula) into fixative consisting of 1% GA, 2% PFA in 0.1 M PIPES, 1 mM CaCl2, pH 7.4 for structural studies, or 4% PFA in 0.1 M PIPES, 1 mM CaCl2, pH 7.4 for immunolabeling. Embryos younger than E14.5 will be fixed as a whole; for older embryos only the heads will be fixed after careful removal of the skull. Fixation is performed at room temperature for 30 min to prevent depolymerization of microtubules, and

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CHAPTER 6 Analysis of primary cilia in the developing mouse brain

Immunofluorescence

(A)

FIGURE 3 Example results of primary cilia analysis by immunofluorescence (A) or time-lapse live imaging (B, C). (A) Immunofluorescence for primary cilia of E10.5 (left) and E14.5 (right). Insets in (A) show magnifications of single fluorescence channels for primary cilia (top, marked by Arl13b in magenta), centrosomes (middle, marked by g-tubulin in green), and the overlay (bottom) at the ventricular surface. Images are 10-mm z-stacks of 0.5-mm single z-planes. (B) Still images of wide-field live imaging of dissociated dorsolateral telencephalon showing DIC (top row only), membrane-EGFP (green), and Arl13b-mKate2 (magenta) signals in a dividing apical progenitor (white dashed lines) in cultured dissociated dorsolateral telencephalon from E12.5 mouse embryos. The arrow indicates the primary cilium membrane and its inheritance by one daughter cell (indicated by yellow dashed lines). The second daughter cell is indicated with red dashed lines. Images are 15-mm z-stacks of 0.3-mm single z-planes. Time intervals are indicated as hours:minutes at the top. The insets in the bottom row show magnifications of the ciliary membrane as marked by Arl13b-mKate2. (C) Still images of multiphoton live imaging of organotypic slices showing membrane-tdTomato (green) and EGFPArl13b (“Fire” lookup table as indicated) signals in a dividing apical progenitor (white dashed lines) in organotypic slices from dorsal telencephalon of E14.5 mouse embryos. The arrow indicates the primary cilium membrane and its inheritance by one daughter cell (indicated by yellow dashed lines). The second daughter cell is indicated with red dashed lines. Images are 25-mm z-stacks of 0.58-mm single z-planes. Time intervals are indicated as hours:minutes at the top. The boxed areas show magnifications of the ciliary membrane as marked by EGFP-Arl13b. Scalebar 10 mm, insets, 2 mm. (See color plate)

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(A)

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FIGURE 4 Cilia investigation by conventional EM and serial block faceeSEM. (A) Schematic drawing of the crucial steps in sample preparation for conventional TEM and SBF-SEM. Steps 1e4 (as indicated in italics on top) are shared by both imaging protocols. For the final step (5), samples for conventional EM have to be trimmed precisely for serial sectioning, whereas rough trimming (but with a clean edge) of the SBF-SEM sample is sufficient. However, for SBF-SEM samples, a good tissue contact to the metal pin for conductivity is required. (B) Examples of transmission electron micrographs of an apical (top, as indicated in white) and a basolateral (bottom) primary cilium in the mouse dorsolateral telencephalon at E10.5. The basal bodies of the cilia are shown at higher magnifications (right panels). The adherens junction belt (arrows) next to the ventricle (v), appendages (slim arrowhead: distal; wide

4. Methods

then continued at 4  C overnight. Tissues can be stored in diluted fixative for longer periods (if necessary up to a month without obvious effects on the general morphology), but must not be stored in pure buffer after (reversible) paraformaldehyde fixation (Hayat, 2000).

4.3.1.2 Vibratome sectioning As embryos are far too big for embedding as a whole, vibratome sectioning after fixation (Figure 4(A1)) provides several advantages: (1) specific brain regions can easily be selected, (2) sections from a single embryo can be treated under different conditions, and (3) the penetration of solutions or infiltration with embedding media is superior on the thin slices (200 mm). However, the slices are rather fragile, especially after strong fixation and heavy metal treatment, and have to be treated with appropriate care. For performing the vibratome sectioning, embryos are washed after fixation with 0.1 M PIPES, 1 mM CaCl2 and placed in prewarmed (37  C) 4% low-melting agarose in buffer in disposable molds. After cooling down (4  C), blocks containing the embryo in the correct orientation are trimmed with a razor blade and mounted with water-polymerizing glue on the vibratome sample support. In the vibratome, 200-mm sections are cut in a trough filled with PBS containing 0.9 mM CaCl2/ 0.5 mM MgCl2 (for keeping membrane integrity). The sections are transferred with a fine brush into a 3-cm Petri dish containing PBS þ CaCl2/MgCl2. Sections are further processed immediately or stored in 1% PFA in buffer overnight at 4  C.

4.3.2 Conventional plastic embedding and serial sectioning For observation of the morphology of cilia in the embryonic mouse brain by TEM, conventional plastic embedding and ultra-microtomy is performed (Figure 4(A3e 5), left). This will allow the observation of the cilia (e.g., their localization within the cell or their substructures such as appendages) at a high resolution in the tissue context without specific labeling. To this end, the vibratome sections of

=

arrowhead: subdistal), and the ciliary necklace (small lines) are indicated. Scalebars: left: 500 nm; right: 100 nm. (C) Examples of SBF-scanning electron micrographs of primary cilia in the mouse dorsal telencephalon at E14.5. The contrast has been inverted. Acquisition parameters: accelerating voltage: 1.9 kV; current: 100 nA; high vacuum; dwell time: 1.5 ms/ px; pixel resolution: 7.5 nm/px; section thickness: 50 nm; immersion lens backscattered electron detector. Top left panel, apical primary cilium of an apical progenitor protruding into the ventricular lumen (v). Right panel, series of 50-nm sections through a basolateral primary cilium of a basal progenitor cell with ventricular (v) contact (section number at top right of each panel). From the basal body (section 1e5), the axoneme is protruding into the extracellular space (tip on section 7e9, arrowhead). Bottom left panel, primary cilium of a delaminated basal progenitor in the subventricular zone. The cilium is sitting in a membrane pocket with a small opening to the extracellular space. Arrows indicate adherens junctions, arrowheads the ciliary tips. Scalebar, 500 nm.

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(glutaraldehyde-) fixed embryonic brains are washed with phosphate buffer (3)/ water (1) and postfixed with 1.5% potassium hexocyano ferrate (II), 1% osmium tetroxide for 30 mine1 h at room temperature in the dark. This staining cocktail will bind preferentially to the unsaturated fatty acids of the membranes. The hexocyanoferrate-reduced osmium will therefore stress membranes at the expense of cytoplasmic structures (ribosomes). In case visibility of cytoplasmic features is desired, plain osmium tetroxide should be used. After water washes, the sections are incubated with 0.2% tannic acid (which will further enhance the membrane contrast by binding to carbohydrate moieties (Simionescu & Simionescu, 1976)) for 10 min at room temperature, followed again by washes in water. The slices are stained overnight in the cold with 0.5% uranyl acetate in 25% methanol (for better penetration). After washing with water, the sections are dehydrated through a graded series of ethanol. As a final dehydration step, three washes with water-free (stored over molecular sieve) ethanol should follow. Next, sections are infiltrated in two steps with increasing concentration of resin (Epon replacement) versus ethanol (1:2, 2:1) for 1 h each. Slices are infiltrated in pure resin overnight. After another change of resin, they are flat embedded between a Teflon-coated microscope slide and a sheet of Aclar plastic foil (cut to the size of a slide), which can easily be removed after polymerization. Two strips of Aclar (ca. 200 mm thick, 1 mm wide) are used as spacers to prevent squeezing of the slices (as depicted in Figure 4(A3)). The glass-resin/tissue-Aclar sandwich is put into the oven, covered with additional microscope slides for weights, and polymerized at 60  C at least overnight. The polymerized samples are peeled off from Aclar or slide, and small pieces (0.5e1 mm2) of the brain region of interest are cut with a fresh razor blade (Figure 4(A3e4); avoid electrostatic “jumps” of the small resin pieces!). Avoid bending the blade edge while cutting, but rather “chop” the pieces without force or tilting. To avoid lifting (and losing) the tiny samples, coat the surface of an empty resin block with some glue and invert it onto the sample for mounting. As glue, unpolymerized resin (kept frozen as a small aliquot in a micro-vial from the embedding day) or super-glue can be used (Figure 4(A5), left). After another day of polymerization in the oven (in upright position), excess resin can be trimmed away with a razor blade with special attention to clean top and bottom edges. The sample is cut in an ultra-microtome to ribbons of 50e70-nm-thick sections that are collected on Formvar-coated slot grids and poststained (standard protocol) with uranyl acetate and lead citrate prior to observation on a standard room temperature TEM. Examples of apical and BL cilia are shown in Figure 4(B). For finer details and analysis of the 3D arrangement, tomographic tilt series through 300-nm sections at high tension (300 kV) can be performed (as described elsewhere (McEwen & Marko, 2001; O’Toole, Winey, McIntosh, & Mastronarde, 2002)). Cilia on a well-defined position (e.g., protruding into the ventricular lumen) are relatively easy to spot in the conventional TEM sections. However, spotting cilia within the tissue can be very time consuming. In addition, an analysis of

4. Methods

the position and structure of the cilia in a more quantitative manner by conventional TEM is very difficult due to the limited field of view at the appropriate TEM magnification.

4.4 VISUALIZATION OF NEURAL PROGENITOR PRIMARY CILIA BY SBF-SEM Electron microscopical analysis of cilia in the tissue context requires good preservation of the tissue, sufficient resolution for fine structure, as well as a large field of view for orientation. Only with the help of the third dimension is it possible to judge whether a cilium is truly located to the basolateral plasma membrane or rather in an intracellular membrane vesicle (or a ciliary pocket). Although it is feasible to solve this question by serial TEM sections for a single cilium, doing a statistical analysis is close to impossible. The localization of a 1-mm-long non-apical cilium within a >100-mm-long, slender and curving BP is an example of a very challenging task due to the large dimensions, and would certainly require extensive three-dimensional imaging. In recent years, large-scale 3D methods have become quite popular. They involve a combination of cutting and SEM imaging (for a comparison see Briggman and Bock (2012)). These methods comprise array tomography (Hayworth, Kasthuri, Schalek, & Lichtman, 2006; Horstmann, Korber, Satzler, Aydin, & Kuner, 2012; Micheva & Smith, 2007), FIB (focussed ion beam)-milling (Knott, Marchman, Wall, & Lich, 2008), as well as serial block face-microtomy (Denk & Horstmann, 2004) that has pioneered the field. In the first case, the sections are cut and automatically collected on wafers or slides outside the SEM. For the latter two methods, the samples are directly sectioned within the electron microscope and the freshly exposed block surface imaged. The cutting by a focussed ion beam (FIB-SEM) or an oscillating diamond knife (SBF-SEM) is a destructive process, where a specific tissue region can only be imaged at the respective time and it is not possible to re-inspect this area at a later time-point. The image data for these 3D-SEM methods are collected from the backscattered electron image and are usually contrast inverted to yield an optical appearance resembling the “well-known” TEM micrographs (examples are given in Figure 4(C)). The 3D methods give an additional clear advantage over conventional serial sections as they are almost perfectly aligned already at acquisition, making it easier to follow a single cell or cilium over multiple sections. However, to yield a resolution that is equal to TEM sections or at least sufficient to identify morphological details, sample preparation has to be adjusted. The goals are (1) highest possible resolution, achieved by as little penetration of the electron beam into the block as possible (i.e., low accelerating voltage of the electrons) and by good cutting properties of the sample, (2) nevertheless a sufficient signal-to-noise ratio and (3) as little charging of the sample by trapped electrons as possible. To achieve these goals, the sample has to be strongly impregnated with heavy metals. SBF-SEM samples from embryonic mouse telencephalon are embedded roughly as described for conventional TEM imaging (depicted in Figure 4(A1e5)).

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However, several additional contrasting steps (based on a protocol (“OTO”) from the Ellisman lab: ncmir.ucsd.edu/sbfsem-protocol.pdf) have been introduced to reduce charging and to improve the signal-to-noise ratio (“OTOTO”). Consecutive addition of reduced osmium tetroxide layers will result in a strong membrane contrast as relatively little cytoplasmic features (e.g., ribosomes) are preserved. As a disadvantage, the heavy metal coat may mask fine details, and it renders the membranes more brittle, sometimes resulting in cracks in the membrane appearance. For some sample, less membrane contrast can be applied (OTO) (Hanker, Deb, Wasserkrug, & Seligman, 1966), depending on the abundance of empty resin spaces (e.g., ventricular lumen, blood vessels) prone to charging, the membrane composition (neuronal membranes stain better with osmium tetroxide than progenitor membranes), or the location of the cilia aimed for (ventricle or within the tissue).

4.4.1 SBF-SEM contrasting In detail, the embedding procedure consists of a primary osmication step (2% osmium tetroxide/1.5% potassium hexacyano ferrate) for 30 mine1 h at room temperature in the dark, then an enhancement step with 1% thiocarbohydrazide (TCH) (Hanker et al., 1966) for 10e20 min in water (dissolved at 60  C) or alternatively 0.2% tannic acid for 10 min (in case cytoplasmic structures in addition to membranes should be visible). This is followed by another osmication step (1% OsO4, 30 min, room temperature), succeeded by a second round of 1% TCH enhancement for 20 min. A final incubation in 1% osmium tetroxide for 30 min at room temperature is added. After the heavy metal treatment, the samples are thoroughly washed with water to get rid of excess stain trapped in the tissue or on the walls of the dish (at least 3  10 min water). This is followed by a contrasting step with 0.5% uranyl acetate in 25% methanol (for better penetration into the tissue) overnight at 4  C. After several washes in water, en bloc staining with lead aspartate is performed for 30 min at 60  C.

4.4.2 SBF-SEM embedding Samples are rinsed with water prior to dehydration with a graded series of ethanol. Resin infiltration is performed with two intermediate steps (1:2, 2:1 mixtures with ethanol) and pure resin overnight. After overnight incubation with the unpolymerized resin, another change of resin is recommended. Finally, the slices are embedded between a Teflon-coated slide and an Aclar coverslip with 200-mm spacers (cut from Aclar sheet) and polymerized at 60  C. The resin to be used is a hard mixture of Durcupan that gives slightly better contrast, or EPON replacement that is less viscous than Durcupan and therefore easier to handle, especially for fragile vibratome sections (Kushida, 1964; Luft, 1961). Both resins have been developed for stability in the electron beam; nevertheless after longer beam exposure (e.g., for focussing) cutting artifacts may be observed due to beam damage of the resin.

4. Methods

4.4.3 Checking and mounting of SBF-SEM samples Prior to serial block face scanning, the sample quality should be checked in the TEM. The sample is directly mounted either on the 3View microtome pins (Figure 4(A5), right) or on a blank resin stub (using the hard EPON mixture used for embedding as a glue). In case of a precious sample, it may be sufficient to mount and section an adjacent tissue region for the TEM investigation. The sample is trimmed and 70-nm sections cut as for standard plastic samples and viewed in the TEM without additional contrasting. If the sample quality is appropriate (high contrast even at low magnification; good quality of structural preservation), a resin-block-mounted sample can carefully be removed from the stub with a razor blade and remounted on the 3View sample pins, again using hard EPON as glue. Alternatively, a conductive resin (CW2400, containing silver flakes) may be used for glueing, although this should not be included in the final sectioned area to prolong the lifetime of the diamond knife. This step is redundant for samples directly mounted on the SEM pins. For better conductivity, the sample on the 3View pin should be orientated such that some tissuecontaining resin (rather than empty resin) makes contact to the pin surface/glue. Small sample dimensions (0.2e0.5 mm in square) are better for good cutting properties in the 3View microtome, although bigger samples are possible. After mounting on the pin, the sample is trimmed at the ultra-microtome with the edge of a glass knife at roughly 90 angles. Since sections will be discarded at cutting and no ribbons are needed, the trimming can be less precise than for serial TEM sectioning. The 3View stage does not allow tilting of the sample, so, already at the ultra-microtome, the sample surface should be trimmed without inclination. As a last trimming step, the surface of the sample can be cut with a diamond knife. This will greatly facilitate alignment in the 3View microtome (by light reflection). For reduction of surface charging, the trimmed sample can be coated with a thin layer of carbon (light brown color on an adjacent filter paper) by evaporation. A coating with silver paint (as used for standard SEM) can serve the same purpose, however can damage the diamond knife.

4.4.4 SBF-SEM investigation At the 3View microtome, mounting and alignment of the sample with respect to the diamond knife should be done as precisely as possible to prevent knife damage and to save time in approaching. The SEM beam should be well aligned prior to imaging. Check the cross-over and astigmatism and select the appropriate detector (backscattered electron detector). Trial scans should be performed for optimization of the accelerating voltage (1e2 kV), current (0.05e0.2 nA) (or spot size), dwell time of the beam (speed of imaging) (1e5 ms/px), pixel size (minimum ca. 5 nm/px), and section thickness (down to 30 nm; however the “useful” thickness should be kept in mind: in which volume a structural change is expected, to avoid generating redundant data). All settings are microscope-dependent. If possible, the chamber pressure (vacuum) should be adjusted for the best signal-to-noise ratio at the required image dimension.

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Once the settings are optimized, the image stack (single frame or montage) can be acquired (for examples see Figure 4(C)). In particular during the first phase of the scan, images should be checked for signs of section debris (dark patches on images) or uneven sectioning, since the “constant” beam exposure may soften the resin and impair cutting properties. In case the sectioning does not stabilize within the first 20 sections, a change of the settings for acquisition should be considered. Focus and astigmatism, as well as the brightness and contrast settings of the detector, have to be checked at intervals, and readjusted if required (depending on the microscope stability).

4.4.5 SBF-SEM image analysis Sections acquired by Digital Micrograph software (Gatan) at the SBF-SEM will show good alignment already, but it is not sufficient for modeling of structures. The TrakEM2 plugin in the ImageJ-based free software Fiji (Cardona et al., 2012; Schindelin et al., 2012) allows one to conveniently import the image data and montages (3View Import List plugin by Nuno Goncalo Diaz, Portugal) into a virtual stack for fast viewing, to align and stitch the sections with SIFT parameters (Saalfeld, Cardona, Hartenstein, & Tomancak, 2010), and to track and annotate features. The information that can “at a glance” be gathered from these SBF-SEM image stacks is amazing, especially in comparison to the tedious analysis of conventional serial TEM sections. This allows a quantitative and statistical analysis even of rare structures (e.g., ciliary remnants in mitotic cells (Paridaen et al., 2013)). However, the computer memory consumed in a relatively short acquisition time can be huge and means of handling and storing of these large data volumes (external hard drives or server solutions) should be considered in advance. A better nominal z-resolution of the SBF-SEM, which is the sectioning thickness with the oscillating knife (w30 nm minimum), can be achieved than with conventional TEM sections (w50 nm). For the effective z-resolution, it has to be kept in mind that the beam penetration (depending on the accelerating voltage) may cause backscattered electron signals to emanate from a larger z-depth than the thickness of the sections. The effective xy-pixel resolution in the SBF-SEMddetermined by the fixed pixel number of the detector for a given field of view, the signal-to-noise ratio, and the masking by the heavy metal coatdis usually lower than for TEM sections (compare Figure 4(B) and (C)). Therefore, for the investigation of structural aspects of cilia within a tissue, a combination of conventional TEM sections, TEM tomography, and SBF-SEM (or one of the other 3D-SEM techniques) may be ideal.

4.5 VISUALIZATION OF NEURAL PROGENITOR PRIMARY CILIA BY TOKUYASU CRYOSECTIONING AND CLEM The methods described so far visualize subcellular details; however, no information about the molecular composition of these structures is provided. This information can be added by the comparison of the EM data with light microscopic data collected by immunostaining and time-lapse imaging, or by immunolabeling for EM.

4. Methods

For the localization of proteins of interest on the cilia by immuno EM, two general methods are available: (1) labeling prior to embedding or (2) labeling after sectioning of embedded samples. The first method involves a protocol similar to light microscopic staining and therefore yields comparable labeling intensities, but results in a poor structural preservation due to the need of permeabilization for antibody penetration (Humbel, de Jong, Muller, & Verkleij, 1998). The latter method (on-section labeling) can be applied either on plastic-embedded samples (as described elsewhere (Schwarz & Humbel, 2014)) or on sucrose-embedded Tokuyasu cryosections (Liou, Geuze, & Slot, 1996; Tokuyasu, 1973). Labeling of (methacrylate) plastic sections is convenient, even for serial sections, but has the limitation that many epitopes (up to 90%) are destroyed or masked by the resin embedding. The Tokuyasu cryosection approach gives much higher success in labeling. It is widely applied for tissue culture studies, where sectioning is relatively easy, but should be described here in some detail for the embryonic mouse brain, which provides challenges if a good orientation of the highly polarized epithelial cells is aimed for (Figure 5(A)). Both methods for on-section immuno-EM-labeling can be correlated with light microscopic stainings, either on (almost) adjacent sections (Fabig et al., 2012; Schwarz & Humbel, 2014) or on the same section mounted on an EM grid (Cortese, Diaspro, & Tacchetti, 2009; Mironov & Beznoussenko, 2012; Sjollema, Schnell, Kuipers, Kalicharan, & Giepmans, 2012; Vicidomini et al., 2008, 2010). The CLEM approach allows the fast identification of structures (e.g., cilia) of interest by fluorescent LM at a superb z-resolution. The z-resolution in CLEM equals the section thickness (50e200 nm) or even less, as antibodies bind only to the surface of plastic sections (Stierhof, Schwarz, & Frank, 1986). This resolution can be reached neither by confocal nor by high-resolution LM imaging. After the LM evaluation, the fine details of the LM-identified structures can be analyzed by TEM. The CLEM approach will greatly facilitate locating a structure of interest (e.g., cilium) within the section at high EM magnification with a limited field of view. For judging the labeling, it has to be kept in mind that the antibodies will not fully penetrate into the sections: fine details (e.g., distal appendages) visible within the section may not be labeled, because the structure is located below the surface of the section (Stierhof et al., 1986).

4.5.1 Preparation of Tokuyasu samples Details of the Tokuyasu sectioning of the embryonic mouse brain are as follows: 200-mm vibratome sections of PFA-fixed embryonic mouse brains are prepared (Figure 5(A1)). The region of interest (dorsolateral telencephalon) is cut with a sharp scalpel blade from the vibratome section and carefully transferred (between the tips of forceps without completely closing) into a small droplet of phosphate buffer on a parafilm square in a humid chamber. Several pieces of tissue can be collected in one droplet. The humid chamber with the tissue pieces is put into a 37  C incubator for 3 min before replacing the buffer with prewarmed 12% gelatine solution in buffer (about 100 mL). After a short incubation in the gelatine solution, the droplet is

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(A)

(B)

FIGURE 5 Cilia investigation by correlative light and transmission immuno EM. (A) Schematic drawing of the crucial steps (1e8, indicated in italics on top) in sample preparation for correlative light and transmission immuno EM on Tokuyasu cryosections. Details are explained in the main text. (B) Example of CLEM at a mouse dorsolateral telencephalon at E13.5. Top left panel,

4. Methods

covered by a second parafilm square with very gentle squeezing, so that a thin layer (0.3e0.7 mm thick) of gelatine is formed between the parafilm sandwich (as depicted in Figure 5(A2)). After hardening the gelatine in the cold, the specific tissue region is cut with a sharp scalpel blade from the gelatine in a trapezoid shape that later allows easy identification of the top edge of the sample for microtome sectioning at the small side of the trapeze (Figure 5(A2), left). The gelatine-tissue-trapeze is next transferred into 2.3 M sucrose solution in phosphate buffer (in a transparent reaction vial, or in a slip-on lid in a humid chamber). For better cutting properties, polyvinylpyrrolidone can be added to the sucrose solution (at 15%), but then additional care has to be taken not to let the sample surface dry out. Sucrose incubation time is in the range of 2e5 h at 4  C (Figure 5(A3)). Longer incubation is not recommended since this may render samples more brittle for cutting. Samples are removed from the sucrose solution between the shanks of forceps and placed in the correct orientation (long side of the trapeze facing down) onto the microtome cryo-pin. The sample is lifted up again and put on a filter paper triangle, before placing it back onto the cryo-pin. In this way, enough sucrose is drained off, not to have a brittle support, but still enough sucrose will remain to act as good glue. The mounted sample is quickly frozen (by swirling it with cryoforceps) into liquid nitrogen, wherein it can be stored for years (Figure 5(A4)).

4.5.2 Cryosectioning Cutting at the ultra-cryo-microtome is performed as described previously (Liou et al., 1996; Tokuyasu, 1973). In summary, sections are trimmed at
90  C and ultrathin sections are cut at
110 to
100  C (Figure 5(A5)). Sections are picked up with a 1:1 mixture of 2.3 M sucrose and 2% methylcellulose in a 3-mm metal loop as soon as possible or just prior to the freezing point of the droplet (Figure 5(A6)). For a light microscopic investigation at high z-resolution, the sections (70e400 nm; 150 nm is our preferred thickness for a high z-resolution with good cutting properties) are collected on 12-mm round coverslips. For CLEM or pure TEM

=

LM image of the ventricular side showing mitotic (m) DNA figures (DAPI, blue) and immunofluorescence for Arl13b (red). The arrowheads indicate Arl13b immunoreactivity, and the red arrowhead indicates the Arl13b immunofluorescence of the basolateral cilium shown at a higher (TEM) magnification to the right. Bottom left panel, TEM of the identical position at the ventricle (v) as shown in the immunofluorescence above as can be judged from the arrangement of the mitotic nuclei (m). The red arrowhead indicates the identical position as in the light microscopic image. Middle panel, high magnification of the Arl13bimmunogold-labeled basolateral cilium indicated by the red arrowheads in the left panels. Right panel, Arl13b-immunogold-labeled apical primary cilium. 10-nm gold immunoreactivity of Arl13b is highlighted by black arrowheads. Scalebar, left panel: 10 mm; middle and right: 200 nm. (See color plate)

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investigation, the sections (70e90 nm) are collected on Formvar- and carbon-coated 100-mesh copper grids, which are mounted on a microscope slide covered with a paper sticker (Figure 5(A7)). This soft support for the grids will allow easy detachment of the grids with the attached sections/sucrose from the support prior to labeling and is useful for marking individual grids (e.g., sample and quality). Glow discharging of the grids may be good, but should not be performed directly before the cutting as otherwise the sucrose droplet may spread over the rim of the grid. Commercial finder grids may be useful for a precise correlation of structures between EM and LM. However, in our hands the arrangement of the tissue sections or folds with respect to the grid bars, together with the pattern of the DAPI-stained nuclei, are sufficient landmarks to correlate the LM/EM structures. Sections can be immunolabeled either directly after sectioning or after storage in the refrigerator for up to several months (Griffith & Posthuma, 2002). Sections on coverslips can be conveniently stored by fitting seven of them into the lid of a 3-cm Petri dish.

4.5.3 Immunolabeling Prior to immunolabeling the gelatine layer on the surface of the tissue is removed by incubating the sections floating on PBS at 37  C for 30e45 min. This is essential for observation of structures on the tissue surface (e.g., cilia protruding into the ventricular lumen), but may be redundant for the observations of structures within the tissue where the gelatine has not penetrated. Sections on coverslips are labeled (section-side facing up, 50 mL droplets for washing steps, 20 mL of diluted antibodies) on parafilm after drawing a waterrepelling circle around the sections (wax-pen) (Figure 5(A8)). Sections on EM grids are labeled floating on droplets (20e50 mL for washing steps, 5 mL of diluted antibody) on parafilm. To prevent formation of auto-fluorescent salt crystals on accidently submerged grids, these are dipped as soon as possible into distilled water and gently dried on their rear-side with filter paper. Primary antibodies have to be used at a higher concentration for EM than for LM stainings, although the volume used is minimal (5e20 mL; use 10 the LM concentration for a first trial). All incubation steps should be carried out in a humid chamber to prevent evaporation. For CLEM, sections on EM grids can be immunolabeled with both fluorescent and gold-conjugated secondary antibodies (or Protein A-gold). For abundant epitopes, a simultaneous incubation with fluorescent and gold antibodies can be performed; however, for weaker epitopes it is recommended to label first with the gold-conjugated secondary antibody, wash the sections, and after a fixation step (1% GA for 2 min) incubate with the fluorescent secondary antibody. Goldconjugated antibodies show a lower affinity for epitopes than antibodies carrying a small fluorescent tag. This lower affinity is explained by reduced antibody access to the epitope due to steric hindrance by the large gold particles (5e15 nm) and by mutual repulsion between the slightly negatively charged gold particles. The fluorescent label will highlight structures, which would have been overlooked or judged as sparse background staining in the EM labeling. Another labeling option is to detect

4. Methods

the primary antibody with a fluorescent secondary antibody as a first step, and subsequently detect the immunoglobulin species of the fluorescent antibody by a gold-conjugated antibody (or Protein A). Although unspecific staining by the fluorescent antibodies will be enhanced as well, this approach will result in an equally high gold-labeling intensity as the fluorescence. CLEM sections or sections on coverslips are counterstained with 0.01 mg/mL DAPI (final) in the secondary antibody solution, which is important both for orientation and for LMeEM correlation. To remove any traces of buffer from their rear-side, coverslips are quickly dipped into a beaker with distilled water after immunolabeling. For mounting, the coverslip is placed section-side down on a 5-mL droplet of Mowiol on a microscope slide. Avoid air bubbles being trapped by lowering the coverslip from one side. Make sure that the coverslip is mounted completely flat, since any inclination will be reflected in a difference of the focal plane at the LM.

4.5.4 CLEM investigation After immunolabeling, CLEM sections are mounted between a microscope slide and a coverslip with 50% glycerol in PBS and are directly viewed in an epifluorescence LM. Although mounting of the grids with the section-side up seems most logical for LM investigation, we prefer the opposite orientation, showing only the fraction of the tissue between the grid bars that will be visible in the EM. Since the fluorescence contained in the ultrathin sections is quite low, care should be taken not to bleach the fluorescence prior to image acquisition (Figure 5(B), top left). After LM investigation, the grid is removed from the slide (submerging it in a 10-cm Petri dish with distilled water will reduce shear forces when lifting the coverslip) and carefully washed with water (5). The rear-side is dried off with filter paper. The section-side of the grids is incubated on droplets of a 1.8% methylcellulose, 0.3% uranyl acetate mixture on parafilm spread on a cold metal plate on ice for 10 min. The contrasting step is finalized by draining excess staining solution at a 45 angle on filter paper, holding the grid in a 3.3-mm loop of wire (as described in Liou et al. (1996)). The uranyl acetate/methylcellulose incubation will result in a combination of negative contrast (electron-translucent membranes) and positive contrast (electron-dense proteins and DNA/RNA). The completely dried grids can be carefully removed from the loops by pricking the methylcellulose film surrounding the grid with closed forceps and subsequently viewed in the TEM (Figure 5(B), bottom left, and right). The immunolabeling of large tissue sections may not always yield as good an ultrastructure of individual cilia as conventional TEM sections; however, a comparison between the (CLEM-)labeled structures and the high-resolution structure of the plastic sections will reveal further details. In some cases, even the correlation of a fluorescent label (antibody or endogenous FP-tag) with an unlabeled ultrastructure within the same section will provide additional information. This may be applicable to rare epitopes, which can be detected by the more powerful fluorescent antibodies but not by the lower labeling intensity of the immuno-gold antibodies.

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CONCLUSION In recent years, advances in, and combination of, imaging techniques at both the LM and EM level have resulted in remarkable gain in knowledge about primary ciliary structure, function, and dynamics. In the future, we expect that development of, for instance, correlation of 3D EM structure and immunolabeling (as is already possible with array tomography) will provide further advances in the field.

ACKNOWLEDGMENTS We thank Marta Florio who initially set up and optimized the dissociation procedure, and Miguel Turrero Garcı´a for his advice on organotypic slice culture and live imaging. Special thanks go to Jula Peters for excellent technical support and critical reading of the manuscript. We thank the MPI-CBG facilities for their essential contribution in providing excellent animal care and state-of-the-art microscopes.

REFERENCES Arai, Y., Pulvers, J. N., Haffner, C., Schilling, B., Nu¨sslein, I., Calegari, F., et al. (2011). Neural stem and progenitor cells shorten S-phase on commitment to neuron production. Nature Communications, 2, 154. Berbari, N. F., Johnson, A. D., Lewis, J. S., Askwith, C. C., & Mykytyn, K. (2008). Identification of ciliary localization sequences within the third intracellular loop of G proteincoupled receptors. Molecular Biology of the Cell, 19(4), 1540e1547. Briggman, K. L., & Bock, D. D. (2012). Volume electron microscopy for neuronal circuit reconstruction. Current Opinion in Neurobiology, 22(1), 154e161. Cardona, A., Saalfeld, S., Schindelin, J., Arganda-Carreras, I., Preibisch, S., Longair, M., et al. (2012). TrakEM2 software for neural circuit reconstruction. PloS One, 7(6), e38011. Caspary, T., Larkins, C. E., & Anderson, K. V. (2007). The graded response to Sonic Hedgehog depends on cilia architecture. Developmental Cell, 12(5), 767e778. Cortese, K., Diaspro, A., & Tacchetti, C. (2009). Advanced correlative light/electron microscopy: current methods and new developments using Tokuyasu cryosections. The Journal of Histochemistry and Cytochemistry, 57(12), 1103e1112. Denk, W., & Horstmann, H. (2004). Serial block-face scanning electron microscopy to reconstruct three-dimensional tissue nanostructure. PLoS Biology, 2(11), e329. Fabig, G., Kretschmar, S., Weiche, S., Eberle, D., Ader, M., & Kurth, T. (2012). Labeling of ultrathin resin sections for correlative light and electron microscopy. Methods in Cell Biology, 111, 75e93. Farkas, L. M., & Huttner, W. B. (2008). The cell biology of neural stem and progenitor cells and its significance for their proliferation versus differentiation during mammalian brain development. Current Opinion in Cell Biology, 20(6), 707e715. Florio, M., & Huttner, W. B. (2014). Neural progenitors, neurogenesis and the evolution of the neocortex. Development, 141(11), 2182e2194.

References

Follit, J. A., Tuft, R. A., Fogarty, K. E., & Pazour, G. J. (2006). The intraflagellar transport protein IFT20 is associated with the Golgi complex and is required for cilia assembly. Molecular Biology of the Cell, 17(9), 3781e3792. Griffith, J. M., & Posthuma, G. (2002). A reliable and convenient method to store ultrathin thawed cryosections prior to immunolabeling. The Journal of Histochemistry and Cytochemistry, 50(1), 57e62. Guadiana, S. M., Semple-Rowland, S., Daroszewski, D., Madorsky, I., Breunig, J. J., Mykytyn, K., et al. (2013). Arborization of dendrites by developing neocortical neurons is dependent on primary cilia and type 3 adenylyl cyclase. The Journal of Neuroscience, 33(6), 2626e2638. Ha¨ndel, M., Schulz, S., Stanarius, A., Schreff, M., Erdtmann-Vourliotis, M., Schmidt, H., et al. (1999). Selective targeting of somatostatin receptor 3 to neuronal cilia. Neuroscience, 89(3), 909e926. Hanker, J. S., Deb, C., Wasserkrug, H. L., & Seligman, A. M. (1966). Staining tissue for light and electron microscopy by bridging metals with multidentate ligands. Science, 152(3729), 1631e1634. Hayat, M. (2000). Principle and techniques of electron microscopy: Biological applications (4th ed.). Cambridge University Press. Hayworth, K. J., Kasthuri, N., Schalek, R., & Lichtman, J. W. (2006). Automating the collection of ultrathin serial sections for large volume TEM reconstructions. Microscopy and Microanalysis, 12(Suppl. 2), 86e87. Higginbotham, H., Eom, T. Y., Mariani, L. E., Bachleda, A., Hirt, J., Gukassyan, V., et al. (2012). Arl13b in primary cilia regulates the migration and placement of interneurons in the developing cerebral cortex. Developmental Cell, 23(5), 925e938. Horstmann, H., Korber, C., Satzler, K., Aydin, D., & Kuner, T. (2012). Serial section scanning electron microscopy (S3EM) on silicon wafers for ultra-structural volume imaging of cells and tissues. PloS One, 7(4), e35172. Humbel, B. M., de Jong, M. D., Muller, W. H., & Verkleij, A. J. (1998). Pre-embedding immunolabeling for electron microscopy: an evaluation of permeabilization methods and markers. Microscopy Research and Technique, 42(1), 43e58. Knott, G., Marchman, H., Wall, D., & Lich, B. (2008). Serial section scanning electron microscopy of adult brain tissue using focused ion beam milling. The Journal of Neuroscience, 28(12), 2959e2964. Kushida, H. (1964). Improved methods for embedding with Durcupan. Journal of Electron Microscopy, 13(3), 139e144. Lehtinen, M. K., Zappaterra, M. W., Chen, X., Yang, Y. J., Hill, A. D., Lun, M., et al. (2011). The cerebrospinal fluid provides a proliferative niche for neural progenitor cells. Neuron, 69(5), 893e905. Liou, W., Geuze, H. J., & Slot, J. W. (1996). Improving structural integrity of cryosections for immunogold labeling. Histochemistry and Cell Biology, 106(1), 41e58. Louvi, A., & Grove, E. A. (2011). Cilia in the CNS: the quiet organelle claims center stage. Neuron, 69(6), 1046e1060. Luft, J. H. (1961). Improvements in epoxy resin embedding methods. The Journal of Biophysical and Biochemical Cytology, 9, 409e414. McEwen, B. F., & Marko, M. (2001). The emergence of electron tomography as an important tool for investigating cellular ultrastructure. The Journal of Histochemistry and Cytochemistry, 49(5), 553e564.

127

128

CHAPTER 6 Analysis of primary cilia in the developing mouse brain

Micheva, K. D., & Smith, S. J. (2007). Array tomography: a new tool for imaging the molecular architecture and ultrastructure of neural circuits. Neuron, 55(1), 25e36. Mironov, A. A., & Beznoussenko, G. V. (2012). Correlative light-electron microscopy a potent tool for the imaging of rare or unique cellular and tissue events and structures. Methods in Enzymology, 504, 201e219. Nigg, E. A., & Stearns, T. (2011). The centrosome cycle: centriole biogenesis, duplication and inherent asymmetries. Nature Cell Biology, 13(10), 1154e1160. O’Toole, E. T., Winey, M., McIntosh, J. R., & Mastronarde, D. N. (2002). Electron tomography of yeast cells. Methods in Enzymology, 351, 81e95. Paridaen, J. T., Wilsch-Bra¨uninger, M., & Huttner, W. B. (2013). Asymmetric inheritance of centrosome-associated primary cilium membrane directs ciliogenesis after cell division. Cell, 155(2), 333e344. Pedersen, L. B., Veland, I. R., Schroder, J. M., & Christensen, S. T. (2008). Assembly of primary cilia. Developmental Dynamics, 237(8), 1993e2006. Rohatgi, R., Milenkovic, L., & Scott, M. P. (2007). Patched1 regulates hedgehog signaling at the primary cilium. Science, 317(5836), 372e376. Saalfeld, S., Cardona, A., Hartenstein, V., & Tomancak, P. (2010). As-rigid-as-possible mosaicking and serial section registration of large ssTEM datasets. Bioinformatics, 26(12), i57ei63. Saito, T., & Nakatsuji, N. (2001). Efficient gene transfer into the embryonic mouse brain using in vivo electroporation. Developmental Biology, 240(1), 237e246. Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., et al. (2012). Fiji: an open-source platform for biological-image analysis. Nature Methods, 9(7), 676e682. Schwarz, H., & Humbel, B. M. (2014). Correlative light and electron microscopy using immunolabeled sections. Methods in Molecular Biology, 1117, 559e592. Shitamukai, A., Konno, D., & Matsuzaki, F. (2011). Oblique radial glial divisions in the developing mouse neocortex induce self-renewing progenitors outside the germinal zone that resemble primate outer subventricular zone progenitors. The Journal of Neuroscience, 31(10), 3683e3695. Simionescu, N., & Simionescu, M. (1976). Galloylglucoses of low molecular weight as mordant in electron microscopy. I. Procedure, and evidence for mordanting effect. The Journal of Cell Biology, 70(3), 608e621. Sjollema, K. A., Schnell, U., Kuipers, J., Kalicharan, R., & Giepmans, B. N. (2012). Correlated light microscopy and electron microscopy. Methods in Cell Biology, 111, 157e173. Sorokin, S. (1962). Centrioles and the formation of rudimentary cilia by fibroblasts and smooth muscle cells. The Journal of Cell Biology, 15, 363e377. Sotelo, J. R., & Trujillo-Cenoz, O. (1958). Electron microscope study on the development of ciliary components of the neural epithelium of the chick embryo. Zeitschrift fur Zellforschung und Mikroskopische Anatomie, 49(1), 1e12. Stierhof, Y. D., Schwarz, H., & Frank, H. (1986). Transverse sectioning of plastic-embedded immunolabeled cryosections: morphology and permeability to protein A-colloidal gold complexes. Journal of Ultrastructure and Molecular Structure Research, 97(1e3), 187e196. Tabata, H., & Nakajima, K. (2008). Labeling embryonic mouse central nervous system cells by in utero electroporation. Development, Growth & Differentiation, 50(6), 507e511. Tokuyasu, K. T. (1973). A technique for ultracryotomy of cell suspensions and tissues. The Journal of Cell Biology, 57(2), 551e565.

References

Vicidomini, G., Gagliani, M. C., Canfora, M., Cortese, K., Frosi, F., Santangelo, C., et al. (2008). High data output and automated 3D correlative light-electron microscopy method. Traffic, 9(11), 1828e1838. Vicidomini, G., Gagliani, M. C., Cortese, K., Krieger, J., Buescher, P., Bianchini, P., et al. (2010). A novel approach for correlative light electron microscopy analysis. Microscopy Research and Technique, 73(3), 215e224. Walton, J. (1979). Lead asparate, an en bloc contrast stain particularly useful for ultrastructural enzymology. The Journal of Histochemistry and Cytochemistry, 27(10), 1337e1342. Wang, X., Tsai, J. W., Imai, J. H., Lian, W. N., Vallee, R. B., & Shi, S. H. (2009). Asymmetric centrosome inheritance maintains neural progenitors in the neocortex. Nature, 461(7266), 947e955. Wilsch-Bra¨uninger, M., Peters, J., Paridaen, J. T., & Huttner, W. B. (2012). Basolateral rather than apical primary cilia on neuroepithelial cells committed to delamination. Development, 139(1), 95e105. Yeh, C., Li, A., Chuang, J. Z., Saito, M., Caceres, A., & Sung, C. H. (2013). IGF-1 activates a cilium-localized noncanonical Gbetagamma signaling pathway that regulates cell-cycle progression. Developmental Cell, 26(4), 358e368. Yu, Y. C., Bultje, R. S., Wang, X., & Shi, S. H. (2009). Specific synapses develop preferentially among sister excitatory neurons in the neocortex. Nature, 458(7237), 501e504.

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