Inverted selective plane illumination microscopy (iSPIM) enables coupled cell identity lineaging and neurodevelopmental imaging in Caenorhabditis elegans

Abstract

The Caenorhabditis elegans embryo is a powerful model for studying neural development, but conventional imaging methods are either too slow or phototoxic to take full advantage of this system. To solve these problems, we developed an inverted selective plane illumination microscopy (iSPIM) module for noninvasive high-speed volumetric imaging of living samples. iSPIM is designed as a straightforward add-on to an inverted microscope, permitting conventional mounting of specimens and facilitating SPIM use by development and neurobiology laboratories. iSPIM offers a volumetric imaging rate 30× faster than currently used technologies, such as spinning-disk confocal microscopy, at comparable signal-to-noise ratio. This increased imaging speed allows us to continuously monitor the development of C, elegans embryos, scanning volumes every 2 s for the 14-h period of embryogenesis with no detectable phototoxicity. Collecting ∼25,000 volumes over the entirety of embryogenesis enabled in toto visualization of positions and identities of cell nuclei. By merging two-color iSPIM with automated lineaging techniques we realized two goals: (i) identification of neurons expressing the transcription factor CEH-10/Chx10 and (ii) visualization of their neurodevelopmental dynamics. We found that canal-associated neurons use somal translocation and amoeboid movement as they migrate to their final position in the embryo. We also visualized axon guidance and growth cone dynamics as neurons circumnavigate the nerve ring and reach their targets in the embryo. The high-speed volumetric imaging rate of iSPIM effectively eliminates motion blur from embryo movement inside the egg case, allowing characterization of dynamic neurodevelopmental events that were previously inaccessible.

Proper neural circuit assembly requires the coordinated execution of multiple events, including cell migration, axon guidance, and synaptogenesis (1, 2). During neurodevelopment, these events are orchestrated between pre- and postsynaptic partners, resulting in the correct wiring of the nervous system. The mechanisms that enable proper wiring in vivo are not well understood.

Caenorhabditis elegans provides an excellent model to understand how neural circuit assembly occurs in vivo. With only 302 neurons, ∼7,000 synapses, and an available and comprehensive neural connectivity map (3), the nervous system of C. elegans is well characterized and relatively simple. The molecular mechanisms that control neurodevelopmental decisions in the nematode are well conserved throughout evolution (4), and several genetic programs that control terminal differentiation of neuronal identity were first identified and characterized in C. elegans (5, 6). Studies in C. elegans have also significantly contributed to our understanding of neuroblast migration and axon guidance (7, 8). One of the model experimental systems for studying neuroblast migration in nematodes is the pair of canal-associated neurons (CANs) (9–17). Identification of the molecules required for CAN migration also revealed conserved genetic programs required for cellular migration in vertebrates (18, 19). Moreover, the conserved Netrin signaling pathway and its associated receptors (UNC-40/DCC and UNC-5) were first discovered in C. elegans (20–22). In these studies, in vivo labeling and visualization of nematode neurites facilitated identification of downstream molecular factors required for Netrin-dependent guidance (23).

Though genetic studies in C. elegans have revealed the identity of the molecular players required for neurodevelopment, the mechanisms by which conserved genetic programs direct neural circuit formation during embryonic neurulation are less understood. Most neurodevelopmental studies have been conducted in fixed embryos or as end-point genetic analyses in nematode larvae. Real-time inspection of neurulation in the embryo would be of great value in visualizing the neurodevelopmental dynamics that allow innervation of the nematode nervous system. However, directly observing neurodevelopment in the embryo has proven difficult because of fast movement and twitching of the embryo inside the egg shell. Removing this rapid motion with anesthetics is impractical because of the relatively impermeable nature of the egg shell, and conventional 4D microscopy induces too much phototoxicity at the rates and durations necessary for prolonged, blur-free imaging.

Here we report an approach that addresses these problems, and that allows inspection of neurodevelopmental decisions in live nematode embryos. First, we designed iSPIM, an adaptable module that adds plane illumination functionality to any inverted microscope base. This technical advance allowed us to continuously image the nematode embryo, scanning volumes every 2 s for the entire 14 h of embryogenesis with no detectable phototoxicity. Second, we merged two-color iSPIM with automated lineaging techniques to identify cells of interest and visualize their neurodevelopmental dynamics. We investigated four neurons expressing transcription factor CEH-10/Chx10 during early embryogenesis: CANL, CANR, RMED, and ALA. We then examined the cell migration and neurite outgrowth decisions made by these neurons throughout embryogenesis. We found that CANs use somal translocation and amoeboid movement as they translocate to their final position. We also visualized axon guidance and growth cone dynamics as neurites circumnavigate the nerve ring and the body of the worm to reach their targets. Measurement of these events was largely inaccessible before our approach. Our experiments demonstrate the power of iSPIM to facilitate inspection of live neurodevelopmental events, and pave the way for a 4D dynamic neurodevelopmental atlas in C. elegans.

Results

Development of iSPIM.

Most of the neurodevelopmental decisions in C. elegans are made during embryonic neurulation (3, 24). This ontogenic time has remained largely inaccessible for two reasons: (i) Neurulation in the embryo is accompanied by fast movement and twitching, requiring faster volumetric imaging rates to avoid the artifacts induced by motion blur. (ii) Conventional 4D imaging technologies such as point-scanning confocal or two-photon microscopies fail to provide the necessary signal rates. Even if such point-scanning technologies are parallelized, as in spinning-disk or swept-field microscopy, the photodamage induced by continuous high-speed imaging results in developmental defects or arrest. To overcome these hurdles, we sought a fast, minimally invasive microscopy technique suitable for routine use in nematode embryos.

Light sheet-based fluorescence microscopy (LSFM) (25–28) employs parallelized excitation and a perpendicular detection geometry (29) for optically sectioned volumetric interrogation of living samples, thus enabling the study of in toto development (30) or neuronal dynamics (28) at high frame rates while reducing photodamage and photobleaching to levels far below other microscopy techniques. In most implementations (31), the LSFM system is designed around the specimen, requiring novel sample preparation (often embedding the sample in an agarose gel) and precluding the use of conventional sample mounts, such as glass coverslips. We designed iSPIM, a module that offers the advantages of plane illumination while maintaining the flexibility and sample geometry of commercially available inverted microscopes (Fig. 1, Materials and Methods, SI Materials and Methods, and Fig. S1).

Fig. 1.

iSPIM, plane illumination on an inverted microscope base. Two water-immersion objectives are mounted onto a Z translation stage that is bolted directly onto the illumination pillar of an inverted microscope. The demagnified image of a rectangular slit (MASK) is reimaged with lenses (L) and excitation iSPIM objective (EXC OBJ), thus producing a light sheet at the sample (S). For clarity, relay lens pairs between L and EXC OBJ are omitted in this schematic, but are included in Fig. S1. Sample fluorescence is detected (DET OBJ) using appropriate mirrors (M), emission filters (F), lenses (L), and camera. EXC OBJ is fixed in place and the light sheet is scanned through the sample using a galvanometric mirror (not shown). A piezoelectric objective stage (PS) moves DET OBJ in sync with the light sheet, ensuring that detection and excitation planes are coincident. The sample S is mounted onto a coverslip (C) that is placed onto a 3D translation stage, thus ensuring correct placement of S relative to iSPIM objectives. S may also be viewed through objectives (CONV OBJ), dichroic mirrors (DM), and optics in the conventional light path of the inverted microscope.

iSPIM replaces the illumination pillar of an inverted microscope with a mechanical housing for two objectives that illuminate the sample with a light sheet (excitation), and collect the resulting fluorescence (detection). By rapidly translating the light sheet through the sample, the module enables rapid volumetric image collection without requiring specimen movement, similar to previous methods (28). The choice of objectives is mechanically constrained by the need to place them in the 90° orientation required for LFSM, without steric interference with each other or the sample. We thus chose two high NA (0.8), long working distance (3.5 mm), water-immersion (minimizing aberrations due to refractive index mismatch) objectives that satisfied this condition. The objective housing is attached to an automated linear translation stage so that the two objectives may be properly positioned relative to the sample. Samples are mounted on a rectangular coverslip (Fig. S2), and may be translated using an automated 3D mechanical stage and additionally imaged using the conventional light path built into the inverted microscope frame. In the experiments we describe herein, the conventional light path was particularly useful for rapidly finding embryos of appropriate age (e.g., two-cell stage) before iSPIM imaging.

Although iSPIM is compatible with a variety of live samples, we optimized our experiments for C. elegans embryogenesis by focusing the excitation light sheet to a beam waist of ∼1.2 μm at the center of the embryo, thus minimizing diffractive spreading at the edges of our field of view (where we measured a beam waist of ∼3 μm). The resulting spatial resolution, as measured on 100-nm beads embedded in a 2% agarose gel, was 0.52 ± 0.02 μm laterally and 1.70 ± 0.39 μm axially (n = 10 beads, measured over our ∼30 × 50 μm field of view), similar to measurements carried out by others at similar N.A. (32). By rapidly scanning the light sheet in the focal plane (33) (SI Materials and Methods and Fig. S3), we mitigated striping artifacts that result from absorption in the sample plane.

High-Speed iSPIM Imaging of GFP-Histones in C. elegans Embryos from the Two-Cell Stage Until Hatching.

We demonstrated the utility of iSPIM for live samples by imaging transgenic C. elegans embryos labeled with GFP-histone markers. Because C. elegans has an invariant cell lineage, and disruptive phototoxic perturbations do not result in cell lineage compensation, embryos provide a stringent assay for the potentially phototoxic effects of in vivo imaging (34). Furthermore, strains carrying GFP-histone markers have been well characterized by spinning disk microscopy, enabling direct comparison of the potential gains of iSPIM relative to this more workhorse technology (35). As in previous experiments, imaging GFP-histone–labeled C. elegans embryos with spinning-disk confocal microscopy (SDCM) was limited to a rate of ∼1 vol/min from two cells until twitching (Movie S1 and SI Materials and Methods), as significantly higher imaging frequencies or longer durations at this frequency resulted in embryonic arrest due to phototoxicity (36). Imaging the same samples with our iSPIM module allowed continuous volumetric image collection every 2 s from the two-cell stage until hatching (Movie S2), a rate 30× faster than SDCM and close to the hardware limit imposed by our camera. Despite our much higher temporal sampling, we did not detect any obvious abnormalities in the imaged embryos in terms of morphology or in the timing of developmental hallmarks, such as the invariant order and orientation of blastomere divisions, gastrulation, pharyngeal shape, elongation, and twitching. Furthermore, embryos imaged with iSPIM hatched at the expected time (measured temporal variation within 5%, similar to previous studies) (34, 37) into viable larva stage 1 (L1) animals (Fig. 2A and Movie S2). iSPIM thus enabled the noninvasive collection of ∼25,000 volumetric datasets over the entire 14-h period of nematode embryogenesis.

Fig. 2.

High-speed nuclear imaging in the nematode embryo. Embryos with GFP-histones were volumetrically imaged at 30 vol/min from the two-cell stage until hatching. (A) Selected maximum-intensity projections from a representative embryo, highlighting different developmental stages (Movie S2). (Scale bar: 5 μm.) (B) Highlighted nuclei (circled in red, A Top Right) comparing maximum-intensity projections between iSPIM (Left, images sampled every 2 s but shown every 6 s due to space constraints) and spinning-disk confocal microscopy (Right, images sampled every minute, taken on a different embryo).

Despite the 30-fold increase in speed, raw iSPIM images displayed similar signal-to-noise ratio (SNR) to SDCM, and SNR could be further increased with deconvolution (Fig. S4). To assess the value of iSPIM for in toto cell biological studies, we imaged single dividing cells in the intact embryo and compared the spatiotemporal resolution of iSPIM and SDCM (Fig. 2B). Chromosome condensation and the kinetics of cytokinesis were clearly visible using iSPIM, but obscured in similarly nonperturbative SDCM.

Development of Two-Color iSPIM for Coupled Cell Identity Lineaging and Neurodevelopmental Imaging.

During the first 14 h of development, the C. elegans embryo develops from a single-cell zygote to a 558-cell larva through an invariant cell lineage, where lineage identity equates with the fate of the cell (34, 35). Because of the invariant cell lineage of the nematode, C. elegans has long been a workhorse in the identification of transcription factors that instruct neuronal identity and development (5, 6, 38). We reasoned that the imaging rates afforded by iSPIM provided an opportunity to determine nuclear positions while also visualizing the expression pattern of transcription factors; this would allow us to simultaneously achieve two goals: (i) identification of neurons expressing the transcription factor and (ii) visualization of their neurodevelopmental dynamics.

To simultaneously image neurons and nuclei, we developed a worm strain that expressed histone:mCherry under a ubiquitous promoter, and GFP under the CEH-10/Chx10 promoter. The homeodomain transcription factor CEH-10 is a conserved regulatory transcription factor that affects the differentiation and development of a subset of neurons in C. elegans (15, 39, 40). Therefore, ceh-10p:GFP transgenic worms enable visualization of neurons that express this transcription factor. By alternately exciting the embryos with 488-nm and 561-nm lasers and capturing both GFP and mCherry fluorescence, we achieved two-color iSPIM at 12 vol/min (Fig. 3A, Movie S3, and SI Materials and Methods).

Fig. 3.

Dual-color iSPIM and lineaging identifies neuronal cells expressing CEH-10/Chx10. (A) DIC and iSPIM maximum-intensity projections at indicated time points, highlighting ubiquitously expressed histone:mcherry (red) and ceh-10p:GFP (green) at selected times points. iSPIM images are from a representative embryo; DIC pictures were taken on a different animal and are provided only as a reference (Movie S3). (B) Cartoon model to accompany A, also emphasizing location of neurons with respect to the anatomical body plan of the embryo. (C) Computer-derived lineage tree for the identified neurons in A.

Lineage identity of cells expressing ceh-10p:GFP has been previously determined by longitudinal studies in fixed embryos, or by visualizing live embryos at specific time points (39, 40). Using iSPIM, we (i) recorded the dynamic GFP expression pattern of the ceh-10p:GFP strain; (ii) used the histone:mCherry signals to obtain the embryo lineage with the automated lineage tracing software, StarryNite (35, 41) (SI Materials and Methods); and (iii) correlated lineage with cytoplasmic expression pattern to identity the four brightest cells expressing ceh-10p:GFP (Fig. 3C). By 6 h postfertilization (hpf), GFP expression appeared in precursor cell ABalapppaa (bean stage of embryonic development), and subsequently in the daughter cells ALA (ABalapppaaa) and RMED (ABalapppaap; Fig. 3A). GFP expression was also subsequently observed in the canal-associated neurons, CANR (ABalappappa) and CANL (ABalapaaapa; Fig. 3A). The observed pattern of ceh-10p:GFP expression in these neuronal cells, and their subsequent migration through the onset of early twitching (∼7:10:00 hpf, twofold stage), was reproducible across all 12 inspected embryos and is summarized as a cartoon model (Fig. 3B). Having established cell identity and initial migrations, we subsequently used a combination of one- and two-color iSPIM for more detailed analysis of the developmental dynamics of these neurons, from the comma stage through twitching.

CAN Migration.

Most cells in C. elegans undergo short migrations to reach their final positions (34). A few cells, such as the CANs, embark upon long-range migrations during development (Fig. 3B) (13, 34). Previous genetic analysis of CAN migration used the osmoregulatory phenotypes resulting from CAN dysfunction, and end-point analysis of CAN position, to identify molecules required for cell migration (10, 15). These studies resulted in fundamental insights regarding conserved molecular mechanisms of neuronal migration in vivo, further underscoring the value of the nematode system in neurodevelopment and gene discovery. Despite its importance as an experimental system for understanding cell migration, CAN migration in embryos has never been fully described, likely because of the limitations of conventional imaging methods.

Using iSPIM, we visualized CAN development in seven wild-type embryos, and observed that their neurodevelopmental progression is stereotyped across individual animals (Fig. 4A and Movie S4). CANs use multiple strategies as they navigate the different cellular environments in the nematode to reach their target destination. From 6:30:00 to 7:10:00 hpf, CANs slowly migrate longitudinally and posteriorly, traveling ∼10 μm from their birth site where fluorescence was first observed (Fig. 4B). From 6:30:00 to 6:45:00 hpf, migration is passive, likely resulting from cells moving en masse, as the relative positions of the CANs with respect to neighboring nuclei do not significantly change, and their migration rate of 0.15 μm/min is similar to neighboring neurons ALA and RMED (Fig. 4A).

Fig. 4.

Neuronal morphology and migration through early twitching. (A) Selected maximum-intensity projections of ceh-10p:GFP, highlighting migration of neurons after expression of GFP, through onset of early twitching. The outline of the embryo is indicated by the dotted line. (B) Outlines of the migratory path of individual neurons before embryonic twitching. (C) Migration kinetics of CANs from the dataset shown in A, through early twitching. 3D displacements are measured from the anterior tip of the embryo. (D) An example of somal translocation before twitching from a dual-color iSPIM dataset. (E) Higher-magnification view of the blue boxed region in A, showing amoeboid movement of CANs posttwitching as neurons migrate posteriorly. Dual-color images in D were acquired from a representative embryo at 12 vol/min; single-color images for other panels were acquired from another representative embryo at 10 vol/min (Movie S4). In all images, anterior is Left and posterior Right.

From 6:45:00 to 7:10:00 hpf, CANs speed up slightly, moving at an average rate of 0.3 μm/min (Fig. 4C). In this interval immediately preceding twitching, CAN morphology changed from a spherical to a bipolar shape (yellow box in Fig. 4A; also, dual-color example in Fig. 4D), oriented along the anterior/posterior axis and along the direction of migration. The bipolar morphology was characterized by a lamellipodial structure, similar to a pseudopod, in the posterior side of the CANs, and a trailing process in the anterior side of the cell. During this active migratory phase we observed somal translocation of CANs and amoeboid movement (Fig. 4 D and E).

After ∼7:10:00 hpf, CANs preserved their bipolar morphology and appeared to move more rapidly in an amoeboid fashion (Fig. 4E) as they continued their migration toward the posterior end of the embryo (Fig. 4A). This period of active CAN migration coincides with embryonic twitching (jagged lines in Fig. 4C). Despite the large embryonic movements during twitching, we successfully tracked both CANs, finding that cells moved ∼20 μm in 30 min, an average speed of 0.7 μm/min until the end of migration at ∼7:45:00 hpf.

Characterization of Neurite Outgrowth Through Embryonic Twitching.

CAN migration occurs during neurulation. During this same period, most neurons in the nematode simultaneously extend their neurites to reach their target regions (3, 24, 34). Although growth cone morphology in the embryonic ventral nerve cord has been described by fixing embryos at different developmental time points and visualizing axon outgrowth by EM (24), continuous neurite outgrowth during neurulation can now be documented in vivo for C. elegans embryos, along with descriptions of axon outgrowth as neurites circumnavigate the nerve ring.

To examine navigation around the nerve ring, we visualized neurite outgrowth in the ALA neuron (Fig. 5 A and B and Movie S5). Serial EM reconstructions of the nematode nerve ring show that in the adult, the ALA neuron is positioned in the dorsal ganglion behind the nerve ring and has two bilaterally symmetric neurites (3). These neurites enter the nerve ring, circumnavigate it, and exit laterally to project toward the posterior of the animal. Visualization of ALA outgrowth with iSPIM revealed the evolution of this process in vivo, and confirmed that ALA outgrowth was stereotyped across individual animals (four embryos inspected).

Fig. 5.

Neuronal outgrowth through twitching. (A) Selected maximum-intensity projections emphasizing ALA neurite outgrowth from a representative embryo. The outline of the embryo is indicated by the dotted line. The red boxed region highlights ALA (Movie S5). (B) Higher-magnification view. The red dot marks ALA soma; the green star marks the Left neurite outgrowth. (C) Selected projections emphasizing CAN neurite outgrowth, from the same embryo as in A and B (Movie S6). Purple arrows highlight direction of outgrowth. Note that as the embryo is twitching inside the egg case, the relative position of the CANs inside the embryos changes between volumes. High-speed volumetric imaging through iSPIM enabled visualization of outgrowth through twitching with no motion blur. (D) Higher-magnification view, emphasizing growth cones (orange stars). (E) Quantification of anterior extensions (distance from center of cell body to tip of axon) over time, measured from four animals. Data were acquired at 30 vol/min. (F) Higher-magnification view of outgrowth of both anterior (pink arrows) and posterior (red arrows) CAN neurites, from the same embryo as in A–D. The posterior outgrowth became visible ∼12–13 hpf.

ALA neurite extension commenced at ∼6:40:00 hpf. Extension occurred simultaneously in the two bilaterally symmetric neurites extending from the anterior end of the ALA cell body (Fig. 5A). We observed that growth cones project from both left and right sides of the single ALA cell (Fig. 5B). As the neurites grow ventrally, circumnavigating the nerve ring, we found that ALA growth cones display dynamic exploratory behaviors. In all inspected embryos, at ∼7:20:00 hpf, the ALA growth cone bifurcates (Fig. 5B). During bifurcation, extension pauses for ∼10 min, after which the growth cone readopts its unipolar, unbranched morphology and continues navigation. We hypothesize that these bifurcations represent guidance decisions as ALA navigates and turns in the nerve ring. This hypothesis is based on the following observations: (i) pause sites in neurite outgrowth occurred at similar developmental times in different embryos, suggesting they are part of the developmental program; (ii) pause sites occurred at similar locations in the nerve ring of all embryos; and (iii) the locations where growth cones pause and bifurcate correspond to sites where the ALA neuron makes turning decisions in the nerve ring.

We also visualized CAN outgrowth through embryonic twitching (Fig. 5 C and D and Movie S6). CANs cease to actively migrate 8 hpf. During the next hour, CANs retain their bipolar morphology, but do not extend a neurite. However, at 9 hpf, we observed the formation of anterior neurite extensions (Fig. 5C). These extensions persisted throughout the remainder of embryogenesis and eventually developed into growth cones. Growth cone biogenesis was observed as a flattened lamellar structure ∼4-μm thick at the tip of the outgrowing neurite (Fig. 5D, orange stars). Outgrowth of the growth cones reaches speeds of 0.18 μm/min (Fig. 5E) and followed a similar trajectory anteriorly, in contrast to the previous, posterior cell migration trajectory taken by the CAN (Fig. 4). Neurite length in the CANs extended until it reached ∼44 μm and the anterior part of the head of the worm. In addition to the anterior neurite, iSPIM also revealed the outgrowth of the dimmer, posterior CAN neurite (Fig. 5F). Posterior neurite outgrowth became visible ∼12–13 hpf, after anterior neurite extension to the head of the nematode. Although posterior neurite outgrowth commences toward the end of embryogenesis, it does not conclude until after the animal hatches (L1 larva stage).

Discussion

Progress in the field of neurodevelopmental biology has been tightly linked to innovations in imaging technologies, and we believe iSPIM will be of great value in visualizing the neurodevelopmental dynamics that allow innervation of the nematode nervous system. To exemplify the value of iSPIM, we identified and described the cell migration and neurite outgrowth decisions made by a subset of cells, in vivo and with subcellular spatial resolution. CAN neurodevelopment was visualized for ∼7 h from birth near the nose of the worm, subsequent migratory behaviors, and the extension of posterior and anterior neurites. Continuous, volumetric imaging allowed qualitative observations of the cell biological events through cell migration and axon guidance, as well as detailed quantification of the outgrowth kinetics in the twitching embryo. Our high imaging rate also allowed us to visualize ALA neurites as they circumnavigated the nerve ring.

iSPIM allows inspection of developmental events that were previously inaccessible, but significant room for technical improvement remains. Although our light sheet thickness was optimized for C. elegans embryos, resulting in better axial resolution than other experiments on larger samples (27, 30), the sheet was still created from a Gaussian beam. Gaussian beams undergo significant diffraction at increasing distances from the beam waist, coupling the quality of optical sectioning to the field of view and degrading the effective axial resolution at the sample edges. Exciting a fluorescent sample with scanned Bessel beams in an LSFM geometry (42) mitigates this problem and can markedly improve axial resolution, as has been demonstrated on single cells (43). Such a resolution improvement comes at a cost, however, as sidelobes in the Bessel beam profile cause significant out-of-plane illumination. Removing the contaminating effects of the sidelobes requires either a structured illumination approach (necessitating more images per plane, and thus exposing the sample to even more excitation) or multiphoton excitation (potentially leading to nonlinear photobleaching and photodamage).

A better alternative for improving axial resolution may be to collect fluorescence from multiple directions while maintaining the perpendicular LSFM geometry. Fusing the result of datasets taken at each view into a multiview image results in more isotropic resolution (44). This approach has typically been implemented by keeping illumination and detection directions fixed, while rotating the sample so that multiple views may be obtained. Rotating the sample is likely too slow for maintaining the high imaging rates we present here. We note that the benefits of multiview imaging may be realized with iSPIM, and at high speed, by leaving the sample fixed and performing alternate excitation and detection with both LSFM objectives, instead of using one objective for excitation and the other for detection, as presented herein. This scheme would provide two views, but a third could be added by using the conventional objective (Fig. 1) mounted in the inverted microscope base (45).

While inspecting the neurodevelopment of CEH-10–expressing cells, we found that migration and outgrowth are highly stereotyped across embryos. iSPIM in combination with the stereotypy of nematode embryonic development thus provides a unique opportunity to document the 4D wiring decisions of nervous system assembly. Because two-color iSPIM enables simultaneous identification of cells and inspection of their developmental dynamics, understanding the neurodevelopment of specific neurons in the context of neighboring cells and tissues is now possible. Further improving the axial resolution of iSPIM in combination with cell-specific promoters or photoactivatable markers will enable the construction of a 4D dynamic brain atlas in the C. elegans embryo, thus providing a systems-level understanding of neurodevelopment.

Materials and Methods

Worm Strains.

Worms were raised at 20 °C on nematode growth medium seeded with E. coli OP50. Strain BV24 [ltIs44 [pie-1p-mCherry::PH(PLC1delta1) + unc-119(+)]; zuIs178 [(his-72 1kb::HIS-72::GFP); unc-119(+)] V] was used to image nuclei. Strain BV117 [lqIs4 [ceh-10 promoter::GFP + rol-6(su1006)]; itIs37 [pie-1 promoter::mCherry::H2B::pie-1 3′ UTR + unc-119(+)]; stIs10226 [his-72 promoter::HIS-24::mCherry::let-858 3′ UTR + unc-119(+)]] was used for visualizing neuronal cells and processes. Preparation of worm embryos for iSPIM is discussed in detail in SI Materials and Methods.

Instrumentation.

An Olympus IX-81 inverted fluorescence microscope served as the base for iSPIM experiments. We removed the illumination pillar of the microscope and modified it for mounting iSPIM excitation/detection objectives above the sample (Fig. 1). Mounting two high-N.A. objectives in a 90° orientation and in proper placement with respect to the sample is nontrivial, so we designed a custom objective mount for this function. The mount was bolted onto an automated translation stage (LS50; Applied Scientific Instrumentation) for correct positioning of the assembly in Z, and the combined mount/stage was bolted onto the modified illumination pillar. Both the modified illumination pillar and objective housing may be obtained from Applied Scientific Instrumentation. The objective mount also housed a fast piezoelectric objective positioner (PIFOC-P726; Physik Instrumente) that allowed us to move the iSPIM detection objective in sync with the light sheet, thus ensuring coincidence of excitation and detection planes. Further details on iSPIM excitation/detection optics and the power levels used during experiments are covered in SI Materials and Methods.

We also added an automated XY stage equipped with a Z piezo (PZ-2000; Applied Scientific Instrumentation) to the microscope base, for translating the sample to the focal/imaging plane of the iSPIM excitation/detection objectives. Rectangular coverslips containing the sample were placed in an autoclavable stainless steel rectangular chamber with removable bottom (I-3078-2450; Applied Scientific Instrumentation), and sealed in place with an O-ring (1.5-mm thickness, 40-mm inner diameter). This imaging chamber was placed into a temperature-controlled stage insert (Bioptechs, custom design) and the insert mounted to the PZ-2000 stage. Both the insert and imaging chamber were designed to allow unfettered access to both iSPIM objectives and to objectives that mount conventionally into the IX-81. Worm embryos were screened initially using a 10×, 0.3 N.A. air objective (1-U2B524; Olympus) and the 1.6× magnification expander internal to the microscope, using room lighting for illumination and an electron-multiplying CCD (EMCCD) camera (Andor iXon DU-885) mounted to the left side port of the microscope for detection.