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Microfluidic devices have been developed for imaging behavior and various cellular processes in Caenorhabditis elegans, but not subcellular processes requiring high spatial resolution. In neurons, essential processes such as axonal, dendritic, intraflagellar and other long-distance transport can be studied by acquiring fast time-lapse images of green fluorescent protein (GFP)-tagged moving cargo. We have achieved two important goals in such in vivo studies namely, imaging several transport processes in unanesthetized intact animals and imaging very early developmental stages. We describe a microfluidic device for immobilizing C. elegans and Drosophila larvae that allows imaging without anesthetics or dissection. We observed that for certain neuronal cargoes in C. elegans, anesthetics have significant and sometimes unexpected effects on the flux. Further, imaging the transport of certain cargo in early developmental stages was possible only in the microfluidic device. Using our device we observed an increase in anterograde synaptic vesicle transport during development corresponding with synaptic growth. We also imaged Q neuroblast divisions and mitochondrial transport during early developmental stages of C. elegans and Drosophila, respectively. Our simple microfluidic device offers a useful means to image high-resolution subcellular processes in C. elegans and Drosophila and can be readily adapted to other transparent or translucent organisms.
Neurons are highly differentiated cells with long axonal processes that extend from the cell body and end at synapses, structures that are essential for synaptic transmission. Most protein synthesis takes place in the cell body, making axonal transport a vital highway for synaptic development and function. Several cargoes such as synaptic vesicles and mitochondria are transported to synapses by microtubule-dependent molecular motors (1–4). Transport of such cargoes is necessary for synaptic development and function. Transport of cargoes also occurs in dendrites and cilia of neurons and is important for function. An important tool to study the dynamics of neuronal transport is high-resolution time-lapse imaging of fluorescently labelled organelles such as vesicles and mitochondria in culture (5–8), in dissected neurons (9–12) and in vivo(13–18). Axonal transport occurs in the time frame of seconds requiring acquisition rates of 3–8 Hz. There are several cellular processes such as neuronal whole-cell calcium responses to stimulus (19) and endocytic processes (20) that occur at similar time scales. Acquiring appropriate in vivo data for any such biological process is at once a challenge and an opportunity. A model organism easily accessible for in vivo cell biological studies is the nematode Caenorhabditis elegans. Owing to its transparency, short life cycle, simple neuronal circuit and well-characterized genetics, this is a useful model organism for studying many cellular processes including axonal transport in an intact animal.
Using C. elegans as a model, we wished to develop a simple microfluidic device to acquire in vivo data of fast cell biological events across all developmental stages. This requires immobilization of the animals at any developmental stage. Current techniques use glue or anesthesia. Both methods have their advantages and limitations: (i) Cyanoacrylate glue can completely immobilize the worm but is toxic and laborious as each animal needs to be individually glued and it is difficult to implement for very young, small animals (21,22). (ii) Anesthetics are convenient and can be used to assay multiple organisms in parallel experiments but anecdotal evidence suggests that they have effects on subcellular processes. Commonly used anesthetics include sodium azide (23–25), levamisole (14–16,26), tricaine (27,28), 1-phenoxy-2-propanol (1P2P) (29) and muscimol (30,31). Many of these anesthetics act on neuronal receptors and consequently they can influence synaptic transmission (32) and possibly other processes in neurons. We found that these anesthetics have significant effects on flux of pre-synaptic vesicles, the parameter that usually correlates best with observed steady-state cargo distributions. These effects are strongest during the early stages of development.
One means to bypass the problems of anesthetics or glue is the use of microfluidic devices (33). Miniaturized microfluidic devices have been used with intact C. elegans to study many neuronal processes like olfactory behavior (19), neuronal axotomy (34–37), synaptogenesis (38) and high-resolution imaging of green fluorescent protein (GFP) expressed in cells and neurons (24,39–41). However, none of the above studies imaged subcellular processes requiring accurate spatial resolution, which depends upon effective animal immobilization. Immobilization has been accomplished in microfluidic devices using a variety of techniques such as suction channels (35,39), tapered devices (19,38,42), thin membrane layers (24,36), a membrane layer with additional cooling (39), temperature-sensitive gel-based immobilization (43) and application of anesthetic gases (40). Hitherto such devices have been mostly used to image relatively large and stationary fluorescent objects over short time scales, but not to image fast events that require highly accurate spatial resolution.
Here, we illustrate the utility of microfluidic technology to collect biologically accurate high-resolution data for various cellular and subcellular processes occurring at different time scales. We show this by using very simple and easily accessible devices to image axonal (44,45), intraflagellar (14,15,46) and dendritic transport (17) and Q neuroblast migration (47,48) in C. elegans.
We have developed a modified membrane-microfluidic device to acquire subcellular millisecond time scale data by imaging GFP-tagged proteins known to be present on various cargoes. We use the same device with larger dimensions to immobilize and image axonal transport in intact young Drosophila larvae. Moreover, in C. elegans, axonal transport data acquired in this microfluidic device appear to be spared from anesthetic-induced changes in transport. Our simple design can be used for C. elegans at all developmental stages with small modifications. Further, we are able to dynamically image for the first time pre-synaptic vesicle transport during early larval development, hitherto not possible using anesthetic preparations. Using our device, we determined that transport of pre-synaptic vesicles increases as synaptic growth takes place.
The principles behind our device can be applied to study other transparent or translucent model organisms. Our device can be readily adapted to such animals simply by changing device dimensions.
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Additional Supporting Information may be found in the online version of this article:
Figure S1: Effect of frame rate and two gaseous anesthetics on transport characteristics of GFP::RAB-3-marked vesicles. Average values of velocity (A), flux (B), pause frequency (C) and pause time (D) are plotted. Different frame rates for 3, 5, 7 and 9 fps were represented using different bar patterns for both anterograde and retrograde movements. E–H) Show representative kymographs acquired at 3, 5, 7 and 9 fps. The figure also shows the effect of compressed nitrogen and carbon dioxide gas when used in a PDMS device for immobilization. Average velocity (I), flux (J), pause frequency (K) and pause time (L) are plotted for anterograde and retrograde movements. Carbon dioxide gas showed a strong effect on transport. 14 psi CO2 application shows greater variation in transport compared to nitrogen immobilization. Kymographs show both enhanced (N) and reduced (O) transport compared to more uniform flux observed using nitrogen gas (M). Data represented as mean +/− SEM. Statistical significances are denoted by asterisk for p-value < 0.05 (*) and p-value < 0.005 (**).
Figure S2: Kymographs and pause parameters of different immobilization preparations. All images were acquired at 5 fps for 1-day adult and kymographs are plotted with the cell body on the right. Animals were immobilized using 10 mM levamisole (A), 10 mM tricaine (B), 0.5% 1P2P (C) and 6 mM muscimol (D), combination of 10 mM levamisole in a microfluidic device (E), combination of 10 mM tricaine in a microfluidic device (F) and PDMS membrane immobilization alone (G). The white triangle indicates stalled particles in animals immobilized with 10 mM tricaine (B) and PDMS membrane immobilization (G). H) Flux of anterograde and retrograde GFP::RAB-3-marked vesicles and ODR-10::GFP in anesthetized C. elegans acquired at different time-points after immobilization using 10 mM levamisole. Pause frequency and pause time of anterogradely (I and K) and retrogradely (J and L) moving particles immobilized using different anesthetics. Data represented as mean +/− SEM. Statistical significances are denoted by asterisk for p-value < 0.05 (*) and p-value < 0.005(**).
Figure S3: Pause parameters improve with development of C. elegans. Average anterograde and retrograde pause frequency (A and B) and pause time (C and D) decrease through development. Data represented as mean +/− SEM. Statistical significances are denoted by asterisk for p-value < 0.05 (*) and p-value < 0.005(**).
Movie S1: Trapping, imaging of GFP::RAB-3-marked vesicles and release of a 1-day adult in a PDMS membrane-immobilization device. Movie acquired at 5 fps and played back at 15 fps. The movies are oriented such that the anterior of the animal is to the left.
Movie S2: Time-lapse imaging of an L3 animal expressing OSM-6::GFP in a PDMS membrane-immobilization device. Movie acquired at 5 fps and played back at 15 fps. The movies are oriented such that the anterior of the animal is to the left.
Movie S3: Time-lapse imaging of an L3 animal expressing ODR-10::GFP in a PDMS membrane-immobilization device. Movie acquired at 5 fps and played back at 15 fps. The movies are oriented such that the anterior of the animal is to the left.
Movie S4: Time-lapse imaging of an L1 animal in a microfluidic device showing division of the QL expressing soluble GFP into QL.a and QL.p. This occurs approximately 4 h after hatching. The maximum distance between the cell centers was 4.5 µm in frame 5 of the movie. After this frame, the QL.a begins to migrate toward QL.p. Images were acquired every 10 min and played back at 2 fps. Cells are marked with an *. The movies are oriented such that the anterior of the animal is to the left and the right side of the animal is at the bottom.
Movie S5: Time-lapse imaging of an L1 animal in a microfluidic device showing migration of QL.a across QL.p. Images were acquired every 5–15 min and played backed at 2 fps. Cells are marked with an *. The movies are oriented such that the anterior of the animal is to the left and the right side of the animal is at the bottom.
Movie S6: Time-lapse imaging of an L1 animal in a microfluidic device showing the division of QL.a into QL.aa and QL.ap (first division in the movie) and QL.p into QL.pa and QL.pp (second division in the movie). Both divisions are completed by 2 h after the birth of QL.a and QL.p and are completed by 6 h after hatching. Images were acquired every 4–5 min and played back at 2 fps. Cells are marked with an *. The movies are oriented such that the anterior of the animal is to the left and the right side of the animal is at the bottom.
Movie S7: Time-lapse imaging of first instar Drosophila larvae expressing GFP targeted to mitochondria under a PDMS membrane-immobilization device. Movie acquired at 2 fps and played back at 15 fps. The movies are oriented such that the anterior of the animal is to the left.
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