Several people contributed to the work described in this paper. CVG built the experimental setup, and LS, SHC, and CVG performed the experiments. CVG and SHC wrote the manuscript. All authors subsequently took part in the revision process and approved the final copy of the manuscript. We thank Paul Sternberg for the GCaMP strain. Some nematode strains used in this work were provided by the Caenorhabditis Genetics Center (CGC), which is funded by the NIH National Center for Research Resources (NCRR). The MATLAB image analysis program was adapted from that used in 18. The authors were supported by Boston University and The Massachusetts Life Sciences Center.

1. Optical Setup
 <ol>
 <li>Use a standard compound microscope with epifluorescence imaging capabilities. We use a Nikon Eclipse Ti-U inverted microscope with an Intensilight HG Illuminator.</li>
 <li>For best image and signal quality use a high magnification, high numerical aperture objective. We typically use a Nikon X60 1.4 N.A. oil immersion objective. In some cases it is possible to use lower magnification (X40, X20) depending on expression level of the fluorophore and signal strength.</li>
 <li>Use a high sensitivity cooled CCD camera, such as an Andor Clara Camera to maximize image sensitivity and minimize background noise.</li>
 <li>For single color fluorophores use the standard microscope fluorescence filter set designed for the particular wavelength of the fluorophore. For GCaMP use a standard GFP filter set optimized for excitation at 488 nm and emission at 525 nm. No further optical modifications are required.</li>
 <li>For ratiometric fluorophores, such as cameleon, use a dual imaging system such that identical images can be simultaneously captured at two separate wavelengths. This is accomplished by projecting the dual images side-by-side onto the camera. A simple setup to accomplish this, using basic optical components on an optical grade breadboard, is illustrated in Figure 1. It consists of: 
 <ol class="ualpha">
 <li>A microscope filter set with 440 &plusmn;20 nm excitation filter and 455 nm long pass dichroic mirror. </li>
 <li>An image mask placed at the initial image plane immediately outside the microscope imaging port (where the camera CCD sensor would usually sit). This restricts the field of view such that it will fill only half of the camera sensor. Placing this mask at the first image plane results in sharp image borders and no overlap when the two images are projected side-by-side.</li>
 <li>The first relay lens set one focal length away from the initial image plane such that it collimates the light.</li>
 <li>A dichroic mirror that splits the light into the two separate imaging wavelengths or channels reflecting one color (&gt;515 nm, yellow) while transmitting the other (&lt;515 nm, cyan).</li>
 <li>Emission filters (band-pass filters) further restricting the transmitted light to the specific imaging wavelengths (cyan at 485 &plusmn; 40 nm and yellow at 535 &plusmn; 30 nm),</li>
 <li>The second relay lenses (one for each channel) placed in the beam paths such that it refocuses the light to a second image plane at the CCD camera.</li>
 <li>A pair of mirrors and a second dichroic mirror directs the beam paths such that the two images recombined and are projected side-by-side onto the camera CCD sensor. Figure 4A shows an example of the resulting image.</li>
 </ol>
 </li>
 <li>Initial Optical Alignment: Initiate setup and alignment of the optics with a high contrast image using a low magnification lens and transmitted wide-field light illumination (a black sharpie drawn on a microscope slide works well). Bring the image to focus through the microscope oculars and then switched to the microscope port to be used. With high light illumination view the image on an index card at the initial focal plane (a few cm outside the microscope port). Place the first relay lens (C) in the beam path such that it is one focal distance from the initial focal plane and collimates the light while not deflecting the beam path (i.e. is directly centered in the beam path). Place the mask at the first focal plane, adjusted to restrict half the field of view and produce a sharp boarder in the image. Position the mirrors and filters (D, E, G) such that the beam path remains parallel to the table surface and runs in two parallel paths side-by-side, as shown in Figure 1B. Center the second relay lenses (F) in their respective beam paths and move them back and forth along the paths to bring the images to a focus on an index card placed were the camera CCD sensor will be. If done correctly the two images should be parfocal with the microscope oculars and other camera ports. Finally place the camera in position and fine-tune the alignment.</li>
 <li>Fine Tuning Optical Alignment: Use the dual image viewed by the CCD camera to fine tune the optical alignment. Use a high contrast image and start with lower magnification, switching to higher magnification for final alignment. Position the transmitted image (cyan channel in Figure 1B) to cover one half of the camera's field of view by moving the camera in X and Y. Position the reflected image (yellow channel in Figure 1B) to cover the other half of the field of view by adjusting the last two mirrors (G) in its beam path. Optimize focus by moving the second relay lens for each channel back and forth along the beam path. If left unaltered the optical setup should be stable and only require rare fine tune adjustment.</li>
 </ol>
 2. Sample Preparation and Data Acquisition.
 <ol>
 <li>Acquire animals expressing a calcium sensitive fluorophore in the neuron of interest from the Caenorhabditis Genetics Center (C.G.C.) or other sources. Alternatively transgenic animals can be constructed using standard C. elegans techniques. Single cell expression is not necessary (and sometimes not desirable if multiple neurons are to be investigated, see 12) as long as images and measurements from multiple cells can be readily separated. If the fluorophore transgene is not integrated into the genome, significant variation in expression can occur from animal to animal. In this case, preselecting animals with higher expression levels using a fluorescent dissecting microscope increases the signal-to-noise ratio and improves the measurements. Fluorophore expression levels need to be sufficient to be readily imaged with the camera under the acquisition parameters required by the experiment.</li>
 <li>Make the agarose pads required for the desired immobilization technique (see 2.3 below) by pressing a small drop of molten agarose between two glass slides spaced by two pieces of laboratory tape. If necessary, agarose pads can be transferred to a glass cover slip before mounting the animals. See Figure 2.</li>
 <li>Immobilize C. elegans for imaging using one of the following techniques:
 <ol class="ualpha">
 <li>Pharmacological Paralyzation: Add 0.05% Levamisole to 2% agarose pads (mixing it into the molten agarose before making the pads in 2.2). Pick 5-10 animals onto the pad and cover with a cover slip. Apply small amounts of hot wax mixture (50% paraffin wax and 50% petroleum jelly) to the corners of the cover slip to hold it in position. Allow 5-10 min for the Levamisole to take effect and the animals to become completely still.</li>
 <li>Stiff Agarose Immobilization: C. elegans can be physically immobilized using stiff 10% agarose pads and 0.05 &mu;m diameter polystyrene nanoparticles. Make 10% agarose pads (10% agarose in NGM buffer by weight) as in 2.2. Add ~3 &mu;l of bead solution (1:10 polystyrene microspheres into NGM by volume) to the top of the pad. Quickly, pick 5-10 C. elegans into the pool of bead solution on the pad and gently cover with a cover slip. Use molten wax applied to the corners of the cover slip to hold it in position. (For additional details on this technique see 7 as well as a previous JoVE article 8)</li>
 <li>Gluing: C. elegans can be glued in place using small amounts of veterinary grade adhesive applied to one side of the animal.</li>
 <li>Microfluidic Devices: Numerous microfluidic devices have been designed to physically hold C. elegans in place for imaging.</li>
 </ol>
 </li>
 <li>Place the prepared slide on the microscope and focus on the desired neuron. It is often easiest to find the C. elegans using bright field illumination and low magnification. Then switching to high magnification and fluorescence imaging to find and focus on the specific neuron. </li>
 <li>Stimulus: If a neuronal response to a specific stimulus is desired (light, electric field, gustatory and olfactory cues, temperature etc.), the stimulus apparatus must be designed into the microscope setup, such that it may be administered to the animals in the sample preparation while imaging. </li>
 <li>Acquire images using the microscope CCD camera. Exposure times and excitation light level will depend on the baseline signal and expression level of the fluorophore, with weaker signals requiring longer exposure times with concordantly less time resolution. We typically use ~300 msec exposure times.</li>
 <li>Acquire images either individually or as a time-lapse movie. Because C. elegans neurons generally display slow activation dynamics (i.e. they lack sodium based action potentials), relatively slow frame rates of ~1 sec are often sufficient to measure neuron dynamics. Acquisition rates up to ~90 frames per second have been used although these may be limited by the integration time necessary for weak fluorescence. Individual movies of a single neuron can easily extend for several minutes (~5-10 min) or longer if the immobilization preparation is stable and the frame rate relatively slow.</li>
 </ol>
 3. Data Analysis
 <ol>
 <li>There are a number of different fluorescent imaging software packages available with dynamic and ratiometric analysis capabilities including ImageJ and NIS-Elements. We use a simple custom MATLAB program that measures the average fluorescent signal over a selected region of interest.</li>
 <li>For individual images (or time-lapse movies that remain perfectly still) select a region of interest (ROI) within the imaged neuron and a separate nearby region for the background measurement. For ratiometric images select similar ROI and background regions for each of the dual images. </li>
 <li>Slight movement of the imaged neuron within a time-lapse movie requires additional analysis to automatically select the ROI in each frame, which can be accomplished using numerous object tracking schemes. Our program displays a compressed image of all images in the movie from which a rough boundary (encompassing the object of interests in all frames) is manually selected and an independent background region selected. For the first frame, an ROI is manually selected from within the boundary region. For each subsequent frame, the program selects the brightest pixels from within the boundary region to generate an ROI of the correct size (i.e. the same number of pixels as the manual first frame ROI). For images where the object of interests is sufficiently brighter then the surrounding background within the boundary region this protocol selects a reasonable ROI for each frame.</li>
 <li>The fluorescent signal, F, for each frame is calculated as the average pixel intensity in the ROI minus the average intensity in the background region. Ratiometric values are calculated as the ratio of the two signals (YFP-YFPbackground)/(CFP-CFPbackground)-0.65. The factor of 0.65 compensates for bleed through of the CFP channel into the YFP channel, which will be dependant on the fluorophores as well as the particular filter set used (this value is typically ~0.6 for cameleon setups).</li>
 <li>The relative calcium level for each frame is calculated as the percent change in intensity normalized to an initial baseline value measured from the first frame (or series of frames): &Delta;F/F=(F(t)-Fo)/ Fo, where F(t) is the fluorescence at time t and Fo is the initial baseline value.</li>
 <li>A ROI selecting the cell body generates the strongest most robust signals but it is also possible to select and measure signals from a ROI along the nerve processes.</li>
 </ol>
 4. Problem Solving
 <ol>
 <li>Bleaching: Exposure to excitation light can cause bleaching of the fluorophore and is marked by a steady decrease in signal that is dependent on exposure levels. Ratiometric measurements help to compensate for this. However selective bleaching of one wavelength can still occur (typically CFP bleaches quicker then YFP). If bleaching occurs reduce the light exposure level by reducing exposure times and/or excitation intensity (with neutral density filters in the illumination pathway). If necessary, one can compensate for bleaching by running mock trials (with the same excitation exposure but no neuronal stimulus) and quantifying the time constant of signal decay.</li>
 <li>Immobilization: For the simple image analysis described here it is critical the animal remains relatively still throughout the recording period. C. elegans respond to light exposure and experimental stimuli, making them most likely to move at the time of recording. Slight movement can introduce significant noise into the measurement especially when measuring from axon processes. Ratiometric analysis helps to mitigate these effects to some extent and measurements from the cell body are more robust. Careful execution and refinement of the immobilization procedures will result in completely still animals.</li>
 <li>Background Selection: Careful selection of a background region is necessary for successful image analysis. Background regions should be reasonably uniform and darker then the imaged neuron. It must be sufficiently removed to avoid scattered light (and thus pickup a signal) from the imaged neuron. We find a uniform region 20-50 &mu;m from the ROI works well in most cases.</li>
 <li>Ratiometric Optics: Correct alignment of the imaging optics is critical for generating a clear dual image. Commercial microscope attachments for ratiometric imaging are available. An alternative technique is to employ rapid sequential imaging using a fast filter wheel to rapidly switch between emission wavelengths while recording separate images for each.</li>
 <li>Ratiometric Analysis: The analysis described here is a relatively simple technique that generates robust signals by virtue of averaging over a ROI. It requires a recognizably bright object to select the ROI. More sophisticated ratiometric analysis is possible by precisely aligning and mapping the dual images such that the fluorescent ratio can be calculated pixel by pixel. </li>
 </ol>

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The authors declare that they have no competing financial interests.

Genetically encoded calcium indicators have been widely utilized in C. elegans neurobiology. Numerous groups have employed these techniques to study response of primary sensory neurons to external stimuli as demonstrated here with the ASJ response to an electrical field. Prominent examples include sensation of mechanical touch, specific chemicals, temperature and an electric field 12, 16-19. Activity of interneuron and muscle cells have also been monitored both in response to stimuli and in con...

Here we present practical methods for in vivo calcium imaging in C. elegans neurons. The development of genetically encoded calcium-sensitive fluorophores with high signal-to-noise ratio makes C. elegans a comparatively straightforward and cost effective system for measurement of neurophysiology and activity. Our imaging is done with a standard compound microscope using wide-field fluorescence imaging of commonly available fluorophores. We present several techniques employing various fluorophores and different sample preparations, discussing the strengths and weakness of each. Data is then presented from two example experiments. An excellent additional resource on the techniques described here can be found in WormBook, "Imaging the activity of neurons and muscles" by R. Kerr, (<a href="http://www.wormbook.org" target="_blank">http://www.wormbook.org</a>) 3.
 Two major classes of genetically encoded fluorescent calcium reporters are commonly used in C. elegans: single channel GCaMP and FRET-based cameleon. We will describe methods and show examples for data generated by each. 
 GCaMP is based on a modified Green Fluorescent Protein (GFP) that is sensitive to the surrounding calcium concentration. This is accomplished by fusion of GFP and the high calcium affinity protein calmodulin, such that the binding of calcium by calmodulin brings the GFP molecule into an efficient fluorescent confirmation 2. The recent advancements in these fluorophores generate exceptional signal size with up to 500% increase in fluorescence intensity over a physiological range of calcium levels and reasonably fast kinetics of ~95 msec rise time and ~650 msec decay time 4. Over relatively short time periods (mins), these large signals can allow for lower resolution imaging (lower magnification) and, given a well-behaved initial baseline measurement, negate the necessity for continuous baseline or comparative measurements.
 Cameleon has the advantage of being a FRET-based fluorophore that generates a ratiometric measurement comparing two independent channels or wavelengths 1. It consists of two separate fluorophores (cyan- and yellow-emitting fluorescent proteins, CFP and YFP) linked by a calmodulin protein. The complex is illuminated with blue light (440 nm) that excites the CFP. Binding of calcium brings the fluorophores closer together, increasing fluorescence resonance energy transfer (FRET) from the CFP (donor) to the YFP (acceptor) and causing the CFP emission (480 nm) to decrease and the YFP emission (535 nm) to increase. Relative calcium levels are measured as the ratio of the YFP/CFP intensity. Cameleon kinetics are slower than that of GCaMP, measured in vivo to have a rise time of ~1 sec and a decay time of ~3 sec 5. However, the ratio of oppositely moving signals increases the signal size and compensates for a number of possible artifacts due to changes in fluorophore concentration, motion or focus drift and bleaching. 
 Genetically encoded fluorescent reporters negate much of the sample preparation required with exogenously administered probes and C. elegans small transparent body allows imaging within the intact animal using simple wide field fluorescence. The main technical challenge in sample preparation is therefore to safely immobilize the animals. There are a number of different commonly used techniques each with advantages and disadvantages. Using a pharmacological agent to paralyze the animals is easy to implement and allows the mounting of multiple animals on one preparation (Levamisole, a cholinergic agonist that causes muscle tissue to seize is typically used 6). C. elegans can also be physically immobilized by mounting them on stiff 10% agarose 7, 8. This minimizes impact on animal physiology, allows long-term imaging (hours) and recovery of multiple animals but is more technically difficult. Both of these techniques restrict physical access to the animals (which are under a cover slip) and can therefore only be used with certain experimental stimuli (such as light, temperature, electric field or laser damage). For stimuli where physical access is required, such as touch or administration of chemicals, many studies have successfully glued C. elegans in place (using veterinary grade glue) 9. This is technically more challenging, is a single animal preparation and does not allow animal recovery. Finally, numerous microfluidic devices have been employed that physically restrain C. elegans, preserving animal physiology, allowing exposure to most types of stimuli (depending on the device design) and can enable rapid exchange and recovery of the animals 10, 11. However microfluidics require additional technical skills and capabilities in design, fabrication and implementation. In immobilized animals activity and stimulus response can generally be measured in sensory and interneurons. Activity of motor neurons requires more sophisticated techniques for imaging in moving animals. Here we will present detailed methods employing the two most straightforward techniques of pharmacological paralyzation and immobilization with stiff agarose.
 The methods presented here can be used to measure neuronal activity and cell physiology in C. elegans. We give an example of each: using GCaMP to measure the sensory response of the ASJ neuron to an external electric field, and using cameleon to measure the physiological calcium response to laser damage of a neuron. These examples show the benefits and drawbacks of the two types of fluorophores and illustrate what is possible with the system.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Eclipse Ti-U inverted 
 microscope</td>
 <td>Nikon</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Intensilight HG Illuminator</td>
 <td>Nikon</td>
 <td>C-HGFI</td>
 <td>Fluorescent light source</td>
 </tr>
 <tr>
 <td>CFI Plan Apo VC 60X Oil</td>
 <td>Nikon</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Optical table or 3'X3' optical 
 grade breadboard</td>
 <td>Thor Labs</td>
 <td>&nbsp;</td>
 <td>If an optical table is not used an 
 optical grade breadboard on a 
 solid laboratory bench should 
 suffice.</td>
 </tr>
 <tr>
 <td>Clara Interline Camera</td>
 <td>Andor 
 Technology</td>
 <td>&nbsp;</td>
 <td>High-sensitivity CCD camera</td>
 </tr>
 <tr>
 <td>wtGFP Longpass Emission</td>
 <td>Chroma Technology 
 Corp.</td>
 <td>41015</td>
 <td>GFP filter set for imaging GCaMP</td>
 </tr>
 <tr>
 <td>Filter 440 +/- 10 nm</td>
 <td>Chroma</td>
 <td>D440/20x EX</td>
 <td>excitation filter for cameleon</td>
 </tr>
 <tr>
 <td>Dichroic mirror &gt; 455 nm 
 longpass</td>
 <td>Chroma</td>
 <td>455DCLP BS</td>
 <td>microscope dichroic for cameleon 
 imaging</td>
 </tr>
 <tr>
 <td>Dichroic mirror &gt; 515 nm 
 longpass</td>
 <td>Chroma</td>
 <td>515DCLP BS</td>
 <td>dichroic mirror for cameleon 
 imaging</td>
 </tr>
 <tr>
 <td>Filter 535 +/- 15 nm</td>
 <td>Chroma</td>
 <td>D535/30m EM</td>
 <td>YFP emission filter</td>
 </tr>
 <tr>
 <td>Filter 480 +/- 20 nm</td>
 <td>Chroma</td>
 <td>D485/40m EM</td>
 <td>CFP emission filter</td>
 </tr>
 <tr>
 <td>Lens, 200 mm, Achromat</td>
 <td>Thor Labs</td>
 <td>AC508-200-A1</td>
 <td>Relay lens for FRET optics (3)</td>
 </tr>
 <tr>
 <td>Silver broadband mirror</td>
 <td>Thor Labs</td>
 <td>ME2S-P01</td>
 <td>FRET optics (2)</td>
 </tr>
 <tr>
 <td>NGM buffer</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Levamisole</td>
 <td>Sigma</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Polybead Microspheres</td>
 <td>Polysciences, Inc.</td>
 <td>08691-10, 2.5% by 
 volume, 50 nm 
 diameter</td>
 <td>polystyrene nanoparticles for C. 
 elegans immobilization</td>
 </tr>
 <tr>
 <td>Transgenic strain, Strain 
 gpa-9::GCaMP3(in pha-1; 
 him-5 bkg) </td>
 <td>Sternberg Lab</td>
 <td>Strain PS6388</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Transgenic strain, mec- 
 4::YC3.60</td>
 <td>Gabel Lab</td>
 <td>Strain CG1B</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

in vivo neuronal calcium imaging in c. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

Here we present results from two separate experiments. The first employs GCaMP to measure the response of a specific sensory neuron to a defined external stimulus, giving a good example of how fluorescent calcium reporters can be used to optically monitor neuronal activity in intact C. elegans. The second employs cameleon to measure the intracellular calcium transient triggered within a neuron in response to specific laser damage, thus illustrating how calcium physiology can be measured within a single cell ...

Watch this Scientific Journal Video about In vivo Neuronal Calcium Imaging in C. elegans at JoVE.com

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

in-vivo-neuronal-calcium-imaging-in-c-elegans

Research

JoVE Journal

Neuroscience

24.8K Views.  Boston University School of Medicine. With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

Video: In vivo Neuronal Calcium Imaging in C. elegans

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new method of Ca2+-imaging using the bioluminescent reporter GFP-aequorin (GA) has been developed. This new technique relies on the fusion of the GFP and aequorin genes, producing a molecule capable of binding calcium and&#160;&#8212; with the addition of its cofactor coelenterazine &#8212; emitting bright light that can be monitored through a photon collector. Transgenic lines carrying the GFP-aequorin gene have been generated for both mice and Drosophila. In Drosophila, the GFP-aequorin gene has been placed under the control of the GAL4/UAS binary expression system allowing for targeted expression and imaging within the brain. This method has subsequently been shown to be capable of detecting both inward Ca2+-transients and Ca2+-released from inner stores. Most importantly it allows for a greater duration in continuous recording, imaging at greater depths within the brain, and recording at high temporal resolutions (up to 8.3 msec). Here we present the basic method for using bioluminescent imaging to record and analyze Ca2+-activity within the mushroom bodies, a structure central to learning and memory in the fly brain.

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Here we present a novel Ca2+-imaging approach using a bioluminescent reporter. This approach uses a fused construct GFP-aequorin which binds to Ca2+ and emits light, eliminating the need for light excitation. Significantly this method permits long continuous imaging, access to deep brain structures and high temporal resolution.

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new ...

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its completely mapped nervous system and transparent anatomy, presents an ideal model for understanding real-time neural dynamics using calcium indicators. In combination with microfluidic technologies and experimental designs, calcium-imaging studies using these indicators are performed in both free-moving and trapped animals. However, most previous studies utilizing trapping devices, such as the olfactory chip described in Chronis et al., have devices designed for use in the more common hermaphrodite, as the less common male is both morphologically and structurally dissimilar. An adapted olfactory chip was designed and fabricated for increased efficiency in male neuronal imaging with using young adult animals. A turn was incorporated into the worm loading port to rotate the animals and to allow for the separation of the individual neurons within a bilateral pair in 2D imaging. Worms are exposed to a controlled flow of odorant within the microfluidic device, as described in previous hermaphrodite studies. Calcium transients are then analyzed using the open-source software ImageJ. The procedure described herein should allow for an increased amount of male-based C. elegans calcium imaging studies, deepening our understanding of the mechanisms of sex-specific neuronal signaling.

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of an adapted &#34;olfactory chip&#34; for the efficient calcium imaging of C. elegans males is described here. Studies of male exposure to glycerol and a pheromone are also shown.

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

Toxicity of misfolded proteins and mitochondrial dysfunction are pivotal factors that promote age-associated functional neuronal decline and neurodegenerative disease across species. Although these neurotoxic challenges have long been considered to be cell-intrinsic, considerable evidence now supports that misfolded human disease proteins originating in one neuron can appear in neighboring cells, a phenomenon proposed to promote pathology spread in human neurodegenerative disease.
C. elegans adult neurons that express aggregating proteins can extrude large (~4 &#181;m) membrane-surrounded vesicles that can include the aggregated protein, mitochondria, and lysosomes. These large vesicles are called &#8220;exophers&#8221; and are distinct from exosomes (which are about 100x smaller and have different biogenesis). Throwing out cellular debris in exophers may occur by a conserved mechanism that constitutes a fundamental, but formerly unrecognized, branch of neuronal proteostasis and mitochondrial quality control, relevant to processes by which aggregates spread in human neurodegenerative diseases.
While exophers have been mostly studied in animals that express high copy transgenic mCherry within touch neurons, these protocols are equally useful in the study of exophergenesis using fluorescently tagged organelles or other proteins of interest in various classes of neurons.
Described here are the physical features of C. elegans exophers, strategies for their detection, identification criteria, optimal timing for quantitation, and animal growth protocols that control for stresses that can modulate exopher production levels. Together, details of protocols outlined here should serve to establish a standard for quantitative analysis of exophers across laboratories. This document seeks to serve as a resource in the field for laboratories seeking to elaborate molecular mechanisms by which exophers are produced and by which exophers are reacted to by neighboring and distant cells.

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

This protocol describes approaches for detection and quantitation of large aggregate and/or organelle extrusions (~4 &#181;m) produced by C. elegans cells in the form of membrane-bound exophers. We describe strains, growth conditions, scoring criteria, timing, and microscopy considerations needed to facilitate dissection of this debris expulsion mechanism.