Work in the Hammarlund lab is funded by: NIH grants R01 NS066082-01 and T32GM007223, the Beckman Foundation, and the Ellison Medical Foundation.

1. Assembling the system
The specific components of our system are outlined below. When properly assembled, the user should be able to focus the microscope with one hand, move the stage with the other (either with the mouse or the joystick), activate the laser with the foot pedal, and have a clear view of the on-screen image.
<ol>
	<li>MicroPoint laser mounted to the epi-fluorescence port of a compound microscope. We use a Nikon 80i, but any research-grade upright or inverted scope should work. The MicroPoint system will include a custom dichroic, which should match the fluorophore that labels the cells you wish to cut. Cutting cells labeled with alternative fluorophores requires additional dichroics. Additionally, a pulse generator and foot pedal come with the MicroPoint system.</li>
	<li>100x, 1.4 NA oil objective for performing surgery and assessing regeneration, plus additional objectives as necessary for locating the sample on the slide. We use a Nikon Plan Apo VC for surgery, and find that a 4x is useful for coarse focusing and finding the sample.</li>
	<li>Motorized stage with joystick. A stage with extremely high accuracy and repeatability is unnecessary, since the stage is only used for targeting. We use a Prior OptiScan II (see discussion).</li> 
	<li>Camera. The camera needs to provide a frame rate sufficient to focus and target the labeled neurite with the 100x objective, while visualizing the image on the computer monitor. We prefer attenuating the illumination light during the axotomy procedure to avoid potential confounding effects from photo-bleaching or damage (see 1.7, below). Thus, the camera needs to be sensitive enough to accommodate this reduced light level. We use a Hamamatsu Orca 05G, and find that imaging at 16.3 Hz is sufficient. Equivalent cameras could be substituted.</li>
	<li>Computer, monitor, and imaging software that can drive the camera. Additionally, we use the imaging software to manipulate the motorized stage with a mouse (see discussion).</li>
	<li>Air table. This requirement may vary depending on location, but since surgery is performed with a 100x objective, we find vibration isolation to be essential.</li>
	<li>Light supply with light guide and shutter mechanism for illuminating the fluorophore. The light guide will attach to the MicroPoint. The shutter mechanism is useful to reduce the illumination intensity independently of the laser. We use a Prior Lumen 200 or a Leica EL6000, and attenuate up to 80% of total light intensity for axotomy.</li>
</ol>
2. Laser setup
A detailed protocol to initially align and focus the laser is provided with the MicroPoint laser system. We assume that procedure has been followed successfully upon initial setup. This includes aligning the laser with the crosshairs in the eyepiece. Here we provide a protocol for regular maintenance of the laser's focus and intensity.
As noted in the MicroPoint manual, the pulsed nitrogen laser is capable of severely damaging the human eye. Care should be taken to never look directly into the laser beam. Once installed, the barrier filter should safely block radiation through the eyepieces.
Cutting neurons in an organism the size of a grain of sand can be challenging. Since the area of damage done by the laser is very small, it is essential to know the three-dimensional location of the laser's focus in order to target the sample precisely. The laser focus is found using the mirrored slide. First, we check that the z location of the laser focus matches the focus of the imaging path. Next, the x-y location of the damage focus is marked with crosshairs in the imaging software.
Focusing in the z-plane
<ol>
	<li>Push in the neutral density filter ND16 (Figure 1a). We use neutral density filters in the epi-illumination path to attenuate the laser while focusing. For even finer focusing, push in both the ND16 and ND4 fluorescent filters. </li>
	<li>Set the MicroPoint laser to 1 pulse with the switch turned to 'Select' (Figure 1b), and then move the filter wheel to the appropriate long-pass barrier filter (Figure 1c). </li>
	<li>Place the mirrored slide on the stage and focus on pinholes within it with the 4X objective using transmitted light.</li>
	<li>Add a drop of immersion oil to the mirrored slide and refocus on pinholes with the 100X objective. Open the camera shutter with the optical path switching lever (Figure 1d) and the laser shutter (Figure 1e). If necessary, reduce transmitted light intensity so the edges of the pinholes are crisp: this will aid in focusing exactly.</li>
	<li>Press the foot pedal. This should produce a small hole in the mirror if the laser is focused properly.</li>
	<li>Focus the microscope slightly above the plane of the mirrored slide and shoot the laser again. This should make an even smaller hole than the first shot, or leave no mark at all. Repeat by focusing below the mirrored slide.</li>
	<li>If a hole is not produced in 2.5, or if a larger hole is produced when focusing above or below the surface of the mirror, refocus the laser with the focus ring on the ablation head. Verify that the hole is still properly aligned with the crosshairs in the eyepiece. With consistent use, the z-plane focus rarely needs to be adjusted.</li>
</ol>
Focusing in the x-y plane 
<ol start="8">
	<li>Follow steps 2.1 through 2.3 above and press the foot pedal to make a small hole in the mirror slide.</li>
	<li>Initiate 'Live' capture mode from the Elements 'Acquire' menu. In the 'Measure' toolbar select 'ROI editor', then select the 'double cross' tool and drag the crosshairs to align them with the center of the new hole. Click 'Save ROI' and then 'Exit editor' and resume Live capture mode.</li>
	<li>Activate the 'mouse XY' setting in Elements to manipulate the stage with the mouse.</li>
	<li>Pull the neutral density filter(s) back to the original position(s), set the laser pulses to 10, and remove the mirrored slide.</li>
</ol>
Adjusting the power of the laser
<ol start="12">
	<li>The power of the laser should be adjusted to the minimum power necessary to cut the neuron of your choice. Adjust the power by moving the attenuator plate (Figure 1f) while cutting living worms (described below). If the laser power is too high, it will cause peripheral damage or blast a hole in the worm. If the laser power is too low, it will not sever the neuron. Once the system has been set up, the attenuator position rarely needs to be changed. If the laser becomes weak, and the attenuator plate needs to be moved to continue cutting, it could be a sign that the dye in the dye cell needs to be replaced (see discussion).</li>
</ol>
3. Immobilize the worms
<ol>
	<li>Prepare a solution of 3% molten agarose in M9 (22mM KH2PO4, 42mM Na2HPO4, 85mM NaCl, 1mM MgSO4). Dispense 5 mL of molten agarose into a 15 mL falcon tube. Fill another tube with water. Place tubes in a modular heating element set to 55&deg;C. Fill the heating element with water to create mini water baths for each tube. Note: agarose can be prepared in large quantities in advance and re-melted for subsequent experiments. </li>
	<li>Place two layers of labeling tape on each of two slides and a clean slide between these slides. Dispense a drop of agarose using a plastic bulb pipette to the middle of the clean slide and place another slide perpendicular to the first so that the resulting agarose pad is the thickness of two pieces of tape (Figure 2). Remove one of the two slides from the pad after 30 seconds. Rinse and store the plastic pipette in the water-filled falcon tube for use with subsequent slides.</li>
	<li>Next, place 3-5 &mu;L of 0.10 &mu;m or 0.05 &mu;m polystyrene beads in the center of the pad. Add 5-10 worms expressing fluorescent protein in the neurons of interest. If the neurons of interest are asymmetrically distributed on the right or left side of the worm, use a platinum pick to flip worms so that the correct side is facing up. Worms must be placed on slides within 10 minutes of preparing the agarose pads.</li>
	<li>Carefully place a cover slip on the worms. After placing the cover slip, do not move it relative to the agarose pad as this will disrupt the cuticle and destroy the worms. Place the prepared slide on the microscope stage.</li>
</ol>
4. Cut neurons
<ol>
	<li>Direct light into the eyepieces with the optical path switching lever. Focus on the worm to be cut with the 4X objective, place a drop of oil on the cover slip, and switch to the 100X objective. Return the optical path to the camera.</li>
	<li>Use the mouse to move the neuron of your choice under the center of the crosshairs. Press the foot pedal to fire the laser and sever the neuronal process. If needed, two sets of pulses can be used to sever the neuron. At first, it is helpful to track precisely which neurons have been severed in individual worms in order to assess subsequent regeneration. When performed correctly, laser axotomy produces a small break in the neuron without causing significant damage to the surrounding skin tissue or other neurons (see Figure 3). In certain cases, a small scar may form around the site of injury.</li>
	<li>Recover the worms by removing the cover slip in a direct upward movement, being careful that the worms remain on the agarose pad. Do NOT slide the coverslip off. Then, cut the pad around the worms with needle-nose forceps and place the section of agarose on a freshly seeded NGM plate containing OP508. Place 10 &mu;L of sterile M9 on the pad to free the worms from the beads and remove agarose.</li>
</ol>
5. Score neurons for regeneration
<ol>
	<li>Prepare agarose pad as outlined in step 2.2.</li>
	<li>Place worms in 3-5 &mu;L of polystyrene beads. Apply a coverslip. Slides can be prepared sequentially, or alternatively all the worms can be prepared in advance and the slides kept in 10 cm culture dishes, each containing a damp Kimwipe to keep the agarose moist. </li>
	<li>Use the 100X objective to visualize the severed neurons. In many neurons, a neuronal stump distal to the cut site will remain. We use the presence of this remnant as an indication that the neuron was indeed severed (Figure 4). The presence of this stump makes it possible to distinguish a neuron that has been cut and regenerated from one that was not cut. Note: both the distal and proximal stumps initially retract after being cut.</li>
	<li>Assess regeneration. A variety of parameters can be determined. One simple assay is whether the injured neuron has formed a growth cone (Figure 4a) and regenerated, or not (Figure 4b). Alternatively, neurite length can be traced and compared.</li>
</ol>
6. Representative results
As an example, we describe characterization of regeneration of the &gamma;-Aminobutyric acid (GABA) motor neurons. These neurons, which reside in the ventral nerve cord and extend processes circumferentially to the dorsal nerve cord, are essential for proper locomotion9. GABA neurons can be visualized by expressing a genetically encoded fluorophore, such as green fluorescent protein (GFP), under the control of the unc-47 or unc-25 promoters (strains EG1285 and CZ1200, respectively, available from the C. elegans Genetic Center).
Mount L4-stage worms as described, flip the worms so the GABA neurons on their right sides are facing up, and place the cover slip. Cut 1-3 of the posterior commissures in each worm, midway between the dorsal and ventral cords. Avoid cutting commissures that cross or fasciculate with other commissures, as these are difficult to score later. Recover worms as described.
18 to 24 hours after a successful axotomy, the worms will have recovered from surgery and exhibit normal locomotory and egg-laying behaviors. Discard worms that are dead or sick. Remount the remainder as described, and assess regeneration. We usually aim to assess at least 30 cut neurons per experimental condition. In oxIs12 animals, typically 60-70% of severed L4 GABA neurons will have initiated a growth cone structure, evidenced by a broadening of membrane at the tip and the extension of several neurite branches, and migrated from the cut site (Figure 4a). In many cases, the growth cones will have migrated to reconnect with the dorsal nerve cord, an adjacent neuronal commissure, or the distal stump. The remaining 30% of neurons will have either initiated no growth, forming a stump proximal to the site of axotomy, or will have extended only small filopodia (Figure 4b). 
<img src="/files/ftp_upload/2707/2707fig1.jpg" alt="Figure 1"/> 
Figure 1. The UV pulsed laser axotomy system. Specific components are: (a) ND filters (b) pulse selector (c) filter wheel lever (d) optical path switching lever (e) laser shutter and (f) attenuator plate.
<img src="/files/ftp_upload/2707/2707fig2.jpg" alt="Figure 2"/> 
Figure 2. Preparing an agarose pad. To prepare an agarose pad of the desired thickness, two layers of tape (green) are placed on two slides. A slide is placed between the first two, and a drop of agarose is then added to the clean slide. Finally, a fourth slide is placed perpendicularly to the first three, resulting in a pad that is the thickness of two pieces of tape.
<img src="/files/ftp_upload/2707/2707fig3.jpg" alt="Figure 3"/> 
Figure 3. Representative GABA neuron axotomies. In a typical experiment, GABA neuron commissures are severed at the middle of the lateral aspect of the worm. A severed commissure is shown immediately before (left panel) and after (right panel) axotomy. All images were taken with a 100X oil objective.
<img src="/files/ftp_upload/2707/2707fig4.jpg" alt="Figure 4"/> 
Figure 4. Representative results of GABA neuron axotomies. 24 hr post-axotomy severed axons are scored for regeneration. A regenerating axon with a growth cone (arrow) is shown in (a) while a non-regenerating axon is seen as a proximal stump (arrow) in (b). The distal stump of each cut axon is shown in each panel (arrowhead) and is an indication that the axon has been severed.

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No conflicts of interest declared.

A variety of laser systems have been used to cut neurites in C. elegans, and several studies have examined their performance in detail3,7,10,11. The MicroPoint laser used in our protocol is a turnkey system that is easy to set up and maintain, and is available at a low cost to researchers compared to a Ti-Sapphire laser system. Compared to a Ti-Sapphire system, however, the MicroPoint laser is expected to cause damage to a larger area, which may be disadvantageous for some applications. If a Ti-Sapphire system is desired, an excellent protocol on building such a system is available12. 
The current protocol can be performed on a variety of neurons in C. elegans; however, we note that differences in regenerative abilities between distinct types of neurons are expected13. Additionally, different transgenic backgrounds can affect regenerative success. Although the percentage of regenerating GABA neurons is fairly consistent between different transgenic marker strains, we have noted an overall slight increase in regeneration in juIs7614 vs. oxIs12 15 worms. Differences between markers of the touch neurons have also been described2.
We prefer to immobilize the worms with microbeads rather than anesthetics as the beads result in faster and more consistent immobilization16. This is also advantageous as we are able to observe regeneration free of any possible confounding effects due to anesthesia17. An alternative anesthetic-free method for immobilization is the use of microfluidic devices. The use of microfluidics for axotomy has been extensively described17-22.
We find that with consistent use, the laser functions best when the Coumarin 440 in the dye cell is changed once each week following the procedure outlined in the MicroPoint manual. If necessary, the attenuation slider can be used to increase power, but this may be an indication of old dye or laser misalignment. Also, the dye cell has a limited lifetime and will eventually need to be rebuilt or replaced (See Troubleshooting).
It can be difficult to maneuver the target neurite under the crosshairs that mark the laser focus, particularly if the animal is incompletely paralyzed. We find that a manual stage is not optimal for this purpose, although it is certainly usable. A joystick-controlled motorized stage is more precise, and we find that using software that supports image dragging with the mouse to move the motorized stage is best. Nikon Elements provides this feature and is described in this protocol, but the free Micromanager package, as well as other imaging software, may have similar functionality. A different way of fine targeting is to move the laser focus, rather than the animal. A galvanometer beam-steering mechanism is available as an add-on to the MicroPoint laser, if this approach is preferred. 
Besides its application to the study of neuronal regeneration, the laser can be used to ablate other cell types, such as skin, muscle, or specialized cells, or to disrupt specific neuronal synapses23-27. Moreover, pathways that regulate the degeneration of neurons that accompanies injury or disease could be investigated with this system. As such, the use of pulsed lasers will continue to shed light on both the genetic factors and the cell biological changes that facilitate neuronal regeneration and other relevant processes.
Troubleshooting:
Described here are some common problems and their associated solutions.
<ol type="a">
	<li>The laser is not cutting properly. This is likely because either the laser is out of focus (see section 2), or the dye in the dye cell needs to be replaced (see MicroPoint manual). Alternatively, the dye cell may need to be replaced or rebuilt.</li>
 <li>The worms are moving on agarose pad. This is generally a sign that the agarose pad is too moist. We find that pads need to be left for approximately 30 seconds before worms are placed on them to circumvent this problem. You can also try using the smaller sized (0.05 &mu;m) microbeads.</li>
 <li>The recovery of worms is difficult and inefficient. This can be caused by incorrect coverslip removal or by a dry agarose pad. If the pad is too dry, you may notice that the axons develop a beaded appearance. In some cases, this is because the worms were not placed on the pad soon enough after it was made. Alternatively, an old stock of agarose solution may have a higher than 3% concentration after repeated melting.</li>
</ol>

<table border="1">
 <tbody>
 <tr>
 <th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments </th>
 </tr>
 <tr>
 <td>0.05 &mu;m Polystyrene Beads</td>
 <td>Polysciences, Inc.</td>
 <td>08691</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>0.10 &mu;m Polystyrene Beads</td>
 <td>Polysciences, Inc.</td>
 <td>00876</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Agarose GPG/LE</td>
 <td>American Bioanalytical</td>
 <td>00972</td>
 <td>Ultra pure</td>
 </tr>
 <tr>
 <td>Falcon 14 ml Polystyrene Round-Bottom Tube</td>
 <td>BD Biosciences</td>
 <td>352057</td>
 <td>17 x 100 mm style, nonpyrogenic</td>
 </tr>
 <tr>
 <td>Thermo Scientific Plain precleaned microscope slides</td>
 <td>Erie Scientific Company</td>
 <td>420-004T</td>
 <td>3" x 1" x 1 mm</td>
 </tr>
 <tr>
 <td>Cover Slips</td>
 <td>VWR</td>
 <td>48366 205</td>
 <td>18 mm x 18 mm No. 1 1/2</td>
 </tr>
 <tr>
 <td>KIMTECH Science Kimwipes</td>
 <td>Kimberly-Clark </td>
 <td>34155</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>BD Falcon 100 x 15mm Style Petri Dish</td>
 <td>BD Biosciences</td>
 <td>351029</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tape blank 3/4w x 500L</td>
 <td>TimeMed </td>
 <td>T-512</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Immersion oil</td>
 <td>Nikon</td>
 <td>&nbsp;</td>
 <td>Type A, nd=1.515</td>
 </tr>
 <tr>
 <td>EG1285 or CZ1200</td>
 <td>C. elegans Genetic Center</td>
 <td>http://www.cbs.umn.edu/CGC/strains/</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>OptiScan II</td>
 <td>PRIOR Scientific</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>NIS-Elements Ar or Br</td>
 <td>Nikon</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>MicroPoint Ablation Laser System</td>
 <td>Photonic Scientific</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Compound microscope</td>
 <td>Nikon</td>
 <td>Eclipse 80i</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Hamamatsu Camera</td>
 <td>Hamamatsu Photonics</td>
 <td>Model C8484-05G01</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dell Precision T3400 PC with Intel Core 2 Duo</td>
 <td>Dell</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Windows XP Professional</td>
 <td>&nbsp;</td>
 <td>Microsoft</td>
 <td>Version 2002, Series Pack 3</td>
 </tr>
 <tr>
 <td>Dual dry bath Incubator</td>
 <td>Fisher Scientific</td>
 <td>&nbsp;</td>
 <td>Analog controls</td>
 </tr>
 <tr>
 <td>4X Plan Fluor objective</td>
 <td>Nikon</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>100X Plan Apo VC oil objective</td>
 <td>Nikon</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dumont #5/45 forceps</td>
 <td>Dumont</td>
 <td>11251-35</td>
 <td>Dumoxel standard tips, can also use #7</td>
 </tr>
 <tr>
 <td>Dissecting Microscope</td>
 <td>Nikon </td>
 <td>SMZ800</td>
 <td>With NI-150 High Intensity Illuminator</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about In vivo Laser Axotomy in C. elegans at JoVE.com

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.

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 ...

Research

JoVE Journal

Neuroscience

18.2K Views.  Yale University School of Medicine. 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.

Video: In vivo Laser Axotomy 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 ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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 ...

C. elegans Tracking and Behavioral Measurement

We have developed instrumentation, image processing, and data analysis techniques to quantify the locomotory behavior of C. elegans as it crawls on the surface of an agar plate. For the study of the genetic, biochemical, and neuronal basis of behavior, C. elegans is an ideal organism because it is genetically tractable, amenable to microscopy, and shows a number of complex behaviors, including taxis, learning, and social interaction1,2. Behavioral analysis based on tracking the movements of worms as they crawl on agar plates have been particularly useful in the study of sensory behavior3, locomotion4, and general mutational phenotyping5. Our system works by moving the camera and illumination system as the worms crawls on a stationary agar plate, which ensures no mechanical stimulus is transmitted to the worm. Our tracking system is easy to use and includes a semi-automatic calibration feature. A challenge of all video tracking systems is that it generates an enormous amount of data that is intrinsically high dimensional. Our image processing and data analysis programs deal with this challenge by reducing the worms shape into a set of independent components, which comprehensively reconstruct the worms behavior as a function of only 3-4 dimensions6,7. As an example of the process we show that the worm enters and exits its reversal state in a phase specific manner.

C. elegans Tracking and Behavioral Measurement

We have developed a video-rate tracking microscope system that can record and quantify C. elegans behavior at high resolution and high speeds. We have also developed computational methods to reduce the dimensionality of the worm images to a fundamental set of measurements that completely describe the shape of the worm.

We have developed instrumentation, image processing, and data analysis techniques to quantify the locomotory behavior of C. elegans as it crawls on the ...

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps shape attention and also protects cells from over-stimulation. Adaptation within the olfactory circuit of C. elegans was first described by Colbert and Bargmann1,2. Here, the authors defined parameters of the olfactory adaptation paradigm, which they used to design a genetic screen to isolate mutants defective in their ability to adapt to volatile odors sensed by the Amphid Wing cells type C (AWC) sensory neurons. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor3 for 30 min they will adapt their responsiveness to the odor and will then ignore the adapting odor in a chemotaxis behavioral assay for ~1 hr. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor for ~1 hr they will then ignore the adapting odor in a chemotaxis behavioral assay for ~3 hr. These two phases of olfactory adaptation in C. elegans were described as short-term olfactory adaptation (induced after 30 min odor exposure), and long-term olfactory adaptation (induced after 60 min odor exposure). Later work from L'Etoile et al.,4 uncovered a Protein Kinase G (PKG) called EGL-4 that is required for both the short-term and long-term olfactory adaptation in AWC neurons. The EGL-4 protein contains a nuclear localization sequence that is necessary for long-term olfactory adaptation responses but dispensable for short-term olfactory adaptation responses in the AWC4. By tagging EGL-4 with a green fluorescent protein, it was possible to visualize the localization of EGL-4 in the AWC during prolonged odor exposure. Using this fully functional GFP-tagged EGL-4 (GFP::EGL-4) molecule we have been able to develop a molecular readout of long-term olfactory adaptation in the AWC5. Using this molecular readout of olfactory adaptation we have been able to perform both forward and reverse genetic screens to identify mutant animals that exhibit defective subcellular localization patterns of GFP::EGL-4 in the AWC6,7. Here we describe: 1) the construction of GFP::EGL-4 expressing animals; 2) the protocol for cultivation of animals for long-term odor-induced nuclear translocation assays; and 3) the scoring of the long-term odor-induced nuclear translocation event and recovery (re-sensitization) from the nuclear GFP::EGL-4 state.

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

Here we describe a molecular readout of long-term olfactory adaptation in Caenorhabditis elegans. The Protein Kinase G, EGL-4, is necessary for stable adaptation responses in the primary sensory neuron pair called AWC. During prolonged odor exposure EGL-4 translocates from the cytosol to nucleus of the AWC.

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps ...

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.

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.

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, ...

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this protocol focuses on how to image dividing neuroblasts, which are found underneath the epidermal cells and may be important for epidermal morphogenesis. Tissue formation is crucial for metazoan development&#160;and relies on external cues from neighboring tissues. C. elegans is an excellent model organism to study tissue morphogenesis in vivo due to its transparency and simple organization, making its tissues easy to study via microscopy. Ventral enclosure is the process where the ventral surface of the embryo is covered by a single layer of epithelial cells. This event is thought to be facilitated by the underlying neuroblasts, which provide chemical guidance cues to mediate migration of the overlying epithelial cells. However, the neuroblasts are highly proliferative and also may act as a mechanical substrate for the ventral epidermal cells. Studies using this experimental protocol could uncover the importance of intercellular communication during tissue formation, and could be used to reveal the roles of genes involved in cell division within developing tissues.

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes how to image dividing cells within a tissue in Caenorhabditis elegans embryos. While several protocols describe how to image cell division in the early embryo, this protocol describes how to image cell division within a developing tissue during mid late embryogenesis.

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this ...

Laser Nanosurgery of Cerebellar Axons In Vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to understand the mechanism that promotes neuronal regeneration, an in-depth analysis of the neuronal types that can remodel after injury is needed. Several studies showed that damaged climbing fibers are capable of regrowing also in adult animals1,2. The investigation of the time-lapse dynamics of degeneration and regeneration of these axons within their complex environment can be performed by time-lapse two-photon fluorescence (TPF) imaging in vivo3,4. This technique is here combined with laser surgery, which proved to be a highly selective tool to disrupt fluorescent structures in the intact mouse cortex5-9.
This protocol describes how to perform TPF time-lapse imaging and laser nanosurgery of single axonal branches in the cerebellum in vivo. Olivocerebellar neurons are labeled by anterograde tracing with a dextran-conjugated dye and then monitored by TPF imaging through a cranial window. The terminal portion of their axons are then dissected by irradiation with a Ti:Sapphire laser at high power. The degeneration and potential regrowth of the damaged neuron are monitored by TPF in vivo imaging during the days following the injury.

Laser Nanosurgery of Cerebellar Axons In Vivo

Two-photon imaging, coupled to laser nanodissection, are useful tools to study degenerative and regenerative processes in the central nervous system with subcellular resolution. This protocol shows how to label, image, and dissect single climbing fibers in the cerebellar cortex in vivo.

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to ...

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 ...