We thank Mei Ann Lim for critical reading of the manuscript and other members of our lab for their discussions throughout the course of this work. This work was funded by Yamada Science Foundation and Royal Society Short Visit Fellowships and EU Marie Curie International Incoming Fellowship GRR to K.K., and BBSRC Project Grant (BB/H002278/1) and Wellcome Trust Equipment Grant (073228/Z/03/Z) to A.H.

1. Collection of Staged Larvae
<ol>
	<li>Place 15 female and 15 male adult flies in a cylindrical Perspex cage to lay eggs on a Petri dish with agar and grape juice supplemented with a small scoop of yeast paste for 3 hr at 25 &deg;C. Change the plate 3-4 times a day, and discard the first plate (i.e. from the O/N egg-lay). Discard also the plates from the first day. From the second day on, keep the plates with eggs at 25 &deg;C. After 7 days of collecting eggs, start with a new set up of parent flies.</li>
	<li>After about 24 hr, larvae hatch from the eggs. By hooking larvae with a thinned paintbrush or with a pair of forceps, transfer 35 larvae from the grape agar to a vial containing standard fly food (10 ml) (Figure 1A).</li>
	<li>Maintain the vials containing larvae for 3 days (96 hr After Egg Laying, AEL) at 25 &deg;C.</li>
</ol>
2. Dissection of Larval Ventral Nerve Cords for Culture
<ol>
	<li>Clean bench and hands with 70% ethanol. Soak 4 staining blocks, 2 pairs of forceps and a needle with 70% ethanol, and let air dry.</li>
	<li>Prepare 4 staining blocks with the following media: one with Distilled Water (DW) to clean and to pool cleaned larvae; a second one with 2 ml of Shield and Sang M3 insect medium with 1% penicillin and Streptomycin, Ecdysone-free (M3 PS medium) to dissect the larvae; a third with 2 ml of M3 PS medium to pool dissected ventral nerve cords (VNCs); and a fourth with 2 ml of M3 PS medium to stab the VNCs. All of the reagents must be pre-warmed to RT.</li>
	<li>Add water to the vial containing 96 hr AEL larvae. Then, by using a spatula, gently spread food with larvae from the vial on to a wet paper towel. Transfer 10 larvae to a staining block containing DW to wash off the food. Replace DW 6 times. Then replace with M3 PS medium.</li>
	<li>Transfer one of the larvae to a staining block containing 2 ml of M3 PS medium.</li>
	<li>Under a dissecting microscope, dissect a larval brain carefully using standard methods 12, but to minimize tissue damage, take special care by proceeding in the following way. Place the larva dorsal side up, and hold the dorsal side at one third from the anterior end with 2 pairs of forceps (one in each hand). Whilst holding the anterior section, pull away the posterior end with the forceps, and tear the epidermis. The brain and the VNC should be visible from the posterior edge of the anterior half. Carefully isolate the brain and VNC complex by removing the fat body, gut and epidermis. When the VNCs are damaged too much, the VNCs will degenerate fully or partially even without stabbing injury. Take into account the following points:</li>
</ol>
<ul>
	<li>Do not pull or tear the imaginal discs and peripheral nerves from the thoracic VNC as this will damage the VNC. To cut the peripheral nerves, hold the peripheral nerves with two pairs of forceps placed close each other, and then pull the nerves with the forceps. Imaginal discs and mouth parts can be left attached to the VNC.</li>
	<li>Leave the ring gland and lymph gland attached to the brain.</li>
	<li>Cut the gut at the point closest to the brain, where not much food is contained.</li>
</ul>
<ol start="6">
	<li>By using a P20 pipetman with the tip cut off and widened a little, transfer the VNC to a staining block containing fresh M3 PS medium. Before transferring the VNC, first pipette up and down only the medium used for the dissection in order to prevent the VNC from sticking to the tip.</li>
</ol>
3. Stabbing Injury to the Drosophila Larval VNC
This section describes how to perform stabbing injury to the VNCs, and to prepare stabbed-VNCs for time-lapse analysis and immunostaining. Post-stabbing culture conditions and duration are described in section 5 (for time-lapse analysis) and in section 6 (for immunostaining).
<ol>
	<li>After pooling enough VNCs (4-5 for time-lapse and over 24 for immunostaining), transfer one VNC to a clean staining block containing M3 PS.</li>
	<li>Orient the VNC so that the dorsal neuropile is in view under the dissecting microscope (Figure 1B). This is the most natural orientation for VNCs to lie in when attached to the optic lobes.</li>
	<li>Stab manually using a tungsten needle under the dissection microscope, from the dorsal side virtually at a right angle and aiming for the abdominal half of the VNC (Figure 1B).</li>
</ol>
Take particular attention to the following:
<ul>
	<li>Stab only once.</li>
	<li>Stab until the needle hits the bottom of the glass. However, be careful not to damage the needle tip.</li>
	<li>The stem of the needle holder must not touch the medium during stabbing.</li>
	<li>The wound is not visible under the dissection microscope.</li>
</ul>
Note: Stab-injury unrelated degeneration
The procedure can result in cellular degeneration within the VNC distinct from the stabbing lesion. Degeneration results in holes in the neuropile, and occasionally holes have been observed in the cortex of non-stabbed specimens. Specimens with degeneration affecting the neuropile or widespread degeneration affecting also the VNC surface must be discarded. Degeneration can be identified as follows:
<ul>
	<li>Rough surface, especially at the lateral/ventral area of the thorax, at 22 hr of stabbing. In extreme cases, the VNC looks protruded.</li>
	<li>Holes or vacuoles in the thoracic neuropile, which can be visible as holes in background signal with fluorescence immunostaining.</li>
</ul>
4. Maintenance of Needle and Forceps
<ul>
	<li>Routinely check whether the needle tip is sharp enough. The needle tip can be bent or blunt after using it for several experiments. In this case, sharpen the tip using an Arkansas stone or equivalent.</li>
	<li>The tips of the forceps must meet perfectly. The tips can get damaged with use. In this case, adjust the tip using an Arkansas stones or equivalent.</li>
</ul>
5. Culture and Time-lapse Recording of Stabbed VNCs
To visualize the axonal neuropile in the living VNC, a GFP protein trap line that labels all axons - G9 13 - can be used, and to visualize all glial cells (except the midline glia) the glial driver repoGAL4 can be used to express the UASdsRed S197Y 14 reporter. By crossing G9 flies to UASdsRedS197Y;;repoGAL4 flies, VNCs from progeny larvae are obtained with green axons and red glia that can be recorded in living tissue.
<ol>
	<li>After pooling 4-5 VNCs, transfer 1 VNC to a clean staining block containing 2 ml of M3 PS, and stab the abdominal half of the VNC as indicated in section 3.</li>
	<li>Transfer the stabbed VNC to a Poly-L-lysin coated 3.5 mm glass bottom Petri dish containing 1 ml of M3 PS. Place the VNCs dorsal side down. Gently push the VNC using the flat side of a pair of forceps to allow the VNC to stick to the dish.</li>
	<li>Gently add 1 ml of M3 PS 15% FBS rendering the final concentration of FBS to 7.5%.</li>
	<li>Acquire images using laser scanning confocal microscopy. Scan the VNC, collecting a Z-section series throughout its entire thickness. We used a Leica SP2 inverted confocal microscope with a temperature controlled environmental chamber. Any equivalent confocal microscope should work, however it may require optimization of the settings for the scanning. The settings for our time-lapse confocal microscopy were as follows: Temperature of environmental chamber: 25 &deg;C; 20X objective with 4 times zoom; scanning mode xyzt, with 512x512 pixel resolution, z=1 &mu;m steps and 1-hour or 2-hour intervals.</li>
</ol>
<ul>
	<li>The Leica SP2 confocal microscope has a limitation in file size for scanning. With these settings, 8-9 timepoints can be scanned. With other confocal microscopes, it ought to be possible to acquire image stacks for longer time points, i.e. over the span of up to 24 hr.</li>
</ul>
6. Culture and Immunostaining of Stabbed and Fixed VNCs
<ol>
	<li>Prepare a 24-well tissue culture plate containing 500 &mu;l of M3 PS 7.5% FBS per well.</li>
	<li>Repeat dissection of more than 24 VNCs for a 24-well tissue culture dish.</li>
	<li>By using the P20 pipetman with the tip cut-off, transfer 12 non-stabbed VNCs from the pool to the culture dish, using 1 VNC per well.</li>
	<li>Stab the rest of VNCs in the pool as described in section 3. Transfer each stabbed VNC to a well each.</li>
	<li>Place the 24-well dish in the 25 &deg;C incubator for a desired duration depending on the experiment. Cell integrity is unaltered for at least 24 hr.</li>
	<li>After culture, transfer the 12 VNCs in 250 &mu;l of 4% formaldehyde PEM in a 1.5 ml tube by using the P20 pipetman with the tip cut-off. Then, the fixative is carefully pipetted out, taking care not to damage the VNCs. A small amount of fixative sufficient to cover the samples should remain in each tube. Subsequently, add fresh fixative, and gently agitate for 50 min at RT.</li>
	<li>Rinse 2 times with 0.3% Triton X PBS, and then wash 2 times for 10 min with 0.3% Triton X PBS at RT.</li>
	<li>Block VNCs by incubating with 10% normal goat serum in 0.3% Triton X PBS for 1 hr at RT. The samples can be stored at 4 &deg;C for at least one month.</li>
	<li>Incubate VNCs with primary antibodies for more than 20 hr at 4 &deg;C.</li>
	<li>Rinse 2 times with 0.3% Triton X PBS, then wash 3 times for 10 min with 0.3% Triton X PBS at RT.</li>
	<li>Incubate VNCs with secondary antibodies for more than 16 hr at 4 &deg;C.</li>
	<li>Rinse 2 times with 0.3% Triton X PBS, then wash 3 times for 10 min with 0.3% Triton X PBS at room RT (in the dark if using fluorescent secondary antibodies).</li>
	<li>Replace PBS with 50% glycerol: 50% PBS for at least an hour.</li>
	<li>Then replace with 80% glycerol: 20% PBS for at least an hour.</li>
	<li>Mount a VNC in a window made on 2 layers of cello-tape (approximately 0.06 mm) on a microscopy glass slide. Adjust the orientation, and place a coverslip (18x18 mm) over the window. Two VNCs can be mounted on a slide using this method.</li>
</ol>
Using a variety of primary antibodies, different cellular responses to injury can be analyzed (e.g. changes in cell number and cell shape).
7. Analyze Data Using ImageJ and a Range of Plugins
If the VNC does not have stabbing unrelated degeneration, the sample must be counted in for the analysis regardless of lesion size. As stabbing is done manually, the lesion size differs each time. Thus it is important to analyze statistically the effect of stabbing, whenever possible.
<ol>
	<li>Wound size measurement. The size of the wound is an indicator of repair 15. The stabbing lesion is visible as devoid of GFP expression. Lesion area can be measured lapse data using freely available ImageJ software, as follows:
	<ol>
		<li>Using ImageJ, open a stack of confocal images from "File" menu, select "Import" and from here select "Image sequence". This will turn the collection of individual images into a stack.</li>
		<li>Next change the "stack" into a "Hyperstack" by going to "Image" menu, select "Hyperstack", then select "Stack to Hyperstack".</li>
		<li>Set the voxel size by using "Properties" from "Image" menu. In data acquired using the Leica SP2 confocal microscope, the voxel size can be obtained from the scanning-software, and from the metadata text file saved together with images. With our setting as described in Section 3, pixel dimensions are xy=0.366211 &mu;m and z=0.99709 &mu;m.</li>
		<li>By examining all the slices in one time point, draw the maximum outline of the lesion area with the polygon-selection tool in the tool bar. This is the Region Of Interest (ROI).</li>
		<li>Add the ROI to the ROI manager by going to "Analyze" menu, scroll down and select "Tools", then "ROI manager", then "add".</li>
		<li>Repeat this for the all the time points.</li>
		<li>Click the "Measure" button in the ROI manager to obtain the area size of each ROI.</li>
	</ol>
	</li>
	<li>Correction of VNC movement during time-lapse recordings. "Stackreg" and "Turboreg" plugins 16 (from the ImageJ website <a href="http://rsbweb.nih.gov/ij/" target="_blank">http://rsbweb.nih.gov/ij/</a>): these plugins allow to correct small sample movements during time-lapse. Build a stack with an equivalent representative optical section through all the time-points and apply the plugins. It helps to visualize how the lesion changes over time. The details on these methods and the instruction manual are available from the research group that developed these plugins. (<a href="http://bigwww.epfl.ch/thevenaz/stackreg/" target="_blank">http://bigwww.epfl.ch/thevenaz/stackreg/</a>).</li>
	<li>Automatic counting of glial cells. "DeadEasy glia" plugin (<a href="http://www.biosciences-labs.bham.ac.uk/hidalgo/Software.html" target="_blank">www.biosciences-labs.bham.ac.uk/hidalgo/Software.html</a>): this was developed to count the number of REPO-positive glial cells in the Drosophila larval VNC and to examine the change in glial number caused by stabbing injury17. The plug-in works accurately, with an error of 0.1% false positives and 4.3% false negatives 17. To use this plug-in, first label all the glia (except the midline) in the specimen using immunofluorescence with anti-REPO antibodies. Then install the plug-in in ImageJ, open a stack of confocal images and run plug-in. It counts glial cells automatically in about 30 seconds.</li>
	<li>Automatic counting of apoptotic cells. "DeadEasy Caspase larvae" plug-in (<a href="http://www.biosciences-labs.bham.ac.uk/hidalgo/Software.html" target="_blank">www.biosciences-labs.bham.ac.uk/hidalgo/Software.html</a>) 18: This was developed to count the number of cells labeled with anti-cleaved-Caspase3, and a new version was adapted for the larval VNCs. Stabbing injury causes an increase in programmed cell death 11. To use this plug-in, first label apoptotic cells in the specimen using immunofluorescence with anti-cleaved-Caspase-3 antibodies. Then install the plug-in in ImageJ, open a stack of confocal images and run plug-in. It counts apoptotic cells automatically in about 30 seconds.</li>
</ol>
8. Reagents
<ol>
	<li>Culture medium: Shield and Sang M3 insect medium, 7.5% fetal bovine serum, 1% penicillin and 1% Streptomycin.</li>
	<li>0.01% Poly-L-lysin in sterile DW.</li>
	<li>Fixative: 4% formaldehyde, Ultra pure in PEM (0.1 M PIPES, 2 mM EGTA, 1 mM MgSO4) solution.</li>
	<li>Blocking solution for immunostaining: 10% normal goat serum in 0.3% Triton X PBS.</li>
	<li>Anti-Glutamine Synthetase 2 antibodies: 1:250 in blocking solution.</li>
	<li>Anti-GFP antibodies: 1:1,000 in blocking solution.</li>
	<li>Anti-REPO antibodies: 1:250 in blocking solution.</li>
	<li>Anti-ELAV antibodies: 1:250 in blocking solution.</li>
	<li>Anti-cleaved Caspase 3 antibodies: 1:1,000 in blocking solution.</li>
	<li>Secondary antibodies: 1:250 in blocking solution.</li>
</ol>

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No conflict of interest declared.

We have established a protocol for stabbing injury to the Drosophila larval CNS to investigate the cellular responses to injury, repair and regeneration. Larval VNCs are dissected and stabbed, after which they are filmed with time-lapse microscopy or fixed for fluorescence immunostaining to visualize glia and neurons, apoptosis or cell division. The progression of the lesion over time can be measured. This method is accompanied by purposely developed software for the quantitative and statistical analysis of cell number changes upon injury and during repair.
We stabbed 96-hour AEL larvae (and these were either fixed or allowed to develop further), a developmental stage prior to the transition to pupa and then adult fly. At 96 hr, VNCs are large enough for stabbing; they are in the middle of third instar stage, and thus are not undergoing pupation yet; and the nervous system is already fully functional similar to that of the adult. It might be possible to stab slightly later in larvae and the precise timing must be chosen to suit the research question. However, depending on the questions addressed, it will generally be necessary to culture the VNCs for some time to observe the cellular responses to injury. Some time after 120 hr AEL pupation starts, a period when the CNS is remodeled and is thus best avoided. Thus the time window for stabbing plus culture in the larva is rather restricted. Using larvae still has a great technical advantage over using adults: whilst, similarly to the adult, the nervous system is already fully functional, analyzing the response to injury is considerably easier and faster in larvae.
When comparing the size and morphology of brains and VNCs dissected and cultured in a dish to VNCs that are dissected at an equivalent later time point without culturing in a dish, it appears that development is slower in culture than in vivo. In other respects, development carries on normally in culture, tissue integrity is preserved and cells are alive showing multiple responses. Wound expansion, neuropile repair and glial proliferation take place within 22 hr post-stabbing. This shows that cellular responses to injury take place in culture and there is most likely no need to maintain the explants for longer than a day. If longer term culture were desired, the protocol may require further optimization such as using a culture plate insert5.
It is important to identify good quality samples from the degenerating ones. Optimizing this protocol takes some skill and inevitably degeneration will occur in some specimens. The VNC can acquire a 'cauliflower' appearance that reflects a breakdown of tissue integrity (Figure 5B). Degeneration is also recognized by the vacuolization of the VNC independently of the stabbing, which can be present in intact, non-stabbed specimens (Figure 5C). These samples must be discarded. Degeneration is most likely caused by rough dissection, which can tear nerves and the protective layer of surface glia. Thus great care must be taken to dissect gently. Other influencing factors include the culture medium, which must be kept clean and with antibiotics, adhering strictly to the washes and timings as indicated in the protocol. Finally, the length of the needle and needle holder, size and sharpness of the needle are very important. The needle holder can contaminate the culture medium and a blunt needle can cause a large injury that cannot repair itself and will lead to degeneration. It is important to routinely maintain the needle sharp.
In this protocol, we took advantage of a protein trap line of flies to visualize the neuropile in parallel to the visualization of glial processes using the traditional GAL4-UAS system. These tools could also be combined with other binary expression systems such as the LexA 19 and Q-system 20, which are independent of GAL4. This would enable the analysis of interactions for instance between axons and glial processes, in response to injury. By further combining it with other genetic tools such as reporters for dendrites 21 or calcium influx 22, this method provides a great opportunity to analyze the cell biology behind the injury response of glial cells, neuronal axons and dendrites in the CNS. Finally, this method can be combined with standard genetics, mutations and over-expression of genes, to test gene function in responses to injury and regeneration.
This protocol has successfully led to the discovery of a gene network underlying the regenerative glial response to CNS injury 11. Given the evolutionary conservation of gene function, unraveling these cellular events and gene functions in fruit-flies is likely to provide significant insights into the understanding of mammalian CNS response to injury and regeneration.

Regeneration in animals reveals that cells sense when organism growth is ordered and complete, and how structural integrity of an organism is achieved and maintained. Understanding these mysterious abilities of cells is of great interest in biology. Promoting regeneration is one of the key goals for medical neuroscience. In humans, the damaged central nervous system (CNS) does not regenerate. Rodent models of spinal cord injury are used to understand how cells respond to injury. However, ethical concerns, high costs and the slow life cycle of the animals constrain progress.
The fruit-fly Drosophila is a widely used model organism in developmental biology and neuroscience. Thanks to its powerful genetic tools and short life cycle, Drosophila has recurrently led to the discovery of gene functions and gene networks with relevance for the understanding of humans and disease. There is abundant evidence of evolutionary conservation of gene function from flies to humans.
Over recent years, several paradigms for nervous system injury have been established in Drosophila. Some consist of damaging peripheral nerves that run along the wing or axotomy of peripheral nerves, including sensory 1 and motor axons 2 . However, the peripheral and central nervous systems differ in many respects, and it is well known that in many animals the peripheral nervous system can regenerate whereas the CNS cannot. Thus, to understand CNS regeneration, direct injury to the CNS is more appropriate. Stabbing injury with a needle has been successfully applied to the Drosophila adult brain to investigate the response to injury 3,4. Using another approach, Ayaz et al. severed CNS axons in cultured adult brains with a Piezo power microdissector, and analyzed their regeneration for 4 days 5. The advantage of these later experimental set ups is that they focus on the brain, which is undoubtedly of great interest. The disadvantage is that the brain is a lot more complex than the VNC, which deals only with motor and sensory control. Working on the adult brain is also more time consuming. The larva is ready for experiments in 4 days, whereas it takes about 10 days for adult eclosion, and then additional 5 days for their maturation. The adult brain is also more difficult to handle, because it is encapsulated in thick cuticle, which is difficult to remove. The VNC has the further attraction of being functionally equivalent to the vertebrate spinal cord.
The Drosophila larva is widely used as a model organism for neuroscience 6-9. Although strictly speaking the larva is in a developmental phase, it can also be considered a fully functional animal. The larva has locomotion, multiple senses including olfaction, taste and nociception, and learning and memory. Thus, the larval VNC was chosen as the ideal model for CNS injury.
Larval and pupal VNCs can be dissected and cultured in dish, still retaining cellular integrity for up to 24 hr 10. This suggested that injury could be applied to the VNC, which could then be recorded in time-lapse for over this period of time, or cultured and fixed at any desired time point within this period.
Here, we present an experimental paradigm for CNS injury using the Drosophila larval VNC. The VNC is first dissected from the larva, and stabbing injury is applied manually with a tungsten needle. The VNC is then placed on a glass-bottom coverslip, and filmed with time-lapse laser scanning confocal microscopy. Alternatively, the VNCs can be cultured for a desired period of time, and the cellular effects of injury can be analyzed in fixed specimens using immunostaining and confocal microscopy. The wound area can be measured, and quantitative analysis of cell number (cell proliferation and cell death) can be carried out with purposely developed software. Large sample sizes can be easily handled, resulting in statistically validated results. These methods have been successfully combined with the powerful genetics of Drosophila to discover a gene network underlying the glial regenerative response to CNS injury 11.

<table border="1">
	<tbody>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Staining block</td>
			<td>Brunel Microscope</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Forceps No. 5</td>
			<td>e.g. Fine Science Tools</td>
			<td>e.g. 11251-20</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tungsten needle: rod diameter, 0.5 mm; tip size, 1 &mu;m; length, 2 inch</td>
			<td>Roboz Surgical Instrument</td>
			<td>RS-6065</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>Roboz Surgical Instrument</td>
			<td>RS-6060</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Arkansans stones: Repair Kit for Dumont Forceps</td>
			<td>Fine Science Tools</td>
			<td>29000-00</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>35 mm Petri dish with 27 mm glass base</td>
			<td>Iwaki</td>
			<td>3930-035</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Leica SP2-AOBS confocal inverted microscope with environment chamber</td>
			<td>Leica</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>Shield and Sang M3 insect medium (ecdysone free)</td>
			<td>Sigma</td>
			<td>S3652-500 ml</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Penicillin and streptomycin</td>
			<td>Invitrogen</td>
			<td>15070-063</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline (PBS)</td>
			<td>See 12</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Sigma</td>
			<td>F7524</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Poly-L-lysin</td>
			<td>Sigma</td>
			<td>P1399-25mg</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Formaldehyde, 10%, methanol free, Ultra Pure</td>
			<td>Polysciences</td>
			<td>04018-1</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mouse anti-glutamine synthetase antibodies</td>
			<td>Millipore</td>
			<td>MAB302</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rabbit anti-GFP antibodies</td>
			<td>Life technologies</td>
			<td>A11122</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mouse anti-REPO antibodies</td>
			<td>Developmental Studies Hybridoma bank</td>
			<td>8D12</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rat anti-ELAV antibodies</td>
			<td>Developmental Studies Hybridoma bank</td>
			<td>7E8A10</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rabbit anti-active caspase 3 antibodies</td>
			<td>Abcam</td>
			<td>ab13847</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Normal goat serum</td>
			<td>Vector Laboratories</td>
			<td>S-1000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Anti-rabbit Alexa Fluor 488</td>
			<td>Life technologies</td>
			<td>A11034</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Anti-mouse Alexa Fluor 647</td>
			<td>Life technologies</td>
			<td>A21236</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Anti-rat Alexa Fluor 647</td>
			<td>Life technologies</td>
			<td>A21247</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

an injury paradigm to investigate central nervous system repair in drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

Here we show how to perform stabbing injury to the Drosophila larval VNC and analyze the cellular responses to injury using time-lapse and immunostaining confocal fluorescence microscopy.
For time-lapse data, the lesion is visualized as a GFP-negative area within the neuropile of specimens bearing the G9 axonal marker (Figure 2). Shortly after stabbing, small GFP-negative areas, which look like holes or vacuoles, start to appear (Figure 2A). Such GFP-negative areas generally enlarge up to around 6 to 8 hr after stabbing (Figure 2A). Subsequently, the GFP-negative areas shrink and may even disappear (Figure 2B). By 22 hr after stabbing, the area occupied by the wound is generally smaller than the maximum area was at 6 to 8 hr after stabbing (Figures 2A, B). Similarly the DsRed-negative area in glial processes initially increases too, but shrinks by 22 hr after stabbing. Interestingly, often DsRed-positive glial processes fill the GFP-negative holes in the neuropile prior to their disappearance (Figure 2C).
Using immunostaining in fixed specimens, fine glial processes and their response to injury can be visualized with better resolution than with time-lapse images. This can be done, for instance, using the repoGAL4 glial driver to induce the expression of a membrane tethered reporter (e.g. UAS-mCD8GFP) in flies to visualize all glial cells (except the midline glia). This can also be combined with other glial markers, such as anti-GS2, which labels neuropile-associated glial cells (Figures 3A, B). Here we show that although the ventral nerve cord is stabbed from the dorsal side, stabbing injury results in a dent ventrally (Figure 3A, arrowheads). Stabbing appears to affect more severely the neuropile and neuropile-associated glial cells than surface and cortex glial cells (Figure 3B). Glial processes appear disorganized in the neuropile, whereas they still maintain their mesh like organization in the cortex. Injury results in GS2-positive cellular debris, which is distinct from normal cells as debris fragments are much smaller and not connected to a cell body. This reveals damage to neuropile-associated glial cells (Figure 3B). Degenerating neuropile-associated glial cells were also observed by electron microscopy 15. Anti-cleaved-Caspase 3+ apoptosis staining is observed in neuropile-associated glial cells 15, showing they undergo apoptosis upon stabbing injury. Neuropile associated glial cells were also shown to phagocytose neuronal debris 15, and GS2+ signal might also reveal the engulfment of apoptotic neurons by glial processes. Cleaved-Caspase+ apoptotic cells are also observed in the cortex, at least some of which correspond to Elav+ neurons (Figure 3C).
Using fixed samples and immunostaining also enables quantitative analyses of cellular responses to injury, such as effects in proliferation and apoptosis. For this, we purposely developed two ImageJ plug-ins, DeadEasy Caspase Larva and DeadEasy Glia, to count automatically the number of glial cells and apoptotic cells, respectively. They have been validated to work on larval VNCs and are very accurate. Using these, it is possible to observe an increase in the number of REPO-positive glial cells caused by stabbing injury, by 22 hr (Figure 4A)15. There is also an increase in the number of anti-cleaved-Caspase-3 apoptotic cells by 6 hr post-stabbing (Figure 4B)15. Such increase in glial proliferation and apoptosis in response to injury in the Drosophila CNS is reminiscent of the injury response in the vertebrate CNS.
A crucial aspect of this protocol is the quality of the VNC dissections. It is difficult to tell at the time of dissection whether the VNCs have been damaged or not in the process. It is important to take particular care to perform gentle dissections. However, bad quality samples will inevitably be produced, and it is essential that these are identified at later stages and discarded. In our hands, VNC degeneration appears to be unrelated to injury, but instead caused by rough dissection. We have not observed a critical size in the injury wound, and we analyze all injured samples that are not degraded. When VNCs maintain their integrity, the VNC surface tends to look smooth and shiny, and no holes in the neuropile are observed (Figure 5A). Degradation of VNCs can be recognized by 24 hr post-dissection from the rough surface of the VNC ventral and lateral areas (Figure 5B). Degradation of the neuropile unrelated to stabbing injury can also occur, and it is recognized as neuropile holes in the background signal of immunostained specimens. Samples with these signs of degeneration must be discarded (Figure 5C). In our hands, the success rate for dissection and injury in intact and stabbed control (yw) flies are both around 70%. This rate will vary with the skill of the person performing the experiments, and with the genotype.
<img alt="Figure 1" src="/files/ftp_upload/50306/50306fig1.jpg" /> 
 Figure 1. Schematic drawing of larval staging and stabbing injury. (A) At day 0, flies are allowed to lay eggs in a grape juice Petri dish for 3 hr. At day 1, the hatched first instar larvae (L1) are collected and placed in vials containing standard yeast food, where they are kept for another 3 days. At day 4, larvae are collected from food vial, and used for the stabbing experiments. (B) Lateral view of the larval central nervous system. The abdominal region of the ventral nerve cord is stabbed with a needle from the dorsal side. Anterior is to the left, and posterior to the right.
<img alt="Figure 2" src="/files/ftp_upload/50306/50306fig2.jpg" /> 
 Figure 2. Time-lapse progression of VNC lesion. Confocal microscopy on live VNCs with GFP-labeled axons and DsRed-labeled glial cells. Genotype: UASDsRed/+;G9/+;repoGAL4/+. (A) The lesion is recognized by the lack of fluorescence on the axonal neuropile. After stabbing, GFP-negative holes appear, thereafter increasing in size and number. Projections of 5 optical sections from each time point after stabbing injury. (B) The axonal neuropile lesion shrinks from 9 hr after stabbing. These images are single optical sections from different time points after stabbing injury. To identify equivalent positions in each sample, the same slice number from each time point in the stack of time-lapse data was chosen. By comparing the patterns visualized with G9 and repoGAL4&gt;UASmCD8GFP in the area not affected by the stabbing, the slices were verified as being from equivalent positions in the Z axis. Dashed lines in (A,B) indicate the edge of the wound. (C) High magnification of wounds on the neuropile. GFP-negative holes were filled with DsRed-positive glial processes, and subsequently disappeared (arrowheads). Horizontal view. Anterior up.
<img alt="Figure 3" src="/files/ftp_upload/50306/50306fig3.jpg" /> 
 Figure 3. Cellular characterization of VNC lesion. VNCs bearing a membrane tethered GFP reporter for all glial cells (except midline glia) stained with anti-GFP and anti-GS2, a neuropile-associated glial cell marker. Genotype: +/+;UASmCD8GFP/+;repoGAL4/+. (A) VNCs were stabbed from the dorsal side, but this causes a dent ventrally (arrowheads). Sagittal view of single optical sections, dorsal up and anterior left. (A, B) Stabbing injury affects more severely neuropile-associated glial cells than cortex and surface glial cells. Injury results in GS2-positive cellular debris (arrowheads in B). (B) Horizontal view of single optical sections, anterior up. Arrowheads point to GS2+ cell debris. (C) Colocalisation of the apoptotic marker anti-cleaved-Caspase-3 and the neuronal marker anti-Elav showing that upon injury some of the dying cells in the cortex are neurons. np, neuropile; cx, cortex.
<img alt="Figure 4" src="/files/ftp_upload/50306/50306fig4.jpg" /> 
 Figure 4. Automatic counting of glial cells and apoptotic cells using DeadEasy. (A) Example of using 'DeadEasy Larval glia' to count REPO positive glial cells. Left: VNC ventral halves stained with anti-REPO antibodies; middle: output from DeadEasy Larval glia showing cells identified by the plug-in; right: merge. The numbers on the right are the number of REPO-positive cells counted automatically in an entire stack of optical confocal sections in about 30 seconds. The number of glia increases in the stabbed VNCs. (B) Example of using 'DeadEasy Caspase larva'. Left: projections of 5 optical confocal sections stained with anti-cleaved-CASPASE 3 antibodies; middle: output showing cells identified by the plug-in; right: merge. The numbers on the right are the number of CASPASE-positive cells counted automatically in an entire stack of optical confocal sections in about 30 seconds. The number of apoptotic cells increases in the stabbed VNC.
<img alt="Figure 5" src="/files/ftp_upload/50306/50306fig5.jpg" /> 
 Figure 5. Examples of degeneration. (A) A healthy VNC. There are no signs of degeneration. (B) The side of the thorax is degenerating (arrowheads) in a stabbed specimen. Asterisk indicates lesion site. (C) Holes in the thoracic neuropile (white arrowhead) and cortex (orange arrowhead) in a non-stabbed VNC. Specimens with damaged neuropile and extensive vacuolization must be discarded. Here VNCs were stained with the neuropile associated glial marker anti-Ebony. Horizontal view, anterior up.

Watch this Scientific Journal Video about An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila at JoVE.com

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

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13.2K Views.  University of Birmingham. An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

Video: An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

Production of Lentiviral Vectors for Transducing Cells from the Central Nervous System

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing therapeutic approaches. Lentiviral vectors are attractive tools in transduction of neurons and other cell types in CNS as they transduce both dividing and non-dividing cells, support sustained expression of transgenes, and have relatively large packaging capacity and low toxicity 1-3. Lentiviral vectors have been successfully used in transducing many neural cell types in vitro 4-6 and in animals 7-10.
Great efforts have been made to develop lentiviral vectors with improved biosafety and efficiency for gene delivery. The current third generation replication-defective and self-inactivating (SIN) lentiviral vectors are depicted in Figure 1. The required elements for vector packaging are split into four plasmids. In the lentiviral transfer plasmid, the U3 region in the 5' long terminal repeat (LTR) is replaced with a strong promoter from another virus. This modification allows the transcription of the vector sequence independent of HIV-1 Tat protein that is normally required for HIV gene expression 11. The packaging signal (&Psi;) is essential for encapsidation and the Rev-responsive element (RRE) is required for producing high titer vectors. The central polypurine tract (cPPT) is important for nuclear import of the vector DNA, a feature required for transducing non-dividing cells 12. In the 3' LTR, the cis-regulatory sequences are completely removed from the U3 region. This deletion is copied to 5' LTR after reverse transcription, resulting in transcriptional inactivation of both LTRs. Plasmid pMDLg/pRRE contains HIV-1 gag/pol genes, which provide structural proteins and reverse transcriptase. pRSV-Rev encodes Rev which binds to the RRE for efficient RNA export from the nucleus. pCMV-G encodes the vesicular stomatitis virus glycoprotein (VSV-G) that replaces HIV-1 Env. VSV-G expands the tropism of the vectors and allows concentration via ultracentrifugation 13. All the genes encoding the accessory proteins, including Vif, Vpr, Vpu, and Nef are excluded in the packaging system. The production and manipulation of lentiviral vectors should be carried out according to NIH guidelines for research involving recombinant DNA (<a href="http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf" target="_blank">http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf</a>). An approval from individual Institutional Biological and Chemical Safety Committee may be required before using lentiviral vectors. Lentiviral vectors are commonly produced by cotransfection of 293T cells with lentiviral transfer plasmid and the helper plasmids encoding the proteins required for vector packaging. Many lentiviral transfer plasmids and helper plasmids can be obtained from Addgene, a non-profit plasmid repository (<a href="http://www.addgene.org/" target="_blank">http://www.addgene.org/</a>). Some stable packaging cell lines have been developed, but these systems provide less flexibility and their packaging efficiency generally declines over time 14, 15. Commercially available transfection kits may support high efficiency of transfection 16, but they can be very expensive for large scale vector preparations. Calcium phosphate precipitation methods provide highly efficient transfection of 293T cells and thus provide a reliable and cost effective approach for lentiviral vector production.
In this protocol, we produce lentiviral vectors by cotransfection of 293T cells with four plasmids based on the calcium phosphate precipitation principle, followed by purification and concentration with ultracentrifugation through a 20% sucrose cushion. The vector titers are determined by fluorescence- activated cell sorting (FACS) analysis or by real time qPCR. The production and titration of lentiviral vectors in this protocol can be finished with 9 days. We provide an example of transducing these vectors into murine neocortical cultures containing both neurons and astrocytes. We demonstrate that lentiviral vectors support high efficiency of transduction and cell type-specific gene expression in primary cultured cells from CNS.

In this protocol we describe production, purification and titration of lentiviral vectors. We provide an example of lentiviral vector-mediated gene delivery in primary cultured neurons and astrocytes. Our methods may also apply to other cell types in vitro and in vivo.

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are tightly regulated, affecting the final dynamics of protein abundance. Various regulatory mechanisms exist at the translation step, rendering mRNA levels alone an unreliable indicator of gene expression. In addition, local regulation of mRNA translation has been particularly implicated in neuronal functions, shifting &#39;translatomics&#39; to the focus of attention in neurobiology. The presented method can be used to bridge transcriptomics and proteomics.
Here we describe essential modifications to the technique of polyribosome fractionation, which interrogates the translatome based on the association of actively translated mRNAs to multiple ribosomes and their differential sedimentation in sucrose gradients. Traditionally, working with in vivo samples, particularly of the central nervous system (CNS), has proven challenging due to the restricted amounts of material and the presence of fatty tissue components. In order to address this, the described protocol is specifically optimized for use with minimal amount of CNS material, as demonstrated by the use of single mouse spinal cord and brain. Briefly, CNS tissues are extracted and translating ribosomes are immobilized on mRNAs with cycloheximide. Myelin flotation is then performed to remove lipid rich components. Fractionation is performed on a sucrose gradient where mRNAs are separated according to their ribosomal loading. Isolated fractions are suitable for a range of downstream assays, including new genome wide assay technologies.

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

This protocol illustrates essential modifications of polyribosome fractionation in order to study the translatome of in vivo CNS samples. It allows global assessment of translation and transcription regulation through the isolation and comparison of total RNA to ribosome bound RNA fractions.

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

Anatomically Inspired Three-dimensional Micro-tissue Engineered Neural Networks for Nervous System Reconstruction, Modulation, and Modeling

Functional recovery rarely occurs&#160;following injury or disease-induced degeneration within the central nervous system (CNS)&#160;due to the inhibitory environment and the limited capacity for neurogenesis. We are developing&#160;a strategy to simultaneously address&#160;neuronal and axonal pathway loss within the damaged CNS. This manuscript presents the fabrication protocol for micro-tissue engineered neural networks (micro-TENNs), implantable constructs consisting of neurons and aligned axonal tracts spanning the extracellular matrix (ECM) lumen of a preformed hydrogel cylinder hundreds of microns in diameter that may extend centimeters in length. Neuronal aggregates are delimited to the extremes of the three-dimensional encasement and are spanned by axonal projections. Micro-TENNs are uniquely poised as a strategy for CNS reconstruction, emulating aspects of brain connectome cytoarchitecture and potentially providing means for network replacement. The neuronal aggregates may synapse with host tissue to form new functional relays to restore and/or modulate missing or damaged circuitry. These constructs may also act as pro-regenerative&#160;&#34;living scaffolds&#34; capable of exploiting developmental mechanisms for cell migration and axonal pathfinding, providing synergistic structural and soluble cues based on the state of regeneration. Micro-TENNs are fabricated by pouring liquid hydrogel into a cylindrical mold containing a longitudinally centered needle. Once the hydrogel has gelled, the needle is removed, leaving a hollow micro-column. An ECM solution is added to the lumen to provide an environment suitable for neuronal adhesion and axonal outgrowth. Dissociated neurons are mechanically aggregated for precise seeding within one or both ends of the micro-column. This methodology reliably produces self-contained miniature constructs with long-projecting axonal tracts that may recapitulate features of brain neuroanatomy. Synaptic immunolabeling and genetically encoded calcium indicators suggest that micro-TENNs possess extensive synaptic distribution and intrinsic electrical activity. Consequently, micro-TENNs represent a promising strategy for targeted neurosurgical reconstruction of brain pathways and may also be applied as biofidelic models to study neurobiological phenomena in vitro.

This manuscript details the fabrication of micro-tissue engineered neural networks: three-dimensional micron-sized constructs comprised of long aligned axonal tracts spanning aggregated neuronal population(s) encased in a tubular hydrogel. These living scaffolds can serve as functional relays to reconstruct or modulate neural circuitry or as biofidelic test-beds mimicking gray-white matter neuroanatomy.

Functional recovery rarely occurs&#160;following injury or disease-induced degeneration within the central nervous system (CNS)&#160;due to the inhibitory ...

Quantifying Acute Changes in Renal Sympathetic Nerve Activity in Response to Central Nervous System Manipulations in Anesthetized Rats

Renal sympathetic nerve activity (RSNA) and mean arterial pressure are important parameters in cardiovascular and autonomic research; however, there are limited resources directing scientists in the techniques for measuring and analyzing these variables. This protocol describes the methods for measuring RSNA and mean arterial pressure in anesthetized rats. The protocol also includes the approaches for accessing the brain during RSNA recordings for central nervous system (CNS) manipulations. The RSNA recording technique is compatible with pharmacologic, optogenetic, or electrical stimulation of the CNS. The approach is useful when an investigator will measure short-term (min to h) autonomic responses in non-survival experiments to correlate anatomically with CNS nuclei. The approach is not intended to be used to obtain chronic (survival) recordings of RSNA in rats. Discharges in RSNA, averaged rectified RSNA, and mean arterial pressure can be quantified and analyzed further using parametric statistical tests. Methods for obtaining venous access, recording mean arterial pressure telemetrically, and brain fixation for future histological analysis are also described in the article.

Methods for measuring sympathetic and cardiovascular responses to central nervous system (CNS) manipulations are important for advancing neuroscience. This protocol was developed to assist scientists with measuring and quantifying acute changes in renal sympathetic nerve activity (RSNA) in anesthetized rats (non-survival).

Renal sympathetic nerve activity (RSNA) and mean arterial pressure are important parameters in cardiovascular and autonomic research; however, there are ...