We thank Thomas Wang for developing the prototype von Frey filaments, Patrick J. Huang for improving the mechanical probe assay, the Bloomington Drosophila Stock Center for the control (w1118) and ppk-Gal4&#62;UAS-mCD8-GFP fly stocks, and Galko lab members for critically reading the manuscript. This work was supported by R21NS087360 and R35GM126929 to MJG.

1. Mechanical probe construction
<ol>
	<li>Cut each nitinol filament (Figure 1B), perpendicular to its long axis, to the specified length (Figure 1M&#8211;N) using a small wire cutter (Figure 1C). The filaments come in three different pre-set diameters (Figure 1B). 
	NOTE: The lengths specified here are a guide to achieve the approximate pressures indicated, using a similar protocol for construction of the mount. Ultimately, regardless of the length of the filament cut, and the depth of the hole in the mount, the filaments must be measured/calibrated on a balance to obtain the exact force/pressure value.</li>
	<li>Examine the tip of the filament under a stereomicroscope to ensure no sharp or irregular edges remain as these could cause tissue damage to the skin of the larvae and interfere with calibration.</li>
	<li>Manually smoothen the sharp edges of the mechanical probe using a sharpening stone until no sharp irregularities persist (Figure 1D).</li>
	<li>Make a hole towards the end of a wooden popsicle stick (Figure 1E) using a hypodermic needle (see Table of Materials). Insert the needle at least halfway through the height of the popsicle stick (Figure 1E). This creates a chamber for insertion of the nitinol filament.</li>
	<li>Apply wood glue to a single nitinol filament (Figure 1F) and insert the glue-coated filament into the needle slot in a wooden popsicle stick (Figure 1G). Allow to dry for ~5 h.</li>
	<li>Calibrate each mechanical probe by pressing it against a scale until the mechanical probe bends (Figure 1H&#8211;L). This is the point of maximum force that can be recorded in grams. Depending on the filament diameters (pre-set) and lengths (user-determined) a full range of forces and pressures can be generated.</li>
	<li>Convert the mass recorded in step 1.6 to force in millinewton (mN) using the formula f = ma (Force is equal to mass multiplied by gravitational acceleration). f: force; m: mass; a: gravitational acceleration (9.8 m/s2) (Figure 1M).</li>
	<li>Finally, convert the calculated force to pressure (force/area) in kilopascal (kPa) by dividing the measured force by the surface area of the filament tip (Figure 1M). To calculate the area, convert the diameter of the different nitinol filaments from inches (0.04&#8221;, 0.06&#8221;, and 0.08&#8221;) to centimeters. Then, &#960;r2 (where, r = the nitinol filament radius) determines the area (see Figure 1M). Preparing multiple probes using filaments of different diameters and lengths will generate a full set spanning the responsive range for Drosophila larvae (sample set shown in Figure 1N). 
	NOTE: Check each mechanical probe at least every 3&#8211;4 weeks. When the pressure deviates by more than &#177; 3% from the original measure, a new mechanical probe must be fabricated.</li>
</ol>
2. Preparation of larvae
<ol>
	<li>Raise control strain (w1118) larval progeny or larvae containing the transgenes ppk-Gal4&#62;UAS-mCD8-GFP (for visualizing damage to sensory neurons) on standard food in a 25 &#176;C incubator. Typically, stocks are routinely maintained at 18 &#176;C but both parents and larval offspring are reared at 25 &#176;C on standard cornmeal food for experiments. 
	NOTE: Adult flies (five males and ten females, 1:2 ratio) are kept in the fly vials, to allow egg laying, for about 24 h. The time after egg laying (AEL) begins from when the adults are removed.</li>
	<li>Collect the third instar larvae, after approximately 96 h of egg laying, by gently squirting tap water into the soft fly food containing the larvae. Wandering larvae that have left the food, or which have everted anterior or posterior spiracles, are too large/old for this assay. Second instar larvae (~ less than 4 mm in length) are too small.</li>
	<li>Pour out the contents of the soft fly food into a clean standard size Petri dish (100 mm x 15 mm).</li>
	<li>Using forceps, sort mid third instar, medium sized, larvae (see Figure 2A) from smaller (second instar and early third instar) or larger (late or wandering third instar) larvae. Gentle manipulation with forceps to avoid any tissue damage to the larvae is recommended. 
	NOTE: The transfer using forceps is based mostly on water tension and not by applying pressure to the larvae with the blades of the forceps. An alternative to the use of forceps for maneuvering larvae is soft paint brushes. With either tool, the user should practice transferring the animals, so as not to cause unintended tissue damage that could complicate behavioral measurements.</li>
	<li>Transfer the mid third instar larvae, using forceps, into a small Petri dish (30 mm x 15 mm) containing a small plug of fly food moistened with water at room temperature. Keep the larvae in the small Petri dish until the experiments are performed, but not longer than 20 min. 
	NOTE: Generally, transferring 20&#8211;30 larvae to the food plug will give an adequate number for 20 min of behavioral assays.</li>
</ol>
3. Mechanical nociception assay
<ol>
	<li>Place a mid-third instar larva (using forceps) onto a thin pad of black or dark vinyl under a bright field stereomicroscope. The dark color provides contrast that improves visualization of the larva. It is preferable to have a freely movable piece of dark vinyl because it allows the user to align the larva without touching or hurting it.</li>
	<li>Put the optical fiber lights between the microscope objective lenses and the black or dark vinyl pad; this will allow adequate high contrast illumination for seeing the larva.</li>
	<li>Discard larvae that do not exhibit normal locomotion following transfer to the pad. These can interfere with the normal nociceptive behavioral response. For normal locomotion, see Video 1.</li>
	<li>Wipe away, using a paper towel, any excess water surrounding the larva that might cause the larva to float on the vinyl pad.</li>
	<li>Orient the larva by moving the dark vinyl pad. The head/mouth of the larva should point to the left if you are right-handed and vice versa if you are left-handed (Figure 2A&#8211;B).</li>
	<li>Apply the chosen mechanical probe, typically for 1&#8211;2 s, onto the posterior dorsal side of the larva at approximately abdominal segment A8 (see Figure 2B), until the probe bends and elicits the previously measured amount of pressure (Figure 2C). It is important that the probe presses against the dorsal surface of the larva and compresses the larvae into the underlying pad at the point of probe contact. 
	NOTE: At the point of contact between the tip of the nitinol filament and the dorsal cuticle-epidermis, probes lower than 2,300 kPa, mainly bend without penetrating the cuticle and underlying tissues. Such probes seldom affect larval mortality4. At higher pressures (&#62;5,000 kPa) the probes both bend and, occasionally, penetrate the cuticle and underlying tissues. Puncturing of the larvae impairs larval survival4 and, if observed, these larvae are typically discarded from behavioral analysis.</li>
	<li>Record the behavioral response for each larva. A positive nociceptive response (Video 2) is indicated if the larva shows a complete roll of 360&#730; along the axis of its body within 3 s. Other responses (attempting to turn, fast crawling, and wiggling) are considered negative for the purposes of this assay. 
	NOTE: Larvae stimulated with a subthreshold mechanical stimulus (200 kPa) did not elicit the typical nociceptive or rolling response (Video 3). Some larvae did exhibit fast forward or light touch responses such as changes in the direction of movement.</li>
	<li>Discard the larva and prepare the next one for assay, repeating steps 3.1 through 3.7.</li>
	<li>Repeat steps 3.1&#8211;3.7 until the desired number of larvae is reached (three to six sets of n = 10 larvae were used here for each probe). 
	NOTE: When using lower pressure mechanical probes (174&#8211;462 kPa), the assay will take more time per larva. This is because the tip of longer filaments oscillates more, making it harder to poke the larva in the center of the A8 segment. Practice is necessary with these probes.</li>
</ol>
4. Confocal microscopy to assess neuronal morphology
<ol>
	<li>Place a larva (of genotype ppk-Gal4&#62;UAS-mCD8-GFP to label sensory neurons) previously stimulated with a nitinol filament into an etherization chamber inside a Coplin jar containing a 10 mL beaker carrying a cotton ball soaked with ~1 mL of diethyl ether. Let the larva sit in the chamber for ~5 min. 
	NOTE: A detailed protocol for etherization is provided in a previous study published by our group12.</li>
	<li>Rinse the larva gently from the etherization chamber into a small Petri dish.</li>
	<li>Have ready one microscope slide, two small coverslips (22 x 22 mm), and one long coverslip (22 x 54 mm) (see Table of Materials).</li>
	<li>Add small drops of ether:oil solution (1:5 ratio of ethyl ether to halocarbon oil solution, see Table of Materials) to both ends of the slide, then place the small coverslips on top of the small droplets. This arrangement creates a small space gap where the larva can fit. 
	NOTE: Press the small coverslips against the microscope slide until it is difficult to slip.</li>
	<li>Add some drops of ether:oil solution on the middle of the microscope slide and then place the larva, using forceps, onto the center of the microscope slide (between the small coverslips). Make sure that the anteroposterior axis of the larva is parallel to the short side of the slide and that the dorsal side is facing up.</li>
	<li>Cover the larvae with the long coverslip placed on top of the larva and the two smaller coverslips. 
	NOTE: Generously press the long coverslip until the larva is almost flat.</li>
	<li>Image segment A8 of the larva using a confocal microscope (see Table of Materials) using laser wavelength 488 (GFP). 
	NOTE: Image the larva immediately because the anesthetization via ether will fade quickly (~5&#8211;10 min) and the larva will wake up and move, which will complicate further imaging.</li>
	<li>Capture Z-stack images at a resolution of 1024 x 1024 pixels using a 20x numerical aperture (NA) 0.7 dry objective lens at 1x zoom, step size of 1.5 &#956;m.</li>
</ol>
5. Quantitation of tissue damage
<ol>
	<li>Collect and convert the Z-series stack images, from section 4.8, into a single Z projection (a flattening of multiple images taken at different focal planes into a single composite image). This can be performed using commercially available software (e.g., Olympus Fluoview) or any equivalent open source platform, e.g., Fiji/Image J. Save the single Z projection in the TIFF format.</li>
	<li>Open the image analysis program Fiji/ImageJ.</li>
	<li>Click on File, from the menu bar, and select Open from the window that is displayed.</li>
	<li>Select the stored single image projection, saved in the TIFF format, to be analyzed.</li>
	<li>Click on Edit, from the menu bar, and select the Invert option from the window that is displayed.</li>
	<li>Click on the Image, from the menu bar, then select Adjust, from the window that is displayed, and finally select the Brightness/Contrast option.</li>
	<li>Select the Freehand Shape option from the tool bar to measure the area of the gap (if any).</li>
	<li>Click on Analyze, from the menu bar, and select the Measure option. This will display the area of the gap or wound.</li>
</ol>

<ol>
	<li>Tracey, W. D., Wilson, R. I., Laurent, G., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Painless,+a+Drosophila+gene+essential+for+nociception.">Painless, a Drosophila gene essential for nociception.</a> Cell. 113 (2), 261-273 (2003).</li><li>Zhong, L., Hwang, R. Y., Tracey, W. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pickpocket+is+a+DEG/ENaC+protein+required+for+mechanical+nociception+in+Drosophila+larvae.">Pickpocket is a DEG/ENaC protein required for mechanical nociception in Drosophila larvae.</a> Current Biology. 20 (5), 429-434 (2010).</li><li>Lopez-Bellido, R., Himmel, N. J., Gutstein, H. B., Cox, D. N., Galko, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+assay+for+chemical+nociception+in+Drosophila+larvae.">An assay for chemical nociception in Drosophila larvae.</a> Philosophical Transactions of the Royal Society of London. Series B: Biological Sciences. 374 (1785), 20190282 (2019).</li><li>Lopez-Bellido, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+factor+signaling+regulates+mechanical+nociception+in+flies+and+vertebrates.">Growth factor signaling regulates mechanical nociception in flies and vertebrates.</a> Journal of Neuroscience. 39 (30), 6012-6030 (2019).</li><li>Hu, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sensory+integration+and+neuromodulatory+feedback+facilitate+Drosophila+mechanonociceptive+behavior.">Sensory integration and neuromodulatory feedback facilitate Drosophila mechanonociceptive behavior.</a> Nature Neuroscience. 20 (8), 1085-1095 (2017).</li><li>Kim, S. E., Coste, B., Chadha, A., Cook, B., Patapoutian, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+Drosophila+Piezo+in+mechanical+nociception.">The role of Drosophila Piezo in mechanical nociception.</a> Nature. 483 (7388), 209-212 (2012).</li><li>Babcock, D. T., Landry, C., Galko, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytokine+signaling+mediates+UV-induced+nociceptive+sensitization+in+Drosophila+larvae.">Cytokine signaling mediates UV-induced nociceptive sensitization in Drosophila larvae.</a> Current Biology. 19 (10), 799-806 (2009).</li><li>Babcock, D. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hedgehog+signaling+regulates+nociceptive+sensitization.">Hedgehog signaling regulates nociceptive sensitization.</a> Current Biology. 21 (18), 1525-1533 (2011).</li><li>Turner, H. N., Patel, A. A., Cox, D. N., Galko, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Injury-induced+cold+sensitization+in+Drosophila+larvae+involves+behavioral+shifts+that+require+the+TRP+channel+Brv1.">Injury-induced cold sensitization in Drosophila larvae involves behavioral shifts that require the TRP channel Brv1.</a> PloS One. 13 (12), 0209577 (2018).</li><li>Im, S. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tachykinin+acts+upstream+of+autocrine+Hedgehog+signaling+during+nociceptive+sensitization+in+Drosophila.">Tachykinin acts upstream of autocrine Hedgehog signaling during nociceptive sensitization in Drosophila.</a> eLife. 4, 10735 (2015).</li><li>Cervero, F., Merskey, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+is+a+noxious+stimulus.">What is a noxious stimulus.</a> Pain Forum. 5 (3), 157-161 (1996).</li><li>Chattopadhyay, A., Gilstrap, A. V., Galko, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+and+global+methods+of+assessing+thermal+nociception+in+Drosophila+larvae.">Local and global methods of assessing thermal nociception in Drosophila larvae.</a> Journal of Visualized Experiments: JoVE. (63), e3837 (2012).</li><li>Follansbee, T. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+nociceptive+sensitization+requires+BMP+signaling+via+the+canonical+SMAD+pathway.">Drosophila nociceptive sensitization requires BMP signaling via the canonical SMAD pathway.</a> Journal of Neuroscience. 37 (35), 8524-8533 (2017).</li><li>Im, S. H., Patel, A. A., Cox, D. N., Galko, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+insulin+receptor+regulates+the+persistence+of+injury-induced+nociceptive+sensitization.">Drosophila insulin receptor regulates the persistence of injury-induced nociceptive sensitization.</a> Disease Models &amp; Mechanisms. 11 (5), (2018).</li><li>Jo, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+caspase+activity+is+required+independently+of+apoptosis+to+produce+active+TNF/Eiger+during+nociceptive+sensitization.">Drosophila caspase activity is required independently of apoptosis to produce active TNF/Eiger during nociceptive sensitization.</a> Cell Death &amp; Disease. 8 (5), 2786 (2017).</li><li>Hwang, R. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nociceptive+neurons+protect+Drosophila+larvae+from+parasitoid+wasps.">Nociceptive neurons protect Drosophila larvae from parasitoid wasps.</a> Current Biology. 17 (24), 2105-2116 (2007).</li><li>Guo, Y., Wang, Y., Wang, Q., Wang, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+PPK26+in+Drosophila+larval+mechanical+nociception.">The role of PPK26 in Drosophila larval mechanical nociception.</a> Cell Reports. 9 (4), 1183-1190 (2014).</li><li>Takagi, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Divergent+connectivity+of+homologous+command-like+neurons+mediates+segment-specific+touch+responses+in+Drosophila.">Divergent connectivity of homologous command-like neurons mediates segment-specific touch responses in Drosophila.</a> Neuron. 96 (6), 1373-1387 (2017).</li></ol>

The authors have nothing to disclose.

We modified an established mechanical assay1,2,16 using customized mechanical probes fabricated from nitinol filaments. This metal alloy allows us to use smaller diameter filaments that are appropriate to the size of the Drosophila larvae. Fishing line-based monofilaments have dominated the field of fly mechanical nociception to date2,5,6,16. Our nitinol filaments maintain their shape and measured pressure for approximately ~3&#8211;5 months (in our experience). By varying the length and diameter of the nitinol filaments, the user can generate a wide range of pressures spanning from subthreshold to a nearly complete rolling response. In particular, making subthreshold probes is simpler with the smaller diameter nitinol filaments. Using these probes, we found that pressure, rather than force, elicits more consistent nocifensive behavioral responses4. We demonstrate here, using a well-established UV-induced nociceptive sensitization model7,10,13, that these filaments are also a useful tool for studying mechanical hypersensitivity&#8212;allodynia and hyperalgesia.
Previous studies using mechanical probes fabricated from fishing line have led to a certain variability in behavioral responsiveness2,6,16,17. Several factors may account for this. First, because pressure is the important variable, the buffing of the filament tip so that it is rounded and does not have any sharp edges is critical. Second, reporting pressure values rather than only force is important for the reproducibility of the experiments, because different mechanical probes that generate similar forces can elicit disparate pressures4. Third, it is critical to apply only one mechanical stimulation per larva using noxious probes, because such probes produce a dose-dependent tissue damage at the epidermal4 and sensory neuronal levels (Figure 2E). A second or subsequent noxious mechanical stimulus, after tissue damage has been induced, could conceivably impair the function of the affected peripheral sensory neurons and elicit an altered behavioral response. In another study, larvae stimulated twice with noxious mechanical probes mostly displayed an enhanced behavioral response5, suggesting development of an acute mechanical sensitization (hyperalgesia), which might result from the tissue damage provoked by the first noxious mechanical stimulus. Conversely, other authors6 reported a mixed (increased or decreased) behavioral response, indicating that the altered behavioral response could be due to damage/dysfunction of the neuronal tissue. Stimulating each larva only once eliminates possible variance in behavioral responses resulting either from sensitization or tissue damage. Fourth, we mechanically stimulated segment A8, which is more posterior than previous studies (preferred areas A3&#8211;A4)2,5,16. Probes between ~3,900 kPa and 5,300 kPa applied to either segment A2 or A8 did not show any behavioral differences4. In addition, A8, compared to A2&#8211;A4, is easier to stimulate with mechanical probes that generate lower pressures (&#60;300 kPa) because the larva is thinner in this region and thus more easily compressed. Other studies showed that noxious mechanical stimulation of the posterior end of the larva (delivered by a rigid insect pin, held with forceps) mostly evoked forward locomotion, rather than an aversive or rolling response18. This different behavioral response could be due to differences in the properties of the used materials (bendable nitinol filament vs incompressible insect pin) or to different pressures delivered to the larvae (the pressure value of the insect pin was not reported).
The development of a mechanical nociception assay for Drosophila larvae has enabled the field to discover that different mechanical sensory ion channels and neural circuits mediate mechanical nociception5,6,16,17. However, the study of the mechanical hypersensitivity (allodynia and hyperalgesia) has lagged, compared to sensitization of the other sensory modalities&#8212;heat7,8,10,13,14, cold9, and chemical3. This lag may be due in part to the absence of suitable mechanical probes that can generate a full response range spanning subthreshold to suprathreshold pressures. Of particular importance, especially for assessing mechanical allodynia, are subthreshold probes that do not elicit an aversive rolling response from uninjured larvae. The significance of our improved mechanical probes is that they can be fabricated to span innocuous stimuli (subthreshold ~174 kPa&#8211;200 kPa) or the low to high noxious range (suprathreshold ~225 kPa to ~5,116 kPa). Here, we demonstrate using the nitinol von Frey-like filaments that Drosophila larvae develop both mechanical allodynia and mechanical hyperalgesia after UV irradiation. The mechanical sensitization shows some differences when compared to thermal sensitization. Both the onset and the peak of mechanical sensitization is earlier (~4 h) compared to thermal (heat) sensitization (~8 h for hyperalgesia and ~24 h for allodynia)7. In addition, the mechanical allodynia and hyperalgesia are concomitant (both peak at ~4 h). Furthermore, while heat sensitization (allodynia and hyperalgesia) resolves completely at later time points7, mechanical hypersensitivity exhibited a long tail that remained slightly above baseline. Cold sensitization in Drosophila involves a switch in cold-evoked behaviors9 and the emergence of new cold-evoked behaviors&#8212;a phenomenon that is not observed with mechanical stimulation. These differences in onset, duration, and observed behaviors suggest that each sensory modality may be controlled by different signaling pathways. Combining the sensitization assay described here with the powerful genetic tools available in Drosophila should allow a precise genetic dissection of the mechanical hypersensitivity (allodynia and hyperalgesia) observed.

Drosophila larvae exhibit a characteristic aversive rolling behavior when exposed to different noxious stimuli: thermal1, mechanical2, and chemical3. This behavior is clearly distinct from normal locomotion. Here we describe an improved mechanical assay that can be used to assess mechanical nociception and mechanical sensitization.
In a recent study, we fabricated von Frey-like filaments using nitinol wires4. Probes exerting different forces and pressures were made by varying the lengths and diameters of the nitinol wires forming each probe. Mechanical probes were calibrated and the measured force values (in millinewton, mN) were converted to pressure (kilopascal, kPa), based on the tip area of each probe4. Custom fabrication of mechanical probes allowed to us to generate subthreshold (&#8804;200 kPa) to suprathreshold (225 kPa to 5318 kPa) pressures, which could, in principle, be beneficial for studying mechanical hypersensitivity. Using these improved mechanical von Frey-like filaments, we showed that pressure4, as opposed to the previously examined force2,5,6 correlates more consistently with aversive behavioral responsiveness in Drosophila larvae. The improved mechanical assay described here also helped to identify a conserved Vascular Endothelial Growth Factor (VEGF)-related receptor tyrosine kinase signaling a pathway that regulates mechanical nociception in flies and rats4.
Mechanical allodynia and hyperalgesia, two modalities of hypersensitivity, are relatively understudied in Drosophila larvae, compared to the thermal (heat and cold) and chemical sensory modalities3,7,8,9,10. This is probably due to the lack of specific mechanical probes that span from innocuous stimuli to the high noxious range2,5,6. A normally innocuous stimulus that elicits the typical aversive rolling behavior after Drosophila larvae experience tissue damage3,7 is referred to as allodynia. An exaggerated rolling response to a typically noxious stimulus is known as hyperalgesia7. Noxious stimuli are defined as those that elicit tissue damage and can activate nociceptors11. Noxious stimuli delivered to Drosophila larvae damage either the barrier epidermis, the peripheral nociceptive sensory neurons3,4,7, or both.
In this article, we demonstrate how to custom fabricate and calibrate von Frey-like mechanical probes that are appropriate for Drosophila larvae. Further, we show how to use these probes to assay mechanical nociceptive responses in Drosophila larvae. Finally, we further demonstrate the utility of these probes by using them to demonstrate the presence of mechanical hypersensitivity, both allodynia and hyperalgesia, following tissue damage in Drosophila larvae (see Representative Results).

<table>
	<tbody>
		<tr>
			<td>Beaker</td>
			<td>Fisher Scientific</td>
			<td>02-540C</td>
			<td>Beaker of 10 ml of capacity. Any similar container will do.</td>
		</tr>
		<tr>
			<td>Black (Arkansas) bench stone</td>
			<td>Dan&#8217;s Whetstone</td>
			<td>SKU: I200306B24b-HQ-BAB-622-C</td>
			<td>Used to smoothe any irregularities of the nitinol wire tips. https://www.danswhetstone.com/product/special-extra-wide-black-bench-stone-6-x-2-1-2-x-1-2/</td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Olympus</td>
			<td>FV1000</td>
			<td>Any equivalent confocal microscope will do</td>
		</tr>
		<tr>
			<td>Coplin Jar</td>
			<td>Fisher Scientific</td>
			<td>08-816</td>
			<td>https://www.fishersci.com/shop/products/fisherbrand-glass-staining-dishes-10-slides-screw-cap/08816#?keyword=08-816</td>
		</tr>
		<tr>
			<td>Diethyl ether</td>
			<td>Fisher Scientific</td>
			<td>E138-500</td>
			<td>For anesthetizing larvae.</td>
		</tr>
		<tr>
			<td>Etherization chamber</td>
			<td></td>
			<td></td>
			<td>This is a homemade customized chamber. Please see details of its construction in our previous published paper12. The purpose of the etherization chamber is allow entry of diethyl ether fumes but prevent larval escape.</td>
		</tr>
		<tr>
			<td>Fiber Optic Light Guide</td>
			<td>Schott AG</td>
			<td>A08575</td>
			<td>Schott Dual Gooseneck 23 inch</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fine Science Tool</td>
			<td>FS-1670</td>
			<td>For transferring larvae</td>
		</tr>
		<tr>
			<td>Glue</td>
			<td>Aleene&#39;s</td>
			<td>N/A</td>
			<td>Aleene&#39;s&#174; Wood Glue, formerly called (Aleene&#39;s All-Purpose Wood Glue) 
			https://www.aleenes.com/aleenes-wood-glue</td>
		</tr>
		<tr>
			<td>Graspable holder</td>
			<td>Loew Cornell</td>
			<td>N/A</td>
			<td>Loew-Cornell Simply Art Wood Colored Craft Sticks, 500 pieces.</td>
		</tr>
		<tr>
			<td>Halocarbon oil 700</td>
			<td>Sigma</td>
			<td>H8898-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Hypodermic needle 30G 1/2&#34;L</td>
			<td>Fisher Scientific</td>
			<td>NC1471286</td>
			<td>BD Precisionglide&#174; syringe needles, gauge 30, L 1/2 inches. Used to make a hole into the wooden holder for the nitinol wires</td>
		</tr>
		<tr>
			<td>Large Petridish</td>
			<td>Falcon</td>
			<td>351007</td>
			<td>60 mm x 10 mm Polystyrene Petridish</td>
		</tr>
		<tr>
			<td>Microscope (Zeiss) Stemi 2000</td>
			<td>Carl Zeiss, Inc.</td>
			<td>NT55-605</td>
			<td>Any equivalent microscope will do</td>
		</tr>
		<tr>
			<td>Microscope Cover Glass 22x22</td>
			<td>Fisher</td>
			<td>12-545-B</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Cover Glass 22x40</td>
			<td>Corning</td>
			<td>2980-224</td>
			<td>Tickness 1 1/2</td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Globe Scientific Inc.</td>
			<td>1358Y</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini Diagonal Cutter</td>
			<td>Fisher Scientific</td>
			<td>S43981</td>
			<td>For cutting nitinol filaments</td>
		</tr>
		<tr>
			<td>Nitinol filaments, Diameters: 0.004&#8221;, 0.006&#8221;, 0.008&#8221;</td>
			<td>Mailin Co</td>
			<td>N/A</td>
			<td>Fifteen pieces of each diameter of 12&#8221; length were ordered. 
			https://malinco.com/</td>
		</tr>
		<tr>
			<td>Piece of black vinyl</td>
			<td>Office Depot</td>
			<td>N/A</td>
			<td>We use a small piece of vinyl cut from a binder. Dark color provides contrast. A small piece allows orientation of the larva</td>
		</tr>
		<tr>
			<td>Small Petridish</td>
			<td>Falcon</td>
			<td>351008</td>
			<td>35 mm x 10 mm Polystyrene Petridish</td>
		</tr>
		<tr>
			<td>Spatula</td>
			<td>Fisher Scientific</td>
			<td>21-401-10</td>
			<td>Double-Ended Micro-Tapered Stainless Steel Spatula. Used to place the food in the petri dish</td>
		</tr>
		<tr>
			<td>Wipes</td>
			<td>Fisher Scientific</td>
			<td>06-666A</td>
			<td>Kimpes KMTECH, Science Brand. Used to dry larvae of excess moisture.</td>
		</tr>
		<tr>
			<td>W1118</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>3605</td>
			<td>Control strain for behavioral assays</td>
		</tr>
		<tr>
			<td>ppk-Gal4&#62;UAS-mCD8-GFP</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>8749</td>
			<td>Strain for fluorescent labeling of class IV md neurons</td>
		</tr>
	</tbody>
</table>

an improved assay and tools for measuring mechanical nociception in drosophila larvae

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila larvae, customized metal nickel-titanium alloy (nitinol) filaments. These mechanical probes are similar to the von Frey filaments used in vertebrates to measure mechanical nociception. Here, we demonstrate how to make and calibrate these mechanical probes and how to generate a full behavioral dose-response from subthreshold (innocuous or non-noxious range) to suprathreshold (low to high noxious range) stimuli. To demonstrate the utility of the probes, we investigated tissue damage-induced hypersensitivity in Drosophila larvae. Mechanical allodynia (hypersensitivity to a normally innocuous mechanical stimulus) and hyperalgesia (exaggerated responsiveness to a noxious mechanical stimulus) have not yet been established in Drosophila larvae. Using mechanical probes that are normally innocuous or probes that typically elicit an aversive behavior, we found that Drosophila larvae develop mechanical hypersensitization (both allodynia and hyperalgesia) after tissue damage. Thus, the mechanical probes and assay that we illustrate here will likely be important tools to dissect the fundamental molecular/genetic mechanisms of mechanical hypersensitivity.

We developed customized mechanical probes, using nitinol filaments (Figure 1A,N), to elicit mechanically-evoked behaviors and generated a full behavioral dose response curve using both innocuous and noxious mechanical probes of varying intensity (Figure 2D) demonstrating that these probes can be used to study baseline (in the absence of injury) mechanical nociception.
Our behavioral assay results determined that probes exerting pressures below 200 kPa (~1.57 mN) (Figure 1M), when applied to Drosophila larvae, do not provoke an aversive rolling response (Figure 2D and Video 3). As expected, these subthreshold or non-noxious mechanical probes (175 kPa or 200 kPa) did not elicit visible neuronal tissue damage (Figure 2E). Because they do not induce damage, such probes could be useful to assess mechanical allodynia (hypersensitivity to a normally non-noxious mechanical stimuli). Conversely, suprathreshold or noxious probes (from 462 kPa to 5,116 kPa), elicited an augmented behavioral response (Figure 2D) in a dose dependent manner&#8212;with the higher pressures eliciting stronger behavioral responses. As anticipated, suprathreshold mechanical pressure also induced dose-dependent tissue damage to the peripheral sensory neurons themselves (Figure 2E). The measured area of tissue damage (in &#181;m2 &#177; standard deviation) taken from four larvae for each group were: 2,051.03 &#177; 703.81 (462 kPa), 5,102.29 &#177; 1,004.67 (2,283 kPa), and 12,238.83 &#177; 3,724.11 (5,116 kPa). Thus, pressures greater than or equal to 462 kPa (~63 mN), which evoke an aversive rolling response (in 25% or more of the larvae) and cause visible neuronal tissue damage (Figure 2E), could be appropriate to study mechanical hyperalgesia (hypersensitivity to normally noxious mechanical stimuli). Nociceptive mechanical probes (&#8805;462 kPa) always induce tissue damage (n = 10, evaluated qualitatively) but do not always provoke an aversive rolling response.
To evaluate mechanical hypersensitivity (allodynia and hyperalgesia), we used a well-established Drosophila larval model of nociceptive sensitization that uses ultraviolet light (UV) irradiation to induce tissue damage7,12. This assay has helped to dissect the genetic and cellular mechanisms of thermal nociceptive hypersensitivity8,9,10,13,14,15. To determine whether UV treatment causes mechanical allodynia, mid third-instar control (w1118) larvae were mock-irradiated or UV-irradiated (15&#8211;20 mJ/cm2) (Figure 3A). Then, the larvae were tested behaviorally at 2 h, 4 h, 8 h, 16 h, and 24 h post-treatment with a normally subthreshold mechanical probe (200 kPa, 1.57 mN). Approximately 20% of larvae responded as early as 2 h after UV treatment while 50% responded at 4 h, compared to 6.6% and 8.3% mock UV-irradiated animals, respectively (Figure 3B). This indicates that UV-induced tissue damage causes mechanical allodynia at 4 h post-irradiation. At later time points (8 h, 16 h, and 24 h) the behavioral response of the UV-treated larvae was in the range of 16%&#8211;20% responders (average mean of n = 3&#8211;6 sets of 10 larvae each), slightly increased (but not statistically significant) compared to the mock-irradiated control group (in the range of 3%&#8211;6% of responders, average mean of n = 3&#8211;6 sets of 10 larvae each) (Figure 3B).
To investigate mechanical hyperalgesia, a suprathreshold pressure (462 kPa, 3.63 mN), that normally induces an aversive rolling response in ~20% of larvae (Figure 2D) and causes neuronal tissue damage (Figure 2E), was used. We applied the 462 kPa probe onto the dorsal side of larvae with or without UV-induced tissue damage (Figure 3A). We found that larvae probed at 4 h, 8 h, and 16 h following UV treatment showed a significant increase in the aversive rolling response, with 4 h being the peak of the behavioral hypersensitivity (~60% responsive); mock UV-irradiated animals showed an ~27% of aversive response (Figure 3C). Similar to mechanical allodynia, the behavioral response at 8 h, 16 h, and 24 h of UV-treated animals (in the range of 36%&#8211;42%) was statistically indistinguishable from the non-treated larvae (in the range of 20%&#8211;26%). Larvae at the late third instar stage did show a slight decrease of the baseline behavioral response when compared with the middle third instar stage. We hypothesize this could be either by the increased size of the larvae (Figure 2A) or the increased thickness of the cuticle covering the body. This fact could explain why at a later stage of development the UV treatment does not induce greater mechanical sensitization, as observed 4 h post UV treatment.
Taken together, our results indicate that Drosophila larvae develop both mechanical allodynia and mechanical hyperalgesia following UV-induced tissue damage. The peak time of mechanical allodynia and hyperalgesia is the same, 4 h after UV treatment; however, mechanical hyperalgesia has a more pronounced temporal tail as it returns to baseline more slowly compared to mechanical allodynia.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61911/61911fig01.jpg" /> 
Figure 1: Development of a Von Frey-like tool to evaluate mechanical nociception in Drosophila larvae. (A) Picture of a mechanical probe used to study mechanical nociception in Drosophila larvae. (B) Nitinol filaments and their relative diameters are shown to relative scale. (C) Picture of the diagonal wire cutter used to cut the nitinol filaments. (D) Smoothing the sharp edges of the cut nitinol filament with a sharpening stone. (E) Hypodermic needle used to make a hole into the wooden popsicle stick handle of the probe. The tip of the needle needs to reach at least half the height of the handle stick for secure filament insertion. (F&#8211;G) Attachment of the nitinol filament by gluing into a wooden popsicle stick handle with insertion hole. (H&#8211;L) Calibration of mechanical probes by pressing them against a scale. (M) Values of force (in mN) and pressure (in kPa) generated by different mechanical probes. The length of each nitinol filament used to construct the probes (P1&#8211;P10; P: probe) is detailed in centimeters (cm). (N) A picture of a complete set of mechanical probes, ranging from 174 kPa to 5,116 kPa. <a href="https://www.jove.com/files/ftp_upload/61911/61911fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61911/61911fig02v2.jpg" /> 
Figure 2: Mechanical nociception assay: Von Frey-like filaments generate a dose-response curve of aversive rolling behavior and cause tissue damage to sensory neurons. (A) Pictures of the different stages (second and third instar) of Drosophila larvae. Scale bar: 2 mm. (B) Cartoon of the dorsal view of the third instar Drosophila larvae. The red dot indicates the abdominal segment where the mechanical probe is applied. T: thoracic segment; A: abdominal segment. Other anatomical landmarks are labeled. (C) Cartoon of the assay: A mechanical probe is applied to the dorsal side of the larva until it bends against the surface below and is then held for 2 s. If the pressure is sufficiently high, this elicits an aversive rolling response upon release. (D) Behavioral dose response; each blue dot represents the percent of larvae that responded, with aversive rolling, to the mechanical stimulation within a set of 10 animals. Violin plot of the percent of aversive rolling behavior induced by different mechanical probes. kPa: kilopascals. Box plots represent median (green), whiskers (red) represent the 10th and 90th percentiles. (E) Tissue damage: Third instar larvae (of genotype ppk-Gal4&#62;UAS-mCD8-GFP&#160;to label nociceptive sensory neurons) were probed at dorsal segment A8 with the indicated pressures. Fluorescently labeled paired ddaC class IV&#160;sensory neurons (across the dorsal midline) were then examined (see sections 4 and 5). White areas (red asterisks) represent gaps or tissue damage. Scale bar: 100 &#956;m. In panel B, the larva is shown in the dorsal view, while in C it is the lateral view. Mechanical probes pressed against the dorsal cuticle-epidermis side of the larva produce a depression like-pocket at the point of contact of the tip of the probe and the surrounding areas. The solid black line curved toward the ventral side is the top of the pocket, while the dashed gray lateral line represents the lateral side and the bottom of the pocket. <a href="https://www.jove.com/files/ftp_upload/61911/61911fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61911/61911fig03v2.jpg" /> 
Figure 3: Mechanical hypersensitivity after UV damage. (A) Schematic of the experimental design to test sensitization. Mid third instar were mock treated (non-UV) or UV irradiated. The mechanical nociception assay was then performed at different time points (2 h, 4 h, 8 h, 16 h, and 24 h) following mock treatment or irradiation. (B) Mechanical allodynia: The percentage of larvae exhibiting aversive rolling after probing with a normally subthreshold or non-noxious mechanical stimulus (200 kPa, 1.57 mN) at the indicated time points after mock-treatment or UV irradiation. (C) Mechanical hyperalgesia: The percentage of larvae exhibiting aversive rolling after probing with a normally suprathreshold or noxious mechanical stimulus (462 kPa, 3.63 mN) at the indicated time points after mock-treatment or UV irradiation. Error bars indicate mean +/- SEM. Two-tailed unpaired t-test was used for statistical analysis: *p &#60; 0.05, **p &#60; 0.01; ns: not significant. Each red dot, in panels B and C, represents the mean proportion of 10 larvae, n = 3&#8211;6 sets per timepoint/condition. <a href="https://www.jove.com/files/ftp_upload/61911/61911fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Normal locomotion of Drosophila larvae. <a href="https://cloudflare.jove.com/files/ftp_upload/61911/Video_1_regresolu.mp4" target="_blank">Please click here to download this video.</a>
Video 2: Noxious mechanical stimulation of Drosophila larvae. <a href="https://cloudflare.jove.com/files/ftp_upload/61911/Video_2_regresol.mp4" target="_blank">Please click here to download this video.</a>
Video 3: Subthreshold mechanical stimulation of Drosophila larvae. <a href="https://cloudflare.jove.com/files/ftp_upload/61911/Video_3_Subthreshold_short.mp4" target="_blank">Please click here to download this video.</a>

Watch this Scientific Journal Video about An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae at JoVE.com

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

The goal of this protocol is to show how to perform an improved assay for mechanical nociception in Drosophila larvae. We use the assay here to demonstrate that mechanical hypersensitivity (allodynia and hyperalgesia) exists in Drosophila larvae.

An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

Published assays for mechanical nociception in Drosophila have led to variable assessments of behavior. Here, we fabricated, for use with Drosophila ...

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4.7K Views.  The University of Texas MD Anderson Cancer Center. The goal of this protocol is to show how to perform an improved assay for mechanical nociception in Drosophila larvae. We use the assay here to demonstrate that mechanical hypersensitivity (allodynia and hyperalgesia) exists in Drosophila larvae.

Video: An Improved Assay and Tools for Measuring Mechanical Nociception in Drosophila Larvae

The Modified Hole Board - Measuring Behavior, Cognition and Social Interaction in Mice and Rats

This protocol describes the modified hole board (mHB), which combines features from a traditional hole board and open field and is designed to measure multiple dimensions of unconditioned behavior in small laboratory mammals (e.g., mice, rats, tree shrews and small primates). This paradigm is a valuable alternative for the use of a behavioral test battery, since a broad behavioral spectrum of an animal&#8217;s behavioral profile can be investigated in one single test.
The apparatus consists of a box, representing the &#8216;protected&#8217; area, separated from a group compartment. A board, on which small cylinders are staggered in three lines, is placed in the center of the box, representing the &#8216;unprotected&#8217; area of the set-up. The cognitive abilities of the animals can be measured by baiting some cylinders on the board and measuring the working and reference memory. Other unconditioned behavior, such as activity-related-, anxiety-related- and social behavior, can be observed using this paradigm. Behavioral flexibility and the ability to habituate to a novel environment can additionally be observed by subjecting the animals to multiple trials in the mHB, revealing insight into the animals&#8217; adaptive capacities.
Due to testing order effects in a behavioral test battery, na&#239;ve animals should be used for each individual experiment. By testing multiple behavioral dimensions in a single paradigm and thereby circumventing this issue, the number of experimental animals used is reduced. Furthermore, by avoiding social isolation during testing and without the need to food deprive the animals, the mHB represents a behavioral test system, inducing if any, very low amount of stress.

This protocol describes the modified hole board, which is a behavioral test set-up that comprises the characteristics of an open field and a traditional hole board. This set-up enables the differential analysis of unconditioned behavior of small laboratory mammals as well as the analysis of cognitive abilities.

This protocol describes the modified hole board (mHB), which combines features from a traditional hole board and open field and is designed to measure ...

An Innovative Running Wheel-based Mechanism for Improved Rat Training Performance

This study presents an animal mobility system, equipped with a positioning running wheel (PRW), as a way to quantify the efficacy of an exercise activity for reducing the severity of the effects of the stroke&#160;in rats. This system provides more effective animal exercise training than commercially available systems such as treadmills and motorized running wheels (MRWs). In contrast to an MRW that can only achieve speeds below 20 m/min, rats are permitted to run at a stable speed of 30 m/min on a more spacious and high-density rubber running track supported by a 15 cm wide acrylic wheel with a diameter of 55 cm in this work. Using a predefined adaptive acceleration curve, the system not only reduces the operator error but also trains the rats to run persistently until a specified intensity is reached. As a way to evaluate the exercise effectiveness, real-time position of a rat is detected by four pairs of infrared sensors deployed on the running wheel. Once an adaptive acceleration curve is initiated using a microcontroller, the data obtained by the infrared sensors are automatically recorded and analyzed in a computer. For comparison purposes, 3 week training is conducted on rats using a treadmill, an MRW and a PRW. After surgically inducing middle cerebral artery occlusion (MCAo), modified neurological severity scores (mNSS) and an inclined plane test were conducted to assess the neurological damages to the rats. PRW is experimentally validated as the most effective among such animal mobility systems. Furthermore, an exercise effectiveness measure, based on rat position analysis, showed that there is a high negative correlation between the effective exercise and the infarct volume, and can be employed to quantify a rat training in any type of brain damage reduction&#160;experiments.

This study presents an innovative running wheel-based animal mobility system to quantify an effective exercise activity in rats. A rat-friendly testbed is built, using a predefined adaptive acceleration curve, and a high correlation between the effective exercise rate and the infarct volume suggests the protocol&#39;s potential for stroke prevention experiments.

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An Automated Rapid Iterative Negative Geotaxis Assay for Analyzing Adult Climbing Behavior in a Drosophila Model of Neurodegeneration

Neurodegenerative diseases are frequently associated with a progressive loss of movement ability, reduced life span, and age-dependent neurodegeneration. To understand the mechanism of these cellular events, and their causal relationships with each other, Drosophila melanogaster, with its sophisticated genetic tools and diverse behavioral features, are used as disease models for assessing neurodegenerative phenotypes. Here we describe a high-throughput method to analyze Drosophila adult negative geotaxis behavior, as an indication for possible motor defects associated with neurodegeneration. An automated machine is designed and developed to drive fly synchronization using an initial electric impulse, later allowing the recording of negative geotaxis behavior over a course of secs to mins. Images from the digitally recorded video are then processed with the self-designed RflyDetection software for statistical data manipulation. Different from the manually controlled negative geotaxis assay based on single fly, this precise, fast, and high-throughput protocol allows data acquisition from more than hundreds of flies simultaneously, providing an efficient approach to advance our understanding in the underlying mechanism of locomotor deficits associated with neurodegeneration.

An Automated Rapid Iterative Negative Geotaxis Assay for Analyzing Adult Climbing Behavior in a Drosophila Model of Neurodegeneration

This step-by-step protocol analyzes Drosophila negative geotaxis behavior using an automated multi-cylinder system that hosts hundreds of flies and synchronizes their action by an electric motor. Upon synchronization, fly negative geotaxis behavior is assayed, digitally recorded, and analyzed using the self-designed RflyDetection software.

Neurodegenerative diseases are frequently associated with a progressive loss of movement ability, reduced life span, and age-dependent neurodegeneration. ...

A Simple and Low-cost Assay for Measuring Ambulation in Mouse Models of Muscular Dystrophy

Measuring functional outcomes in the treatment of muscular dystrophy is an essential aspect of preclinical testing. The assessment of voluntary ambulation in mouse models is a non-invasive and reproducible activity assay that is directly analogous to measures of patient ambulation such as the 6-minute walk test and related mobility scores. Many common methods for testing mouse ambulation speed and distance are based on the open field test, where an animal&#39;s free movement within an arena is measured over time. One major downside to this approach is that commercial software and equipment for high-resolution motion tracking is expensive and may require transferring mice to specialized facilities for testing. Here, we describe a low-cost, video-based system for measuring mouse ambulation that utilizes free and open-source software. Using this protocol, we demonstrate that voluntary ambulation in the dystrophin-null mdx mouse model for Duchenne muscular dystrophy (DMD) is decreased relative to wild-type mouse activity. In mdx mice expressing the utrophin transgene, these activity deficits are not observed and the total distance traveled is indistinguishable from wild-type mice. This method is effective for measuring changes in voluntary ambulation associated with dystrophic pathology, and provides a versatile platform that can be readily adapted to diverse research settings.

This protocol describes a flexible, low-cost system for measuring mouse ambulation in an open field activity assay. We show that a 6-minute ambulation assay based on this system detects a decrease in voluntary movement in mdx mice, and accurately distinguishes improvement in a muscle-specific rescue of these animals.

Measuring functional outcomes in the treatment of muscular dystrophy is an essential aspect of preclinical testing. The assessment of voluntary ambulation ...

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen distribution. It is also possible that aberrant behavior of oxygen-sensing neurons could induce hypoxia-like behavior in the presence of normal oxygen levels. In D. melanogaster, development at low oxygen levels results in inhibition of growth and sluggish behavior during the larval phases. However, these established manifestations of oxygen deficit overlap considerably with the phenotypes of many mutations that regulate growth, stress responses or locomotion. As result, there is currently no assay available to identify i) cellular hypoxia induced by a mutation or ii) hypoxia-like behavior when induced by abnormal neuronal behavior.
We have recently identified two distinctive behaviors in D. melanogaster larvae that occur at normal oxygen levels in response to internal detection of hypoxia. First, at all stages, such larvae avoid burrowing into food, often straying far away from a food source. Second, tunneling into a soft substratum, which normally occurs during the wandering third instar stage is completely abolished if larvae are hypoxic. The assay described here is designed to detect and quantitate these behaviors and thus to provide a way to detect hypoxia induced by internal damage rather than low external oxygen. Assay plates with an agar substratum and a central plug of yeast paste are used to support animals through larval life. The positions and state of the larvae are tracked daily as they proceed from first to third instar. The extent of tunneling into the agar substratum during wandering phase is quantitated after pupation using NIH ImageJ. The assay will be of value in determining when hypoxia is a component of a mutant phenotype and thus provide insight into possible sites of action of the gene in question.

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

The protocol describes a simple assay to identify Drosophila melanogaster larvae that are experiencing hypoxia under normal atmospheric oxygen levels. This protocol allows hypoxic larvae to be distinguished from other mutants that show overlapping phenotypes such as sluggishness or slow growth.

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen ...

Measuring Active and Passive Tameness Separately in Mice

Domesticated animals such as dogs and laboratory mice show a high level of tameness, which is important for humans to handle them easily. Tameness has two behavioral components: a reluctance to avoid humans (passive tameness) and a motivation to approach humans (active tameness). To quantify these components in mice, we previously developed behavioral tests for active tameness, passive tameness, and the willingness to stay on a human hand, each designed to be completed within 3 min. The data obtained were used for selective breeding, with a large number of mice analyzed per generation. The active tameness test measures the movement of the mouse toward a human hand and the contact it engages in. The passive tameness test measures the duration of time that a mouse tolerates human touch. In the stay-on-hand test, a mouse is placed on a human hand and touched slowly using the thumb of that hand; the duration of time that the animal remains on the hand is measured. Here, we describe the test set-up and apparatus, explain the procedures, and discuss the appropriate data analysis. Finally, we explain how to interpret the results.

Tameness in animals includes a reduction of avoidance responses to humans (passive tameness) and an increase in active approaches to humans (active tameness). Here, we describe detailed protocols for three behavioral tests (active tameness, passive tameness, and stay-on-hand tests) to separately measure the active and passive tameness of mice.

Domesticated animals such as dogs and laboratory mice show a high level of tameness, which is important for humans to handle them easily. Tameness has two ...

An Automated Method to Determine the Performance of Drosophila in Response to Temperature Changes in Space and Time

Temperature is a ubiquitous environmental factor that affects how species distribute and behave. Different species of Drosophila fruit flies have specific responses to changing temperatures according to their physiological tolerance and adaptability. Drosophila flies also possess a temperature sensing system that has become fundamental to understanding the neural basis of temperature processing in ectotherms. We present here a temperature-controlled arena that permits fast and precise temperature changes with temporal and spatial control to explore the response of individual flies to changing temperatures. Individual flies are placed in the arena and exposed to pre-programmed temperature challenges, such as uniform gradual increases in temperature to determine reaction norms or spatially distributed temperatures at the same time to determine preferences. Individuals are automatically tracked, allowing the quantification of speed or location preference. This method can be used to rapidly quantify the response over a large range of temperatures to determine temperature performance curves in Drosophila or other insects of similar size. In addition, it can be used for genetic studies to quantify temperature preferences and reactions of mutants or wild-type flies. This method can help uncover the basis of thermal speciation and adaptation, as well as the neural mechanisms behind temperature processing.

An Automated Method to Determine the Performance of Drosophila in Response to Temperature Changes in Space and Time

Here we present a protocol to automatically determine the locomotor performance of Drosophila at changing temperatures using a programmable temperature-controlled arena that produces fast and accurate temperature changes in time and space.

Temperature is a ubiquitous environmental factor that affects how species distribute and behave. Different species of Drosophila fruit flies have specific ...

Measuring the Functional Abilities of Children Aged 3-6 Years Old with Observational Methods and Computer Tools

The analysis of functional abilities and their development in early childhood (0-6 years old) are fundamental aspects among young children with certain types of developmental difficulties that can facilitate prevention, through programmed interventions adapted to the needs of each user (student or patient). There are, however, few investigations to date, that have analyzed the use of automated tools for recording and interpreting the results of the initial assessment. Here, a protocol is presented to examine the functional abilities in early childhood in young children, aged between 3-6 years old, with intellectual disabilities, but the protocol can also be used for ages 0 to 6 years. The protocol makes use of a computer application, eEarlyCare, that facilitates the interpretation of the results of systematic observations, which are recorded in natural environments by professionals trained in early intervention. The software can be used to analyze 11 functional areas (Food Autonomy, Personal Care and Hygiene, Dressing and Undressing Independently, Sphincter Control, Functional Mobility, Communication and Language, Daily Life Routines, Adaptive Behavior and Attention) and a total of 114 different behaviors. Its use facilitates the analysis of the observed abilities and greatly assists early intervention. Compared to other observational methods, it allows a more efficient use of personal and material resources. The use of the computer application facilitates the recording of the observation results, which helps with organization and reflection on the observations. The software displays the observation results on-screen compared to normal developmental parameters. This information can be referred to for decision-making about the most suitable intervention program for each user (student or patient). Likewise, clustering techniques are applied to analyze the relation between the type of intellectual disabilities and functional development identified with the software, a relation that is intended to serve as a guide for early-care professional intervention.

We present a protocol to use a computational tool to record and analyze the functional abilities of children aged 3-6 years old. The protocol facilitates the comparison of these abilities throughout their development and can be used to assess developmental difficulties.

The analysis of functional abilities and their development in early childhood (0-6 years old) are fundamental aspects among young children with certain ...

An Experimental Paradigm for Measuring the Effects of Ageing on Sentence Processing

Previous studies have found that older adults have greater difficulties in processing syntactically complex sentences than younger adults. However, the exact regions where the difficulties arise have not been fully identified. In this study, a maze task was implemented to investigate how older adults and younger adults differentially processed two types of sentences with different levels of syntactic complexity, namely subject relative clauses and object relative clauses. Participants were asked to choose between two alternatives at each segment of the sentences. The task required participants to engage in a strictly incremental mode of processing. The reading times for each segment were recorded, allowing the quantification of the difficulty of sentence reading. The task allowed us to identify the exact locations of the processing difficulty and thus facilitate a more accurate assessment of the age-related decline in sentence processing. The results indicate that the effect of ageing was found mainly at the head nouns, but not at other regions of the sentences, a finding which suggests that the maze task is an effective method to identify the exact location of the ageing effect on sentence processing. The implications of this experimental paradigm for investigating the effect of ageing on sentence processing are discussed.

Here, we present a protocol to examine the age-related decline in sentence processing using a maze task that allowed us to localize the processing difficulty at each word of the sentences during reading.

Previous studies have found that older adults have greater difficulties in processing syntactically complex sentences than younger adults. However, the ...

High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster

Drosophila melanogaster provides an excellent model to study the genetic underpinnings of alcohol sensitivity. In contrast to studies in human populations, the Drosophila model allows strict control over genetic background, and virtually unlimited numbers of individuals of the same genotype can be reared rapidly under well-controlled environmental conditions without regulatory restrictions and at relatively low cost. Flies exposed to ethanol undergo physiological and behavioral changes that resemble human alcohol intoxication, including loss of postural control, sedation, and development of tolerance. Here, we describe a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies. The assay is based on video recording of single flies introduced without anesthesia in 24-well cell culture plates in a set-up that enables synchronous initiation of alcohol exposure. The system enables a single person to collect individual ethanol sedation data on as many as 2,000 flies within an 8 h work period. The assay can, in principle, be extended to assess the effects of exposure to any volatile substance and applied to measure effects of acute toxicity of volatiles on other insects, including other fly species.

High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster

Current methods to measure alcohol sensitivity in Drosophila are designed to test groups of flies. We present a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies. The method does not require specialized tools and can be performed in any laboratory using common materials.