Name: constructing a low-budget laser axotomy system to study axon regeneration in c. elegans
Uploaded: 2011-11-15T12:00:00
Duration: 10 min 5 s
Description: Watch this Scientific Journal Video about Constructing a Low-budget Laser Axotomy System to Study Axon Regeneration in C. elegans at JoVE.com

This research was supported by the National Science Foundation, the McKnight Endowment Fund for Neuroscience, the Christopher and Dana Reeve Foundation and Amerisure Charitable Foundation.

<ol>
	<li>Hammarlund, M., Nix, P., Hauth, L., Jorgensen, E. M., Bastiani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axon+regeneration+requires+a+conserved+MAP+kinase+pathway.">Axon regeneration requires a conserved MAP kinase pathway.</a> Science. 323, 802-806 (2009).</li><li>Steinmeyer, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Construction+of+a+femtosecond+laser+microsurgery+system.">Construction of a femtosecond laser microsurgery system.</a> Nature protocols. 5, 395-407 (2010).</li><li>Wu, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Caenorhabditis+elegans+neuronal+regeneration+is+influenced+by+life+stage,+ephrin+signaling,+and+synaptic+branching.">Caenorhabditis elegans neuronal regeneration is influenced by life stage, ephrin signaling, and synaptic branching.</a> Proc Natl Acad. 104, 15132-15137 (2007).</li><li>Gilleland, C. L., Rohde, C. B., Zeng, F., Yanik, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidic+immobilization+of+physiologically+active+Caenorhabditis+elegans.">Microfluidic immobilization of physiologically active Caenorhabditis elegans.</a> Nature protocols. 5, 1888-1902 (2010).</li><li>Raabe, I., Vogel, S. K., Peychl, J., Tolic-Norrelykke, I. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+nanosurgery+and+cell+enucleation+using+a+picosecond+laser.">Intracellular nanosurgery and cell enucleation using a picosecond laser.</a> J. Microsc. 234, 1-8 (2009).</li><li>Hutson, M. S., Ma, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasma+and+cavitation+dynamics+during+pulsed+laser+microsurgery+in+vivo.">Plasma and cavitation dynamics during pulsed laser microsurgery in vivo.</a> Physical review letters. 99, 158104-158104 (2007).</li><li>Venugopalan, V., Guerra, A., Nahen, K., Vogel, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+laser-induced+plasma+formation+in+pulsed+cellular+microsurgery+and+micromanipulation.">Role of laser-induced plasma formation in pulsed cellular microsurgery and micromanipulation.</a> Physical review letters. 88, 078103-078103 (2002).</li><li>Bourgeois, F., Ben-Yakar, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Femtosecond+laser+nanoaxotomy+properties+and+their+effect+on+axonal+recovery+in+C.+elegans.">Femtosecond laser nanoaxotomy properties and their effect on axonal recovery in C. elegans.</a> Opt Express. 16, 5963-5963 (2008).</li><li>O'Brien, G. S., Rieger, S., Martin, S. M., Cavanaugh, A. M., Portera-Cailliau, C., Sagasti, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+axotomy+and+time-lapse+confocal+imaging+in+live+zebrafish+embryos.">Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos.</a> J. Vis. Exp. (24), e1129-e1129 (2009).</li><li>Tsai, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasma-mediated+ablation:+an+optical+tool+for+submicrometer+surgery+on+neuronal+and+vascular+systems.">Plasma-mediated ablation: an optical tool for submicrometer surgery on neuronal and vascular systems.</a> Curr. Opin. Biotechnol. 20, 90-99 (2009).</li><li>Chung, S. H., Clark, D. A., Gabel, C. V., Mazur, E., Samuel, A. D. <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+the+AFD+neuron+in+C.+elegans+thermotaxis+analyzed+using+femtosecond+laser+ablation.">The role of the AFD neuron in C. elegans thermotaxis analyzed using femtosecond laser ablation.</a> BMC Neurosci. 7, 30-30 (2006).</li><li>Shen, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ablation+of+cytoskeletal+filaments+and+mitochondria+in+live+cells+using+a+femtosecond+laser+nanoscissor.">Ablation of cytoskeletal filaments and mitochondria in live cells using a femtosecond laser nanoscissor.</a> Mech. Chem. Biosyst. 2, 17-25 (2005).</li><li>Rao, G. N., Kulkarni, S. S., Koushika, S. P., Rau, K. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+nanosecond+laser+axotomy:+cavitation+dynamics+and+vesicle+transport.">In vivo nanosecond laser axotomy: cavitation dynamics and vesicle transport.</a> Opt. Express. 16, 9884-9894 (2008).</li><li>Rohde, C. B., Yanik, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subcellular+in+vivo+time-lapse+imaging+and+optical+manipulation+of+Caenorhabditis+elegans+in+standard+multiwell+plates.">Subcellular in vivo time-lapse imaging and optical manipulation of Caenorhabditis elegans in standard multiwell plates.</a> Nat. Commun. 2, 271-271 (2011).</li></ol>

No conflicts of interest declared.

There are several good discussions of laser microsurgery with different laser systems3,5-11. Femtosecond IR lasers are the "gold standard" for subcellular laser ablation12 and convenient if associated with an imaging facility, but they are often too costly for individual users. If you require a femtosecond IR laser for imaging your sample because of depth of imaging then you will probably need one for laser axotomy. Transparent tissues with target axons within 30-50 um of the surface are probably feasible with blue and green pulse lasers in the 20 uJ/pulse range targeted through a high na immersion lens. There will be more collateral damage with the nano and pico second lasers compared with the femtosecond laser, especially as the depth of the target axon increases. C. elegans is transparent and all axons are within 20-30 um of the surface. We routinely cut motor axons that are within 5 um of the surface. We have also easily cut axons within the nerve ring that are about 20 um from the cuticle surface. We found the limit for cutting axons to be about 30-50 um through the diameter of an adult worm. It is likely that the laser ablation system described here would work well with many different preparations that fit the criteria of transparency and depth of target axons. Still, it is a bit surprising, given the theoretical advantages of the femtosecond IR lasers, that nano and picosecond 355 nm and 532 nm lasers perform so well for laser axotomy in C. elegans6,13. We see no differences in axon regeneration in response to laser axotomy with nanosecond 440 nm, nanosecond 532 nm, and femtosecond IR lasers.
Solid state 355 nm UV lasers are about the same cost as the 532 nm diode pumped passively Q-Switched solid state laser, but require either higher powers or optics that efficiently pass these shorter wavelengths. Most optics are designed to perform well with visible 400-700 nm light. 355 nm lasers would offer some advantages6 such as reduced plasma threshold, smaller spot size, and a long pass dichroic would efficiently direct 100% of the ablation laser to the target and allow simultaneous imaging of GFP without loss of sample signal. Blue 440 nm lasers would retain the advantages of working with a visible light laser (good performance with standard optics and safety). Unfortunately, the cost of a DPP Q-switched solid state 440 nm laser is 3 times the cost of a 355nm or 532 nm laser of similar power at this time. If we were designing a new system for C. elegans , where target depth is minimal, we would opt for a UV transparent lens (cost about $3,000 for a 40X 1.3na oil lens with 50-70% transmittance at 355nm) and a 355 nm laser producing 5-10 uJ/pulse at 1-20 kHz.
Axon ablation is thought to be due to plasma formation. Shorter pulses will produce plasma thresholds at lower power and the plasma volumes will be smaller6. The goal is to produce the smallest and shortest-lived plasma by adjusting the pulse energy to the lowest power. Larger long-lived plasmas will generate cavitation bubbles that damage surrounding cells. We have generally found that ablation using about 50-100 pulses at the highest frequency (i.e. 2.5kHz) gives the best results. This suggests that damage can be gradually integrated over a series of pulses to better control the extent of damage. The laser ablation system described here is very forgiving if the beam is aligned and expanded accurately. We start cutting axons in adult worms at 10% laser power (average of 0.3 mW measured at 2500 Hz and estimated 1 uJ/pulse) and can cut with limited localized damage through 14% power. You can observe larger areas of damage from 15-39% laser power, but only above 40% power (about 1 mW average measured at 2500 Hz) do the worms explode due to large cavitation bubbles and damage to the worm's cuticle.
The Steinmeyer et al. 20102 paper is an excellent resource for constructing a laser ablation system. Leon Avery also has a nice practical description of a laser ablation system based on an inexpensive N2 gas 337 nm laser and he provides some great practical advice (elegans.swmed.edu/Worm_labs/Avery/). When designing your own system you should first talk with your microscope representative to determine how to target the ablation laser to the microscope objective. Your microscope should be mounted on a vibration isolation table (Newport, TMI, Thor). A breadboard top is optimal, but a solid steel top can still be used with magnetic positioners. A dedicated intermediate module on an upright scope is nice because all optical elements can be left in place. However, it may be less expensive and more convenient to use a camera port (inverted) or a "combiner" attached to an epi-fluorescent port. You need to know the size of the back aperture and the transmittance (at the ablation laser wavelength) of the objective lens you intend to use. Once you decide on a laser (e.g., Crylas, TEEM, Crystal, or CRC) you should contact Thor or Newport for help with selecting the correct optical elements, hardware and safety equipment. We decided on a dual Galilean beam expander to save space, but a simpler Keplerian expander is an option (f2/f1=expansion factor). A custom beam expander will be the least expensive, but Newport, Thor, and several of the laser manufacturers offer excellent quality commercial beam expanders. Some laser manufacturers also offer manual and electronically controlled attenuators that may be cost effective and space saving, but not as versatile as the Glan-Thompson polarizer and half wave plate. You will also need to decide how to control the laser. You can design a LabView based controller, use a commercial TTL generator, or software provided by the laser company. Discuss the options with the specific laser manufacturer.
We have provided a list of the hardware we have used only as a general guide. The system described here cost about $15,000 when added to our existing imaging system. Your local Physics department will often have someone willing to provide a practical introduction to safely working with free beam lasers.
Alternative methods for immobilization and time lapse imaging of C. elegans have been recently described.14
Troubleshooting
The most difficult and critical step is alignment of the centered expanded beam to the optical axis of the microscope. Once you have the beam centered and expanded through the Galilean lenses you should work hard to get it aligned to within 5 um of the microscope's optical axis BEFORE using the steering mirrors for the final alignment to the center. Excessive use of the steering mirrors to align the beam to the microscope will move it off axis through the beam expander. USING EXTREME CAUTION you can attenuate the laser beam with the appropriate ND filter and directly view the laser beam profile on a reflective target (front surface mirror). You can gently nudge the microscope to precisely center the beam in the 60X objective field of view (if it can't be centered in the up/down range you need to adjust the rail height). The beam profile can be used to precisely align the microscope optical axis to give a centered beam that is circular and "flares" symmetrically. An asymmetric flare is an indication that the microscope axis is not perfectly aligned (or the rail is not level). Finally, you should adjust L3 and see an even and symmetrical expansion and contraction of the beam. Focus the microscope precisely on the target surface and then adjust L3 to focus the laser beam to the smallest spot. You should now find that the laser beam is within 5 um of the center when imaged via LSCM or CCD system and the ablation spot is within 1um of Z focus.
If you find you are "blowing up" worms or consistently generating large cavitation bubbles to cut axons your problem is most likely the fine alignment of the ablation laser. Make sure the minimum (&lt;500nm) ablation spot is localized to the Z focus (adjust convergence with L3 lens) and to the XY fiduciary spot (adjust with last kinematically mounted mirror).
Targeting axons closer to the surface will require less laser power. Similarly, smaller animals (L1 and L2) will require less laser power for axotomy compared to targeting the same axons in adults.
If your wild type axons do not regenerate or the axons bleach you should try reducing imaging laser power or decreasing the sampling rate. If your worms die or become sickly try reducing the percent Agarose in 1% steps. Make sure you are not moving the coverslip after mounting worms as this will often cause them to die when using high percentage agarose and microspheres.
If your worms burst or die during a time-lapse session it is due to: 1. Percentage agarose is too high. 2. Damage to cuticle by ablation laser. 3. Movement of coverslip after mounting. If damage to the cuticle is at fault it will only affect mounted worms that have been targeted with the ablation laser.
If your worm moves too much during a time-lapse session try increasing the percentage agarose in 0.5% steps. Healthy axons in healthy worms are always "active" and display a consistent level of fluorescence. If you see a sudden decrease in fluorescence or activity the worm is dying or dead. Multiple beaded or fragmented axons are a sure sign of a dying or dead worm.
If you have trouble keeping your axons in focus during the entire time lapse session there may be several different problems. First check your stage drift by imaging an inert sample on a glass slide over 10 hrs. A few microns drift may be due to thermal instability, but greater than 5 um is probably due to mechanical issues with your microscope. Problems with mounting are more common. Let your mounted worms equilibrate with your microscope stage for 30 minutes before starting your time lapse. Check that your Vaseline seal did not develop leaks during the course of your time lapse session.

<table border="1">
	<tbody>
		<tr>
			<th>
				Name of the reagent</th>
			<th>
				Company</th>
			<th>
				Catalogue number</th>
			<th>
				Comments (optional)</th>
		</tr>
		<tr>
			<td>535 nm Laser</td>
			<td>Crylas</td>
			<td>FDSS532Q3</td>
			<td>(TEEM, Crystal, and CRC also offer comparable lasers)</td>
		</tr>
		<tr>
			<td>All optics and hardware</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>(Thor offers comparable optics and hardware)</td>
		</tr>
		<tr>
			<td>SUPREMA Optical Mount, 1.0-in diameter</td>
			<td>Newport</td>
			<td>SS100-F2KN</td>
			<td>2</td>
		</tr>
		<tr>
			<td>1" diameter post, 1" height</td>
			<td>Newport</td>
			<td>PS-1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>1" diameter post fork</td>
			<td>Newport</td>
			<td>PS-F</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Achromatic Zero-Order Wavel Plate, &frac12; Wave Retardation, 400-700nm</td>
			<td>Newport</td>
			<td>10RP52-1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Rotation Stage, 1" Aperature</td>
			<td>Newport</td>
			<td>RSP-1T</td>
			<td>1</td>
		</tr>
		<tr>
			<td>1" diameter post, 1" height</td>
			<td>Newport</td>
			<td>PS-1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>1" diameter post fork</td>
			<td>Newport</td>
			<td>PS-F</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Glan-Laser Calcite Polarizer, 430-700nm</td>
			<td>Newport</td>
			<td>10GL08AR.14</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Polarizer Rotation Mount</td>
			<td>Newport</td>
			<td>RM25A</td>
			<td>1</td>
		</tr>
		<tr>
			<td>&frac14;" spacer for 1" diameter post</td>
			<td>Newport</td>
			<td>PS-0.25</td>
			<td>1</td>
		</tr>
		<tr>
			<td>&frac12;" height, 1"diameter post</td>
			<td>Newport</td>
			<td>PS-0.5</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Fork, 1" diameter post</td>
			<td>Newport</td>
			<td>PS-F</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Beam Dump</td>
			<td>Newport</td>
			<td>PL15</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Microscope Objective Lens Mount</td>
			<td>Newport</td>
			<td>LH-OBJ1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>1" diameter post, 1" height</td>
			<td>Newport</td>
			<td>PS-1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>1" diameter post fork</td>
			<td>Newport</td>
			<td>PS-F</td>
			<td>1</td>
		</tr>
		<tr>
			<td>High-Energy Nd:YAG Laser Mirror, 25.4 mm Diameter, 45&deg;, 532 nm</td>
			<td>Newport</td>
			<td>10Q20HE.2</td>
			<td>4</td>
		</tr>
		<tr>
			<td>SUPREMA Optical Mount, 1.0 inch diameter, clearance mounting hole</td>
			<td>Newport</td>
			<td>SN100C-F</td>
			<td>2</td>
		</tr>
		<tr>
			<td>High Precision Knob Adjustment Screw, 12.7mm travel, 100TPI</td>
			<td>Newport</td>
			<td>AJS100-0.5K</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Thread adaptor, &frac14;-20 male, 8-32 female</td>
			<td>Newport</td>
			<td>SS-1-B</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Rod Clamp for 1.5 inch diameter rod</td>
			<td>Newport</td>
			<td>340-RC</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Rod, 14 inch (35.5 cm) tall</td>
			<td>Newport</td>
			<td>40</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Rail carrier for X26, square 40mm length</td>
			<td>Newport</td>
			<td>CN26-40</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Steel Rail, 384mm (15") length</td>
			<td>Newport</td>
			<td>X26-384</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Rod Platform for 1.5 inch diameter rod</td>
			<td>Newport</td>
			<td>300-P</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Rod, 14 inch (35.5 cm) tall</td>
			<td>Newport</td>
			<td>40</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Plano-Concave Lens, 12.7mm diameter, -25mm EFL, 430-700nm</td>
			<td>Newport</td>
			<td>KPC025AR.14</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Plano-Convex Lens, 25.4mm diameter, 50.2mm EFL, 430-700nm</td>
			<td>Newport</td>
			<td>KPX082AR.14</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Plano-Convex Lens, 25.4mm diameter, 62.9mm EFL, 430-700nm</td>
			<td>Newport</td>
			<td>KPX085AR.14</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Fixed Lens Mount, 0.5" diameter, 1.0" Axis height</td>
			<td>Newport</td>
			<td>LH-0.5</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Fixed Lens Mount, 1.0" diameter, 1.0" Axis height</td>
			<td>Newport</td>
			<td>LH-1</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Microspheres 0.1um</td>
			<td>Polysciences</td>
			<td>00876</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>RPI</td>
			<td>A20090</td>
			<td>EEO matters</td>
		</tr>
		<tr>
			<td>Muscimol</td>
			<td>Sigma</td>
			<td>M1523</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

constructing a low-budget laser axotomy system to study axon regeneration in c. elegans

Laser axotomy followed by time-lapse microscopy is a sensitive assay for axon regeneration phenotypes in C. elegans1. The main difficulty of this assay is the perceived cost ($25-100K) and technical expertise required for implementing a laser ablation system2,3. However, solid-state pulse lasers of modest costs (&lt;$10K) can provide robust performance for laser ablation in transparent preparations where target axons are "close" to the tissue surface. Construction and alignment of a system can be accomplished in a day. The optical path provided by light from the focused condenser to the ablation laser provides a convenient alignment guide. An intermediate module with all optics removed can be dedicated to the ablation laser and assures that no optical elements need be moved during a laser ablation session. A dichroic in the intermediate module allows simultaneous imaging and laser ablation. Centering the laser beam to the outgoing beam from the focused microscope condenser lens guides the initial alignment of the system. A variety of lenses are used to condition and expand the laser beam to fill the back aperture of the chosen objective lens. Final alignment and testing is performed with a front surface mirrored glass slide target. Laser power is adjusted to give a minimum size ablation spot (&lt;1um). The ablation spot is centered with fine adjustments of the last kinematically mounted mirror to cross hairs fixed in the imaging window. Laser power for axotomy will be approximately 10X higher than needed for the minimum ablation spot on the target slide (this may vary with the target you use). Worms can be immobilized for laser axotomy and time-lapse imaging by mounting on agarose pads (or in microfluidic chambers4). Agarose pads are easily made with 10% agarose in balanced saline melted in a microwave. A drop of molten agarose is placed on a glass slide and flattened with another glass slide into a pad approximately 200 um thick (a single layer of time tape on adjacent slides is used as a spacer). A "Sharpie" cap is used to cut out a uniformed diameter circular pad of 13mm. Anesthetic (1ul Muscimol 20mM) and Microspheres (Chris Fang-Yen personal communication) (1ul 2.65% Polystyrene 0.1 um in water) are added to the center of the pad followed by 3-5 worms oriented so they are lying on their left sides. A glass coverslip is applied and then Vaseline is used to seal the coverslip and prevent evaporation of the sample.

Watch this Scientific Journal Video about Constructing a Low-budget Laser Axotomy System to Study Axon Regeneration in C. elegans at JoVE.com

Constructing a Low-budget Laser Axotomy System to Study Axon Regeneration in C. elegans

Laser axotomy followed by time-lapse imaging is a sensitive way to assay the effects of mutations in C. elegans on axon regeneration. A high quality, but inexpensive, laser ablation system can be easily added to most microscopes. Time lapse imaging over 15 hours requires careful immobilization of the worm.

Constructing a Low-budget Laser Axotomy System to Study Axon Regeneration in C. elegans

Laser axotomy followed by time-lapse microscopy is a sensitive assay for axon regeneration phenotypes in C. elegans1.  The main difficulty of this assay ...

constructing-low-budget-laser-axotomy-system-to-study-axon

Research

JoVE Journal

Bioengineering

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

An Experimental System to Study Mechanotransduction in Fetal Lung Cells

Mechanical forces generated in utero by repetitive breathing-like movements and by fluid distension are critical for normal lung development. A key component of lung development is the differentiation of alveolar type II epithelial cells, the major source of pulmonary surfactant. These cells also participate in fluid homeostasis in the alveolar lumen, host defense, and injury repair. In addition, distal lung parenchyma cells can be directly exposed to exaggerated stretch during mechanical ventilation after birth. However, the precise molecular and cellular mechanisms by which lung cells sense mechanical stimuli to influence lung development and to promote lung injury are not completely understood. Here, we provide a simple and high purity method to isolate type II cells and fibroblasts from rodent fetal lungs. Then, we describe an in vitro system, The Flexcell Strain Unit, to provide mechanical stimulation to fetal cells, simulating mechanical forces in fetal lung development or lung injury. This experimental system provides an excellent tool to investigate molecular and cellular mechanisms in fetal lung cells exposed to stretch. Using this approach, our laboratory has identified several receptors and signaling proteins that participate in mechanotransduction in fetal lung development and lung injury.

Mechanical forces play a key role in lung development and lung injury. Here, we describe a method to isolate rodent fetal lung type II epithelial cells and fibroblasts and to expose them to mechanical stimulation using an in vitro system.

Mechanical forces generated in utero by repetitive breathing-like movements and by fluid distension are critical for normal lung development. A key ...

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

Quantitative Locomotion Study of Freely Swimming Micro-organisms Using Laser Diffraction

Soil and aquatic microscopic organisms live and behave in a complex three-dimensional environment. Most studies of microscopic organism behavior, in contrast, have been conducted using microscope-based approaches, which limit the movement and behavior to a narrow, nearly two-dimensional focal field.1 We present a novel analytical approach that provides real-time analysis of freely swimming C. elegans in a cuvette without dependence on microscope-based equipment. This approach consists of tracking the temporal periodicity of diffraction patterns generated by directing laser light through the cuvette. We measure oscillation frequencies for freely swimming nematodes.
Analysis of the far-field diffraction patterns reveals clues about the waveforms of the nematodes. Diffraction is the process of light bending around an object. In this case light is diffracted by the organisms. The light waves interfere and can form a diffraction pattern. A far-field, or Fraunhofer, diffraction pattern is formed if the screen-to-object distance is much larger than the diffracting object. In this case, the diffraction pattern can be calculated (modeled) using a Fourier transform.2
C. elegans are free-living soil-dwelling nematodes that navigate in three dimensions. They move both on a solid matrix like soil or agar in a sinusoidal locomotory pattern called crawling and in liquid in a different pattern called swimming.3 The roles played by sensory information provided by mechanosensory, chemosensory, and thermosensory cells that govern plastic changes in locomotory patterns and switches in patterns are only beginning to be elucidated.4 We describe an optical approach to measuring nematode locomotion in three dimensions that does not require a microscope and will enable us to begin to explore the complexities of nematode locomotion under different conditions.

Microscopic organisms like the free-swimming nematode C. elegans, live and behave in a complex three-dimensional environment. We report on a novel approach that provides analysis of C. elegans using diffraction patterns. This approach consists of tracking the temporal periodicity of diffraction patterns generated by directing laser light through a cuvette.

Soil and aquatic microscopic organisms live and behave in a complex three-dimensional environment. Most studies of microscopic organism behavior, in ...

Measurement of Tension Release During Laser Induced Axon Lesion to Evaluate Axonal Adhesion to the Substrate at Piconewton and Millisecond Resolution

The formation of functional connections in a developing neuronal network is influenced by extrinsic cues. The neurite growth of developing neurons is subject to chemical and mechanical signals, and the mechanisms by which it senses and responds to mechanical signals are poorly understood. Elucidating the role of forces in cell maturation will enable the design of scaffolds that can promote cell adhesion and cytoskeletal coupling to the substrate, and therefore improve the capacity of different neuronal types to regenerate after injury.
Here, we describe a method to apply simultaneous force spectroscopy measurements during laser induced cell lesion. We measure tension release in the partially lesioned axon by simultaneous interferometric tracking of an optically trapped probe adhered to the membrane of the axon. Our experimental protocol detects the tension release with piconewton sensitivity, and the dynamic of the tension release at millisecond time resolution. Therefore, it offers a high-resolution method to study how the mechanical coupling between cells and substrates can be modulated by pharmacological treatment and/or by distinct mechanical properties of the substrate.

We measured the tension release in an axon that was partially lesioned with a laser dissector by simultaneous force spectroscopy measurement performed on an optically-trapped probe adhered to the membrane of the axon. The developed experimental protocol evaluates the axon adhesion to the culture substrate.

The formation of functional connections in a developing neuronal network is influenced by extrinsic cues. The neurite growth of developing neurons is ...

Planar Gradient Diffusion System to Investigate Chemotaxis in a 3D Collagen Matrix

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to their surrounding area, leading to cell movement and migration. In order to understand the mechanisms that can alter and/or affect cell migration, one can study these forces. In theory, understanding the fundamental mechanisms and forces underlying cell migration holds the promise of effective approaches for treating diseases and promoting cellular transplantation. Unfortunately, modern chemotaxis chambers that have been developed are usually restricted to two dimensions (2D) and have complex diffusion gradients that make the experiment difficult to interpret. To this end, we have developed, and describe in this paper, a direct-viewing chamber for chemotaxis studies, which allows one to overcome modern chemotaxis chamber obstacles able to measure cell forces and specific concentration within the chamber in a 3D environment to study cell 3D migration. More compelling, this approach allows one to successfully model diffusion through 3D collagen matrices and calculate the coefficient of diffusion of a chemoattractant through multiple different concentrations of collagen, while keeping the system simple and user friendly for traction force microscopy (TFM) and digital volume correlation (DVC) analysis.

Cell migration is an important part of human development and life. In order to understand the mechanisms that can alter cell migration, we present a planar gradient diffusion system to investigate chemotaxis in a 3D collagen matrix, which allows one to overcome modern diffusion chamber limitations of existing assays.

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to ...

Construction of Defined Human Engineered Cardiac Tissues to Study Mechanisms of Cardiac Cell Therapy

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that replicate the biofidelity of three-dimensional native human myocardium, while also enabling a controlled level of biological complexity, and allowing non-destructive longitudinal monitoring of tissue contractile function. Initially, human engineered cardiac tissues (hECT) were created using the entire cell population obtained from directed differentiation of human pluripotent stem cells, which typically yielded less than 50% cardiomyocytes. However, to create reliable predictive models of human myocardium, and to elucidate mechanisms of heterocellular interaction, it is essential to accurately control the biological composition in engineered tissues. 
To address this limitation, we utilize live cell sorting for the cardiac surface marker SIRP&#945; and the fibroblast marker CD90 to create tissues containing a 3:1 ratio of these cell types, respectively, that are then mixed together and added to a collagen-based matrix solution. Resulting hECTs are, thus, completely defined in both their cellular and extracellular matrix composition.
Here we describe the construction of defined hECTs as a model system to understand mechanisms of cell-cell interactions in cell therapies, using an example of human bone marrow-derived mesenchymal stem cells (hMSC) that are currently being used in human clinical trials. The defined tissue composition is imperative to understand how the hMSCs may be interacting with the endogenous cardiac cell types to enhance tissue function. A bioreactor system is also described that simultaneously cultures six hECTs in parallel, permitting more efficient use of the cells after sorting.

This manuscript describes the creation of defined engineered cardiac tissues using surface marker expression and cell sorting. The defined tissues can then be used in a multi-tissue bioreactor to investigate mechanisms of cardiac cell therapy in order to provide a functional, yet controlled, model system of the human heart.

Human cardiac tissue engineering can fundamentally impact therapeutic discovery through the development of new species-specific screening systems that ...

Production of Metal Nanoparticles by Pulsed Laser-ablation in Liquids: A Tool for Studying the Antibacterial Properties of Nanoparticles

The emergence of multidrug-resistant bacteria is a global clinical concern leading some to speculate about our return to a "pre-antibiotics" era of medicine. In addition to efforts to identify novel small-molecule antimicrobial drugs, there has been great interest in the use of metal nanoparticles as coatings for medical devices, wound dressings, and consumer packaging, due to their antimicrobial properties. The wide variety of methods available for nanoparticle synthesis results in a broad spectrum of chemical and physical properties which can affect antibacterial efficacy. This manuscript describes the pulsed laser-ablation in liquids (PLAL) method to create nanoparticles. This approach allows for the fine tuning of nanoparticle size, composition, and stability using post-irradiation methods as well as the addition of surfactants or volume excluders. By controlling particle size and composition, a large range of physical and chemical properties of metal nanoparticles can be explored which may contribute to their antimicrobial efficacy thereby opening new avenues for antibacterial development.

The antimicrobial properties of metals such as copper and silver have been recognized for centuries. This protocol describes pulsed laser-ablation in liquids, a method of synthesizing metal nanoparticles that provides the ability to fine tune the properties of these nanoparticles to optimize their antimicrobial effects.

The emergence of multidrug-resistant bacteria is a global clinical concern leading some to speculate about our return to a "pre-antibiotics" era of ...

Three-dimensional Tissue Engineered Aligned Astrocyte Networks to Recapitulate Developmental Mechanisms and Facilitate Nervous System Regeneration

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to replace lost neurons and regenerate axonal pathways. However, during nervous system development, neuronal migration and axonal extension often occur along pathways formed by other cells, referred to as &#34;living scaffolds&#34;. Seeking to emulate these mechanisms and to design a strategy that circumvents the inhibitory environment of the CNS, this manuscript presents a protocol to fabricate tissue engineered astrocyte-based &#34;living scaffolds&#34;. To create these constructs, we employed a novel biomaterial encasement scheme to induce astrocytes to self-assemble into dense three-dimensional bundles of bipolar longitudinally-aligned somata and processes. First, hollow hydrogel micro-columns were assembled, and the inner lumen was coated with collagen extracellular-matrix. Dissociated cerebral cortical astrocytes were then delivered into the lumen of the cylindrical micro-column and, at a critical inner diameter of &#60;350 &#181;m, spontaneously self-aligned and contracted to produce long fiber-like cables consisting of dense bundles of astrocyte processes and collagen fibrils measuring &#60;150 &#181;m in diameter yet extending several cm in length. These engineered living scaffolds exhibited &#62;97% cell viability and were virtually exclusively comprised of astrocytes expressing a combination of the intermediate filament proteins glial-fibrillary acidic protein (GFAP), vimentin, and nestin. These aligned astrocyte networks were found to provide a permissive substrate for neuronal attachment and aligned neurite extension. Moreover, these constructs maintain integrity and alignment when extracted from the hydrogel encasement, making them suitable for CNS implantation. These preformed constructs structurally emulate key cytoarchitectural elements of naturally occurring glial-based &#34;living scaffolds&#34; in vivo. As such, these engineered living scaffolds may serve as test-beds to study neurodevelopmental mechanisms in vitro or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding following CNS degeneration in vivo.

We showcase the development of self-assembled, three-dimensional bundles of longitudinally aligned astrocytic somata and processes within a novel biomaterial encasement. These engineered &#34;living scaffolds&#34;, exhibiting micron-scale diameter yet extending centimeters in length, may serve as test-beds to study neurodevelopmental mechanisms or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding.

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to ...

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved useful as a convenient live imaging platform providing capabilities for precise control of experimental conditions in real time. In this article, we demonstrate live imaging of individual worms employing WormSpa, a previously-published custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes. To illustrate the capability, we performed proof-of-principle experiments in which worms were infected in the device with pathogenic bacteria, and the dynamics of expression of immune response genes and egg laying were monitored continuously in individual animals. The simple design and operation of this device make it suitable for users with no previous experience with microfluidic-based experiments. We propose that this approach will be useful for many researchers interested in longitudinal observations of biological processes under well-defined conditions.

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved ...