Alberto Guiggiani for developing the real time control system, Evelina Chieregatti and Hanako Tsushima for insightful discussions, Giacomo Pruzzo and Alessandro Parodi for the development of custom electronics and software, and Claudia Chiabrera and Marina Nanni for their expert advice and assistance in cell culture preparation.

1. Optical Setup
The entire optical system was described earlier15. Briefly, the optical tweezers system is based on an ytterbium continuous wave (CW) fiber laser operating at 1064 nm (IPG Laser GmbH). A spatial light modulator (SLM) (LCOS-SLM, model X10468-07 - Hamamatsu) varies the phase of the incoming IR laser beam to control the position of the trapping focus spot on the culture dish by computer generated holograms. The freely available Blue-tweezers software (web link on equipment table) generated holograms projected on the spatial light modulator. The interferometer for force spectroscopy measurements was based on a four-quadrant photodiode (QPD, S5980 with C5460SPL 6041 board - Hamamatsu) and a photodiode (PD, PDA100A-EC - Thorlabs).
The laser dissection source was a pulsed sub-nanosecond UV Nd:YAG laser at 355 nm (PNV-001525-040, PowerChip nano-Pulse UV laser - Teem Photonics). An acousto-optic modulator (MQ110-A3-UV, 355 nm fused silica -AA-Opto-electronic) controlled the power of the UV laser delivered to the sample.
The holographic optical tweezers and laser micro-dissector were integrated on a modified upright microscope (BX51 - Olympus) equipped with a 60X, 0.9 NA water dipping objective.The stage of the microscope is composed of a 3-axis linear DC motor micro-positioning system (M-126.CG1, Physics-Instruments) carrying a separate 3-axis piezoelectric nano-positioning stage (P-733.3DD, Physics-Instruments) to combine coarse movement of the sample with the sub-nanometer resolution of the piezo-stage. The microscope stage system was equipped with two control loops synergistically acting to maintain the trapping focus spot at the right position, depending on the selected working mode (position or force clamp, static or dynamic)16. In particular, an internal feedback loop acts on a piezoelectric stage, to keep the bead at a selected distance from the trap center. The other external loop controls the position of the motorized stage to exploit the region spanned by the piezo-actuator on a larger area than its available stroke17. When the piezo-stage reaches the limit of the available stroke in one direction, the external loop moves the micro stage in the opposite direction, thus the piezo recovers toward its central position because it is tracking the trapped bead adhered to the sample. When the piezo-stage reaches the central position of its course range, the micro stage halted. Further details of the system are reported in Guiggiani et al16,17.
A Peltier device (QE1 resistive heating with TC-344B dual channel heater controller - Warner Instruments) controls the temperature of the cell culture under the microscope (37 &deg;C). In the culture, pH and humidity were maintained at physiological conditions by aerating a custom-designed polydimethylsiloxane (PDMS) sleeve (integrating the microscope objective) with humidified carbogen (95% O2, 5% CO2).
2. Cell Culture Preparation
All the experimental protocols were approved by the Italian Ministry of Health. Primary cultures were obtained from hippocampi of mice (C57BL6J, Charles River) at embryonic day 18 (E18).
<ol>
	<li>Neurons were plated at a concentration of 25,000 cells/ml on glass-bottom Petri dishes (P35G-0-14-C - MaTek Corporation). The low concentration of cultured cells is needed to avoid the formation of a dense network already in the first days in vitro. At the mentioned cell concentration, the likelihood of finding isolated cells with a longer neurite not connected to other cells is much higher.</li>
</ol>
3. Bead Coating
<ol>
	<li>Polymer microspheres (&Oslash; 4 &mu;m, COOH-terminated- Bangs Laboratories, product code PC05N/6700) were coated with poly-D-lysine following the procedure described in the polyLink Protein Coupling Kit (Polysciences, product code 19539). The beads are coated with the same molecule used to cover the culture support and favor cells adhesion.</li>
</ol>
4. Choose Isolated Neuron. Detach A Bead from the Culture Substrate, Trap and Move it Next to the Neuron
<ol>
	<li>Put the culture Petri dish on the microscope stage. After setting the focus on the sample by stage motion, correct the position of the illuminating objective condenser to set Kohler-illumination. Aligning the illumination optics to set the Kolher-illumination condition is necessary to improve the bright field imaging quality. Using such alignment conditions, it is also necessary to maximize the IR laser collection efficiency (through objective condenser), from the scattering trapped probe, to perform its interferometric tracking by the QPD and PD.</li>
	<li>Pipette a few &mu;l of the coated microsphere stock solution into the culture dish. Microspheres will deposit and adhere to the culture substrate. The number of &mu;l injected into the culture dish depends on the microsphere concentration of the stock solution. The ideal condition is to have between one and two microspheres per imaging field of view. When too many microspheres are added to the culture dish, they can already attach randomly to the cultured neurons.</li>
	<li>Move around in the culture dish, searching for an isolated neuron with a longer neurite (the axon), and save the position of the microscope stage.</li>
	<li>Move around and search for a bead adhered to the culture support.</li>
	<li>Turn on the IR trapping laser. At this stage, no holograms are projected on SLM, and the position of IR laser spot coincides with the UV laser spot position. Set the axial position of the IR spot 2-3 &mu;m above the culture support surface, by microscope stage motion.</li>
	<li>Set the UV laser power delivered to the sample to less than 1 &mu;W. The UV laser dissector has been previously calibrated14: 
	5-6 &mu;W the glass support is ablated 
	4 &mu;W a neuronal connection is completely dissected 
	2.5 &mu;W a neurite is partially lesioned 
	&lsaquo; 1 &mu;W a shock wave is produced in the culture dish. The optical shock wave is strong enough to detach the bead from the glass support.</li>
	<li>Turn on the UV laser, while leaving the IR laser on, and move the UV spot above the adhered bead by microscope stage motion. The bead detaches from the surface, and it is trapped by the IR laser. Then, turn off the UV laser.</li>
	<li>Move the trapped bead several micrometers above the glass support (20-30 &mu;m). Lifting the bead to this position will prevent it from contacting other cells while it is being moved toward the previously chosen neuron. Bring the bead to the previously saved stage position. Set the stage speed to a low value (10 &mu;m/sec) so the drag fluid force does not exceed the optical trapping force.</li>
</ol>
5. Move the Trap Position with Respect to the Laser Dissector Spot and Axon Position by Computer Generated Hologram, and Calibrate the Optical Tweezers Stiffness
<ol>
	<li>Move the bead toward the glass support to visualize the axon. The bead is held above the cell to avoid contact with it (about 5 &mu;m above the glass support).</li>
	<li>Move the microscope stage to move the UV focus spot on the center of the width of the axon. Save the current stage position.</li>
	<li>Move the IR focus spot position by computer-generated hologram. In this step, the stage is halted in the position chosen in the previous step. When the computer generated hologram is projected on the SLM, the trapped bead is moved with respect to the axon. We move the IR laser spot to have the trapped bead aligned on the center of the neurite, with the UV laser spot centered on the same neurite too. The IR spot and thus the trapped bead are positioned along the neurite 5-10 &mu;m away from the UV spot. We move the position of the IR with respect to the UV spot depending on the neurite geometry. In case the optical tweezers are not equipped with an SLM system, the position can be varied by tilting mirrors, or acoustic optic deflectors, on the IR optical path. The IR laser spot may be positioned 5-10 &mu;m away from the UV spot to avoid optical interaction of the dissecting beam with the trapped probe, which could influence its Brownian motion and alter the force spectroscopy traces.</li>
	<li>After defining the new IR spot position with respect to UV spot and axon position, move the stage to position the trapped bead away from the axon and avoid collision with it. Move the axial position of the trapped bead to about 2 &mu;m above the cover glass.</li>
	<li>Align the QPD to center the interference fringes on it: x and y QPD differential signals are zeros when the QPD is centered. Acquire 5 sec of the Brownian motion of the trapped bead by the interferometer, at 50 KHz sampling rate. Obtain the stiffness (pN/nm) and sensitivity (V/nm) of the optical trap by the power spectrum method18.</li>
</ol>
6. Attach the Bead to the Axon. Perform Axotomy and Simultaneous Force Measurement
<ol>
	<li>Raise the trapped bead position about 4 &mu;m above the cover glass, and move it to the previously saved stage position (see step 3.2).</li>
	<li>Move the trapped bead down toward the axon until it contacts the neurite. The collision between the trapped bead and the axon is monitored by the z QPD signal detecting displacement of the trapped bead in the axial direction.</li>
	<li>Wait 10 sec with the trapped bead pushed against the axon to favor its adhesion to the neurite membrane.</li>
	<li>Move the micro stage to displace the trapped bead from the axon, and thus verify its adhesion to the cell membrane. If the bead adhered, it escapes from the optical trap.</li>
	<li>Move back the laser trap on the adhered bead and switch on the force-clamp loop with force condition equal to zero. In such a way, the piezo-stage repositions the bead, attached to the cell, on the optical trap center. If the system does not have a feedback loop control, the laser trap position can be moved by the microscope stage to center it on the adhered probe. The center of the trap is reached when the QPD signals are the same as when the bead is trapped and not adhered to the cell (x and y QPD signals equal to zero, and the z QPD signal gives the same voltage sum of the four quadrants as when the bead is trapped far from any surface) .</li>
	<li>Set a force-clamp condition on the z axis, positioning the trapping laser slightly over the center of the bead (about 100 nm), to generate a pretension on the adhered trapped probe. In case the system is not equipped with a force-clamp control, the optical trap position can be raised up at steps of 25 nm by the microscope stage. The z QPD signal allows monitoring. Therefore, use this signal to set the position of the laser trap with respect to the adhered probe. Such pre-tension is needed to sense the strain on the membrane after the axonal lesion, and the consequent decrease of Brownian motion of the adhered probe. A similar approach has been proposed to measuring the release of tension in a membrane tether by video imaging19.</li>
	<li>Switch-off the force-clamp feedback to measure the force on the adhered probe in the position-clamp condition (the piezo-stage is blocked).</li>
	<li>Start simultaneous recording of the trapped probe positions through the interferometer (20 KHz sampling frequency), and the cell during laser axotomy time-lapse bright-field imaging (20 Hz frame rate).</li>
	<li>Turn on the UV laser to deliver laser pulses until a lesion becomes visible on the image of the axon (Usually 200-400 optical pulses are needed. Energy per pulse: 25 nJ), then turn off the UV laser.</li>
	<li>Continue recording by the interferometer for about 3 min, until the x, y and z QPD traces reach a plateau.</li>
</ol>
7. Quantify the Total Tension Release
<ol>
	<li>Convert the recorded QPD traces (in Volts) by the calibrated sensitivity of the optical trap, to obtain the respective traces in nanometers.</li>
	<li>Filter the x, y, and z displacement traces by a high pass filter with cut off at 10 Hz.</li>
	<li>Calculate the total variance of the filtered traces representing the Brownian motion of the trapped bead. Calculate the variance at 25 msec steps for overlapping time windows of 500 msec (10,000 data points)20. The high pass filtering of traces excludes the changes of Brownian motion due to the membrane fluctuation or cortical actin motion. The Brownian motion of the bead is reduced by the optical forces and the adhesion forces on the cell membrane. When the membrane is strained because of the release of tension its viscosity is increased, and thus the Brownian motion of the bead, detected immediately after axotomy, starts to decrease.</li>
	<li>Define t0: the beginning of decrease of the Brownian motion variance, after the delivery of UV laser energy. The time instant t0 indicates the beginning of membrane strain.</li>
	<li>Define t1: the end of decrease of the Brownian motion variance (when it reaches a plateau). The time instant t1 indicates the end of membrane strain.</li>
	<li>Perform video tracking of debris or a scratch on the cover glass support, to measure any drift of the sample during the force measurement. To track the particle on the glass support, we first apply thresholds to the bright-field images, to obtain a stack of binary images with the particle in white on a black background. Then, we track the particle center of mass with sub-pixel accuracy (using the freeware ImageJ software).</li>
	<li>Subtract the measured drift from the displacement QPD traces. Multiply the drift-corrected QPD traces by the respective calibrated optical stiffness (kx, ky, kz) to obtain the Fx, Fy, Fz traces in piconewtons.</li>
	<li>Calculate the total force trace as Ftot = &radic;(Fx2+Fy2+Fz2).</li>
	<li>Calculate the total release of force between t0 and t1 as Freleased= Ftot(t1) - Ftot(t0). The force measured at t0 has to be subtracted because it is due to the displacement of the probe from the trap center, when the bead adheres to the neurite.</li>
	<li>Calculate the neurite contact area between the trapped bead and the UV laser spot position: on the bright field image measure the long (L) and short (D) axes of the neurite between the trapped bead and the UV laser spot position. The axon contact area is Aaxon = L x D.</li>
	<li>Normalize the total released force Freleased by the axon contact area Aaxon.</li>
</ol>

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The authors declare that they have no competing financial interests.

We report in this work a quantitative method to compare the neurite adhesion to the culture substrate, by performing simultaneous force spectroscopy measurement during laser induced cell lesion. The measured release of tension is related to the degree of adhesion of the cell to the substrate: cells with a higher number of focal adhesions should release less tension. Measuring the release of tension in terms of piconewtons provides a physical quantity by which to evaluate the axonal adhesion to the culture support in diff...

Optical microscopy is one of the less invasive imaging system available to observe living cells. With the exploitation of effects such as radiation pressure (as in optical tweezers1), or high-energy photon flux (as in laser dissector2), this technology was extended to nano-manipulation. The optical imaging system furnishes a precise control to visualize and manipulate sub cellular targets3. At the same time, thanks to the accurate calibration of the delivered laser power, optical tools accomplish either soft or invasive sample manipulation with unprecedented reproducibility.
Several laboratories integrated, in the same experimental setup, optical tweezers and laser dissector in order to ablate organelles4, to fuse together different cells5, or to stimulate cells by optically driven cargos6,7. While optical tweezers, after calibration of the optical stiffness, allow for the control of applied force to the cell on a piconewton scale, laser dissection systems can modulate optical manipulation, which ranges from membrane photo-poration to ablation of single organelles or dissection of sub-cellular structures. However, laser dissection calibration relies on qualitative assessment of the entity of optical manipulation with respect to the energy delivered to the sample, mainly based on image analysis illustrating morphological changes caused to the specimen8. In the presented method, we demonstrate how to perform force spectroscopy measurement during the laser axonal dissection of a developing neuron, to quantify, on piconewton scale, the force produced by an altered equilibrium in the cytoskeleton structure of a sub-cellular compartment9. Cultured neurons adhere to the substrate, and polarize during development. The polarization phase occurs during the first five days in vitro. At stage two of polarization, one of the extruding neurites becomes longer, and it will differentiate to become the axon10. Axonal elongation in response to towing force at the growth cone has been previously modeled by Dennerl's model11. Recently, this model has been extended12 to include the role of neurite adhesion to the extracellular matrix substrates. This biophysical model, proposed after experimental observations13, showed that pulling forces on growth cone, propagating along the neurite, are modulated by focal adhesions to the substrate. Likewise, axonal lesion produces a local release of tension propagating toward the cell body. Thus, we proposed that measuring such released tension in a location along the axon between the lesion and the cell soma offers the possibility to assess the dampening outcome of unaffected focal adhesions.
We calibrate the necessary energy photon-flux of the laser dissector to control the extent of the inflicted axonal damage, from complete transection to partial lesion. Following the calibration, we repeated partial lesion to the axons of several differentiating neurons and developed the protocol to quantify the tension release, and thus obtained a quantitative parameter to estimate the adhesion of the axon to the substrate14.
In the present work, we describe in detail the developed protocol, which represents a precise experimental procedure to evaluate and compare with piconewton sensitivity the axonal adhesion to the substrate in different experimental conditions such as chemical treatment14, or different types of cell culture support.

<table><tbody><tr><td colspan="4">&lt;strong&gt;REAGENTS&lt;/strong&gt;</td></tr><tr><td>Polymer microspheres, 4 &amp;mu;m, COOH coated</td><td>Bangs laboratories</td><td>PC05N/6700</td><td></td></tr><tr><td>PolyLink Protein Coupling Kit</td><td>Polyscience</td><td>19539</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;EQUIPMENT&lt;/strong&gt; </td></tr><tr><td>IR laser</td><td>IPG Laser GmbH</td><td>YLM-5-SC-LP</td><td>ytterbium continuous wave (CW) fiber laser operating at 1064 nm, with linear polarization</td></tr><tr><td>Spatial light modulator</td><td>Hamamatsu</td><td>LCOS-SLM 10468-07</td><td></td></tr><tr><td>Blue-tweezers software</td><td>Optics group, University of Glasgow</td><td>Free downloadable software</td><td>&lt;a href=&quot;http://www.physics.gla.ac.uk/Optics/projects/tweezers/slmcontrol/&quot; target=&quot;_blank&quot;&gt;http://www.physics.gla.ac.uk/Optics/projects/tweezers/slmcontrol/&lt;/a&gt;</td></tr><tr><td>ImageJ</td><td>Hamamatsu</td><td>Free downloadable software</td><td>&lt;a href=&quot;http://rsbweb.nih.gov/ij/&quot; target=&quot;_blank&quot;&gt;http://rsbweb.nih.gov/ij/ &lt;/a&gt;</td></tr><tr><td>QPD</td><td>Thorlabs</td><td>S5980 with C5460SPL 6041 board</td><td>Four quadrant photo-diode to measure x, y trapped probe displacement</td></tr><tr><td>PD</td><td>Teem Photonics</td><td>PDA100A-EC</td><td>Photodiode to measure z trapped probe displacement</td></tr><tr><td>nano-Pulse UV laser</td><td>AA-optoelctronics</td><td>PNV-001525-040</td><td>Pulsed UVA laser, pulse length 400 ps</td></tr><tr><td>Acoustic Optic Modulator</td><td>Olympus</td><td>MQ110-A3-UV, 355nm fused silica</td><td></td></tr><tr><td>Upright microscope</td><td>Andor</td><td>BX51</td><td>Equipped with a 60, 0.9 NA, water dipping objective</td></tr><tr><td>CCD</td><td>Warner Instruments</td><td>V887ECSUVB EMCCD</td><td></td></tr><tr><td>Peltier device</td><td>Physic Instruments</td><td>QE1 resistive heating with TC-344B dual channel heater controller</td><td></td></tr><tr><td>Microscope stage: micro+piezo stage</td><td>National Instruments</td><td>Three linear stages M-126.CG1 carrying a separate 3-axis piezoelectric nano-positioning stage P-733.3DD</td><td></td></tr><tr><td>Daq</td><td></td><td>NI PCI-6229</td><td>Acquiring the x, y, z position of the trapped probe, and sending feedback loop signals to microscope stage</td></tr><tr><td>Linux Real Time Application Interface (RTAI) machine</td><td></td><td></td><td>Real time feedback loop system, to control stage position, developed on a dedicated PC desktop</td></tr></tbody></table>

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.

The cell generates traction forces on the substrate by its focal adhesions. Force generated by cytoskeletal elements are in equilibrium with the counteracting force of the culture substrate. After laser induced lesion of the neurite, some of the tensed cytoskeleton cables are disrupted and their equilibrated tension is released because the opposing force of the substrate adhesion is eliminated. The released tension is partially distributed on the unaffected focal adhesions, and the bead attached to the cell membrane, hel...

Watch this Scientific Journal Video about Measurement of Tension Release During Laser Induced Axon Lesion to Evaluate Axonal Adhesion to the Substrate at Piconewton and Millisecond Resolution at JoVE.

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

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 ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

10.2K Views.  National Research Council of Italy. 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.

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

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.

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.

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

Mechanical Manipulation of Neurons to Control Axonal Development

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the 10-1000 &mu;dyne range1,2. Force measurements are obtained through observation of the Hookean bending of the glass needles, which are calibrated by a direct and empirical method3. Equipment requirements and procedures for fabricating, calibrating, treating, and using the needles on cells are fully described. The force regimes previously used and different cell types to which these techniques have been applied demonstrate the flexibility of the methodology and are given as examples for future investigation4-6. The technical advantages are the continuous 'visualization' of the forces produced by the manipulations and the ability to directly intervene in a variety of cellular events. These include direct stimulation and regulation of axonal growth and retraction7; as well as detachment and mechanical measurements on any type of cultured cell8.

Application and direct measurements of forces on neurons in the 2-1000 microdyne range are achieved with high precision using calibrated glass needles. This methodology can be used to control and measure several aspects of axonal development, including axonal initiation, axonal tension, velocity of axonal elongation, and force vectors.

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the ...

Neuroscience

Laser Nanosurgery of Cerebellar Axons In Vivo

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

Laser Nanosurgery of Cerebellar Axons In Vivo

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

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

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated calcium channels (VGCCs) located in the release face membrane, is the trigger for vesicle fusion and neurotransmitter release. Crucial to the rapidity of synaptic transmission is the spatial and temporal synchrony between the arrival of the action potential, VGCCs and the neurotransmitter release machinery. The ability to directly record Ca2+&#160;currents from the release face membrane of individual presynaptic terminals is imperative for a precise understanding of the relationship between presynaptic Ca2+&#160;and neurotransmitter release. Access to the presynaptic release face membrane for electrophysiological recording is not available in most preparations and presynaptic Ca2+&#160;entry has been characterized using imaging techniques and macroscopic current measurements &#8211; techniques that do not have sufficient temporal resolution to visualize Ca2+ entry. The characterization of VGCCs directly at single presynaptic terminals has not been possible in central synapses and has thus far been successfully achieved only in the calyx-type synapse of the chick ciliary ganglion and in rat calyces. We have successfully addressed this problem in the giant reticulospinal synapse of the lamprey spinal cord by developing an acutely dissociated preparation of the spinal cord that yields isolated reticulospinal axons with functional presynaptic terminals devoid of postsynaptic structures. We can fluorescently label and identify individual presynaptic terminals and target them for recording. Using this preparation, we have characterized VGCCs directly at the release face of individual presynaptic terminals using immunohistochemistry and electrophysiology approaches. Ca2+ currents have been recorded directly at the release face membrane of individual presynaptic terminals, the first such recording to be carried out at central synapses.

Recording Ca2+ currents at the presynaptic release face membrane is key to a precise understanding of Ca2+ entry and neurotransmitter release. We present an acute dissociation of the lamprey spinal cord that yields functional isolated reticulospinal axons, permitting recording directly from the release face membrane of individual presynaptic terminals.

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Developmental Biology

High-Speed Magnetic Tweezers for Nanomechanical Measurements on Force-Sensitive Elements

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are therefore poised to be useful in the field of mechanobiology. Since the method commonly relies on image-based tracking of magnetic beads, the speed limit in recording and analyzing images, as well as the thermal fluctuations of the beads, has long hampered its application in observing small and fast structural changes in target molecules. This article describes detailed methods for the construction and operation of a high-resolution MT setup that can resolve nanoscale, millisecond dynamics of biomolecules and their complexes. As application examples, experiments with DNA hairpins and SNARE complexes (membrane-fusion machinery) are demonstrated, focusing on how their transient states and transitions can be detected in the presence of piconewton-scale forces. We expect that high-speed MTs will continue to enable high-precision nanomechanical measurements on molecules that sense, transmit, and generate forces in cells, and thereby deepen our molecular-level understanding of mechanobiology.

Here, we describe a high-speed magnetic tweezer setup that performs nanomechanical measurements on force-sensitive biomolecules at the maximum rate of 1.2 kHz. We introduce its application to DNA hairpins and SNARE complexes as model systems, but it will be also applicable to other molecules involved in mechanobiological events.

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are ...

Biochemistry

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

DNA Tension Probes to Map the Transient Piconewton Receptor Forces by Immune Cells

Mechanical forces transmitted at the junction between two neighboring cells and at the junction between cells and the extracellular matrix are critical for regulating many processes ranging from development to immunology. Therefore, developing the tools to study these forces at the molecular scale is critical. Our group developed a suite of molecular tension sensors to quantify and visualize the forces generated by cells and transmitted to specific ligands. The most sensitive class of molecular tension sensors are comprised of nucleic acid stem-loop hairpins. These sensors use fluorophore-quencher pairs to report on the mechanical extension and unfolding of DNA hairpins under force. One challenge with DNA hairpin tension sensors is that they are reversible with rapid hairpin refolding upon termination of the tension and thus transient forces are difficult to record. In this article, we describe the protocols for preparing DNA tension sensors that can be "locked" and prevented from refolding to enable "storing" of mechanical information. This allows for the recording of highly transient piconewton forces, which can be subsequently "erased" by the addition of complementary nucleic acids that remove the lock. This ability to toggle between real-time tension mapping and mechanical information storing reveals weak, short-lived, and less abundant forces, that are commonly employed by T cells as part of their immune functions.

This paper describes a detailed protocol for using DNA-based tension probes to image the receptor forces applied by immune cells. This approach can map receptor forces &gt;4.7pN in real-time and can integrate forces over time.

Mechanical forces transmitted at the junction between two neighboring cells and at the junction between cells and the extracellular matrix are critical ...

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study of this movement in situ was only possible using radioisotopic pulse-labeling, which permits analysis of axonal transport in whole nerves with a temporal resolution of days and a spatial resolution of millimeters. To study neurofilament transport in situ with higher temporal and spatial resolution, we developed a hThy1-paGFP-NFM transgenic mouse that expresses neurofilament protein M tagged with photoactivatable GFP in neurons. Here we describe fluorescence photoactivation pulse-escape and pulse-spread methods to analyze neurofilament transport in single myelinated axons of tibial nerves from these mice ex vivo. Isolated nerve segments are maintained on the microscope stage by perfusion with oxygenated saline and imaged by spinning disk confocal fluorescence microscopy. Violet light is used to activate the fluorescence in a short axonal window. The fluorescence in the activated and flanking regions is analyzed over time, permitting the study of neurofilament transport with temporal and spatial resolution on the order of minutes&#160;and microns, respectively. Mathematical modeling can be used to extract kinetic parameters of neurofilament transport including the velocity, directional bias and pausing behavior from the resulting data. The pulse-escape and pulse-spread methods can also be adapted to visualize neurofilament transport in other nerves. With the development of additional transgenic mice, these methods could also be used to image and analyze the axonal transport of other cytoskeletal and cytosolic proteins in axons.

We describe fluorescence photoactivation methods to analyze the axonal transport of neurofilaments in single myelinated axons of peripheral nerves from transgenic mice that express a photoactivatable neurofilament protein.

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study ...

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

Summary

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Chapters in this video

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

Mechanical Manipulation of Neurons to Control Axonal Development

Laser Nanosurgery of Cerebellar Axons In Vivo

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

High-Speed Magnetic Tweezers for Nanomechanical Measurements on Force-Sensitive Elements

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

DNA Tension Probes to Map the Transient Piconewton Receptor Forces by Immune Cells

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve