This work has been supported by a grant from the European Community [FP7-PEOPLE-2011-IAPP project 285827 (EPIXCHANGE)].

All experimental protocols were approved by the University of Ferrara Ethical Committee for Animal Experimentation and by the Italian Ministry of Health. The ARRIVE (Animal Research: Reporting In Vivo Experiments11) guidelines have been followed.
1. Preparation of Quartz Needles
<ol>
	<li>Clean and sterilize the quartz capillaries (see Table of Materials) by placing them for 5 min in distilled water, 5 min in ethanol 99%, and 5 min in diethyl-ether.</li>
	<li>Leave capillaries under the hood for at least 1 h to let them dry completely.</li>
	<li>Prepare the needles using a laser puller according to parameters that allow the production of a sufficiently long, sufficiently thin needle, depending on the individual needs. 
	NOTE: We use a laser puller (see Table of Materials). Use a two lines program for pulling. The first line uses high power, the largest scanning length (corresponding to FILAMENTS VALUE) and no pulling force. This combination of parameters leads to the melting of the largest portion of the capillary with the lowest extension speed, allowing for the shaping of a long needle with large inner/external size and reduced shank. In this way, the needle preserves a high mechanical resistance and wide inner diameter that allows the passage of stem cells or viral particles without clogging. Velocity and delay of this program line should stop the capillary extension before the tip becomes too small and long, missing the required mechanical and shape characteristics. The second line of the pulling program is executed automatically, once the quartz cools down (5 s). Parameters of power, filament and pulling force are functional to cut a tip with the shortest possible tip length and steep shank, avoiding the separation of the two needles with a tip rupture. Routinely, with our equipment we use the following: (a) Line 1 = power 990, filament 5, velocity 130, delay 110, pull 0; (b) Line 2 = power 970, filament 3, velocity 0, delay 70, pull 0.</li>
	<li>Put the needles in a Petri dish previously cleaned with ethanol 70%.
	<ol>
		<li>Cover the dish with a plastic paraffin film.</li>
	</ol>
	</li>
	<li>Cut the capillaries at the micro-dissector.
	<ol>
		<li>At the start-up, let the laser run an autocalibration process to verify power and energy delivered. Then set the cutting parameters. 
		NOTE: We use a micro-dissector (see Table of Materials) and the following parameters: laser power pulse energy (63 &#181;J), laser beam diameter of 20% of the maximal aperture (maximal aperture: 1.0 &#181;m), and cutting speed of 10% of the maximal speed (maximal speed: 30 &#181;m/s).</li>
		<li>Firmly fix the needle on the microscope table and position its tip at the center of the computer screen.</li>
		<li>On the menu bar, click on the pencil icon and use it to draw a mark for the laser cutting of the needle. 
		NOTE: The cut should be at an angle of about 45 &#176; with respect to the needle surface.</li>
		<li>On the side bar function, select the flag containing the &#34;START CUT&#34; button to allow the laser to cut the previously marked spot.</li>
		<li>If needed, cut side hole(s) by repeating the procedures described in 1.5.2-3; there is no need to reposition the needle (Figure 1).</li>
	</ol>
	</li>
	<li>Post-cut cleaning.
	<ol>
		<li>Connect the needle to a void pump through a plastic tube.</li>
		<li>Turn the pump on and manually set the forward flow rate at a moderate speed (3 &#181;L/min).</li>
		<li>Aspirate the 99% ethanol first and then the distilled water to remove impurities due to cutting.</li>
		<li>Aspirate the 99% ethanol again to ease drying. 
		NOTE: Each step should last at least 3 min.</li>
	</ol>
	</li>
	<li>Store the needles in a properly covered Petri dish until the day of surgery.</li>
	<li>Needles may be re-used; at the end of the experiment, clean with bleach and ethanol to remove the solution and possible tissue residue. For a more complete cleaning, put the needles in a micropipette storage jar with the tip pointing downwards, fill the jar with distilled water until the tips are covered, boil for 1.5 min to remove any leftover tissue, then wash with ethanol.</li>
</ol>
2. Procedure
<ol>
	<li>Following the instructions provided by the pump user manual, select the settings menu to calibrate the pump according to the diameter/volume of the microinjection syringe and set the flow rate at 0.3 &#181;L/min.</li>
	<li>Fill a 10-mL syringe (equipped with a blunt needle (e.g., Hamilton needle, 30 G) and a short polytetrafluoroethylene tube) with sterile water and use it to fill the microinjection syringe (after having very cautiously removed the piston) and the injection assembly parts kit for the manual microinjection pump (MMP). 
	NOTE: The MMP kit is used to securely mount the needle in the stereotactic frame (Figure 2). The kit includes: a polytetrafluoroethylene tubing, luer-to-tubing coupler, pipette holder and gaskets for pipettes with different outer diameters.</li>
	<li>Connect the needle to the MMP kit.
	<ol>
		<li>Flush more water across the system through the 10-mL syringe, and check whether the water comes out of the needle. 
		NOTE: Make sure to remove any air bubble inside the system by repeating the filling steps.</li>
	</ol>
	</li>
	<li>Disconnect the 10-mL syringe from the microinjection syringe while slowly pushing the piston in order to leave a drop of water at the top of the microinjection syringe. Insert the piston in the microinjection syringe.</li>
	<li>Connect the microinjection syringe to the pump and set the flow rate at 0.3 &#181;L/min. 
	NOTE: Wait until there is a drop leaking from the tip of the needle and then let it go for another 3 min.</li>
	<li>Set the flow at 0.1 &#181;L/min and start the surgery procedure. The pump should be ongoing while performing the surgery. 
	NOTE: For the experiments reported in this article, we use male Sprague Dawley rats of 300 g (3 months of age). However, the procedure is applicable to other species, mice in particular.</li>
	<li>Induce general anesthesia using a method approved by local regulations. 
	NOTE:&#160;We use a mix of intraperitoneal injection (i.p.) of 85 mg/kg ketamine and 8 mg/kg xylazine for 20-50 min of anesthesia. It is important that tail and/or toe pinches are used to ensure the animal is fully sedated. Level of anesthesia and basic vitals are assessed before surgery begins and monitored every 10 min&#160;while performing the surgery. Animal body temperature is maintained constant before, during and after surgical procedures using a water re-circulating heating pad. Animals are monitored until fully recovered (e.g. they are upright and ambulatory). In addition, animals receive analgesics (tramadol, 5&#8211;7 mg/kg i.p.) post-operatively every 24 h for three days.
	<ol>
		<li>Shave the animal&#39;s head and&#160;adequately prepare the surgical site for aseptic surgery. Perform a two-stage scrub with iodine-based solution (Betadine) followed by 70% ethanol. Furthermore, to maintain asepsis, wrap the surgical site with a surgical drape (in the video has been omitted for demonstration purposes).&#160;</li>
	</ol>
	</li>
	<li>Position the rat in the stereotaxic frame over a water re-circulating heating blanket. To provide additional pain relief, at the site of surgery apply 0.5% lidocaine solution (do not exceed 7 mg/kg). Pre-sterilize all surgical instruments using a bead sterilizer or liquid cold disinfection by immerging the equipment in either 2% glutaraldehyde plus 7.05% phenol (Sporodicin or Cidex) followed by rinse with sterile water or saline. The disinfection must be performed even between procedures. Make a longitudinal cut over the skull by using a sterilized scalpel. Be gentle to avoid excessive bleeding. Open the scalp using a flat spatula and apply surgical clips on the flaps of skin to keep it open. Very gently, detach the periosteum to expose the bregma and the area of the skull through which the needle will be inserted.</li>
	<li>Mount the arm on the stereotaxic frame (while the flow is ongoing) and move the tip of the needle exactly on top of bregma, paying attention not to break its tip. Annotate the antero-posterior and medio-lateral coordinates of the bregma and translate to the desired coordinates. Guided by a stereomicroscope, lower the needle until its tip almost touches the skull, mark the drilling spot after raising a little the stereotaxic arm.</li>
	<li>Drill (drill tip &#216;: 0.8 mm; drill speed: 1,000 rpm) the skull on the marked spot.&#160;Care must be taken not to penetrate the dura during the drilling procedures. During drilling, drip sterile saline to avoid bone damage and necrosis.</li>
	<li>Cut a small piece of plastic paraffin film and place it on the skull. Using the 10 &#181;L pipet, make a drop of the solution to inject on the plastic paraffin film. Stop the pump, slightly retrieve the pusher of the pump and create a little air bubble in the capillary by gently pulling the piston of the microinjection syringe.</li>
	<li>Lower the stereotactic arm until the tip of the needle reaches the drop. Draw the sample by pulling (very slowly) the piston of the microinjection syringe, avoiding any bubble formation. Replace the pump pusher in contact with the piston and turn on the pump at a flow rate of 0.3 &#181;L/min. Wait until the formation of a drop (a few min) before proceeding to the next step. 
	NOTE: In the present example experiment, artificial cerebrospinal fluid (aCSF) was infused in the striatum (AP: +1.5, ML: &#177;2.7; DV: -5.0) and in the dorsal hippocampus (AP: -3.5, ML: &#177;2.1; DV: -3.5; in mm from dura), 2 &#181;L/site. To facilitate the comparison between the two injection methods, each brain area was infused using a quartz needle in one hemisphere and a 26 G blunt tip or a 30 G bevel-tip stainless steel needle in the other.</li>
	<li>Lower the flow to 0.1 &#181;L/min, nick the dura with a stainless steel bevel-tip needle and slowly lower the stereotactic arm on the dorso-ventral axe, entering the brain tissue. When the desired position is reached, go down an additional 0.1 mm, stop the pump, raise the stereotactic arm by 0.1 mm and start again the pump at the flow to 0.3 &#181;L/min. 
	NOTE: The reason to maintain a slow flow while lowering the needle is to ensure a positive pressure that avoids penetration of small fragments of tissue in the needle itself. Such an event would obstruct the tip, impairing the delivery of solution. Since the flow rate is very slow while lowering the needle and the speed of lowering is about 10 s/mm, this implies that the amounts of solution that remain along the needle track are negligible compared to those injected at the target site. This precaution may be avoided when using needles with side holes.</li>
	<li>Start the timer and wait the needed time depending on the desired volume to inject (for example, 6 min 40 s if injecting 2 &#181;L). Monitor the bubble movement while injecting. At the end, stop the pump and wait 5 min before slowly retrieving the needle. When the quartz needle is completely extracted, start the pump again and check for the formation of a drop at the tip of the needle.</li>
	<li>Seal the opening in the skull surface with bone wax, remove the hemostatic clips and suture the scalp, coat the area around the wound with an antibiotic cream (e.g., neosporin) and return the animal to the home cage. 
	NOTE: Following the euthanasia protocol approved by the institutional ethical committee, for the experiments described in this article, animals were deeply anesthetized with an i.p. injection of 43 mg/kg ketamine and 7 mg/kg xylazine, and decapitated. Brains were removed, postfixed in formalin for 48 h and paraffin embedded. Coronal sections (8 &#181;m) of the brains were sliced using a microtome.</li>
</ol>

<ol>
	<li>Kirik, D., Rosenblad, C., Bj&#246;rklund, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+behavioral+and+neurodegenerative+changes+following+partial+lesions+of+the+nigrostriatal+dopamine+system+induced+by+intrastriatal+6-hydroxydopamine+in+the+rat.">Characterization of behavioral and neurodegenerative changes following partial lesions of the nigrostriatal dopamine system induced by intrastriatal 6-hydroxydopamine in the rat.</a> Exp Neurol. 152 (2), 259-277 (1998).</li><li>Paolone, G., Brugnoli, A., Arcuri, A., Mercatelli, D., Morari, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eltoprazine+prevents+levodopa-induced+dyskinesias+by+reducing+striatal+glutamate+and+direct+pathway+activity.">Eltoprazine prevents levodopa-induced dyskinesias by reducing striatal glutamate and direct pathway activity.</a> Mov Disord. 30 (13), 1728-1738 (2015).</li><li>Paolone, G., Lee, T. M., Sarter, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time+to+pay+attention:+attentional+performance+time-stamped+prefrontal+cholinergic+activation,+diurnality,+and+performance.">Time to pay attention: attentional performance time-stamped prefrontal cholinergic activation, diurnality, and performance.</a> J Neurosci. 32 (35), 12115-12128 (2012).</li><li>Shoichet, M. S., Winn, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+delivery+to+the+central+nervous+system.">Cell delivery to the central nervous system.</a> Adv Drug Deliv Rev. 42, 81-102 (2000).</li><li>Simonato, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progress+in+gene+therapy+for+neurological+disorders.">Progress in gene therapy for neurological disorders.</a> Nature RevNeurol. 9, 277-291 (2013).</li><li>Bj&#246;rklund, H., Olson, L., Hahl, D., Schwarcz, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Short-and+Long-Term+Consequences+of+Intracranial+Injections+of+the+Excitotoxin,+Quinolinic+Acid,+as+Evidenced+by+GFA+Immunohistochemistry+of+Astrocytes.">Short-and Long-Term Consequences of Intracranial Injections of the Excitotoxin, Quinolinic Acid, as Evidenced by GFA Immunohistochemistry of Astrocytes.</a> Brain Res. 317, 267-277 (1986).</li><li>Paradiso, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localized+delivery+of+fibroblast+growth+factor-2+and+brain-derived+neurotrophic+factor+reduces+spontaneous+seizures+in+an+epilepsy+model.">Localized delivery of fibroblast growth factor-2 and brain-derived neurotrophic factor reduces spontaneous seizures in an epilepsy model.</a> Proc Natl Acad Sci U S A. 106 (17), 7191-7196 (2009).</li><li>Falcicchia, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Silencing+Status+Epilepticus-Induced+BDNF+Expression+with+Herpes+Simplex+Virus+Type-1+Based+Amplicon+Vectors.">Silencing Status Epilepticus-Induced BDNF Expression with Herpes Simplex Virus Type-1 Based Amplicon Vectors.</a> PLoS One. 11 (3), 1-17 (2016).</li><li>Munoz, J. L., Coles, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quartz+micropipettes+for+intracellular+voltage+microelectrodes+and+ion-selective+microelectrodes.">Quartz micropipettes for intracellular voltage microelectrodes and ion-selective microelectrodes.</a> J Neurosci Meth. 22 (1), 57-64 (1987).</li><li>Kilkenny, C., Browne, W., Cuthill, I. C., Emerson, M., Altman, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+research:+reporting+in+vivo+experiments:+the+ARRIVE+guidelines.">Animal research: reporting in vivo experiments: the ARRIVE guidelines.</a> J Cereb Blood Flow Metab. 31, 991-993 (2011).</li><li>Torres, E. M., Trigano, M., Dunnett, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Translation+of+cell+therapies+to+the+clinic:+characteristics+of+cell+suspensions+in+large-diameter+injection+cannulae.">Translation of cell therapies to the clinic: characteristics of cell suspensions in large-diameter injection cannulae.</a> Cell Transplant. 24, 737-749 (2015).</li></ol>

The authors have nothing to disclose.

The technique described in this article fulfills the needs outlined in the Introduction for optimizing microinjections that are performed for various purposes12. The needles described here reduce damage to a minimum, essentially non-detectable level; at variance with glass needles (that are prone to bend), quartz needles are rigid and ensure a reliable hit of the desired brain region even in deep coordinates. Moreover, the side hole ensures delivery of the solution even if the tip...

Microinjections have been used for a long time for the delivery of pharmacologically active compounds to modulate neuronal activity in specific brain areas. In addition, they have been used to inject toxins near particular neuronal populations, to mimic neurodegenerative events characteristic of certain diseases, for example 6-hydroxy-dopamine in the nigrostriatal dopamine system to mimic Parkinson&#39;s disease1,2 or the immunotoxin 192 IgG saporin to lesion the cholinergic system3. More recently, microinjection procedures have been used to deliver viral vectors or cells grafts for gene or cell therapy of experimental brain disorders4,5.
The classical type of needles employed in these studies is made of stainless steel. Although easy and practical to use, steel needles have a number of problems6: they are relatively large and may cause tissue damage, with leakage of the blood-brain barrier and activation of astrocytes; moreover, they may produce coring of brain tissue that gets into the needle creating an obstacle or even completely avoiding flow of the desired solution. More recently, glass needles prepared ad hoc from capillaries have been introduced in use7,8. These do not cause significant tissue damage nor astrocyte activation, but are relatively flexible and may bend when introduced in deep structures, reducing the accuracy of localization (personal observations).
There is therefore a need to reduce as much as possible damage (especially when performing experiments to heal damage) while increasing accuracy and reproducibility (i.e., ensure that all solution is delivered and ensure correct localization). Moreover, it would be desirable to use different needle designs to ensure optimal distribution of the injected solution in brain areas with varying geometries. In this article, we describe a protocol to prepare and use quartz needles for microinjections in the rodent brain. Due to the high melting point, quartz capillaries cannot be pulled on a conventional puller and therefore, have not been used in the past to generate needles. Quartz, however, offers some important advantages over glass, in particular high rigidity and break resistance9. Because of their rigidity, quartz needles are ideally suited for injections into ventral brain regions. Because of their high resistance to breakage they can be modeled to include multiple holes, obtaining designs that may prove most effective even when targeting brain regions with complex geometries10.

<table><tbody><tr><td>Quartz capillaries</td><td>Sutter Instruments, Novato, CA USA</td><td>Q100-50-10</td><td>Without filament</td></tr><tr><td>Puller</td><td>Sutter</td><td>P2000</td><td></td></tr><tr><td>Micropipette storage jar</td><td>World Precision Instruments (WPI), Sarasota, FL, USA</td><td>E210</td><td></td></tr><tr><td>Laser microdissector</td><td>Leica Microsystems, Wetzlar, Germany</td><td>LMD6500</td><td></td></tr><tr><td>Hamilton syringe</td><td>Hamilton ILS Innovative Labor Systeme GmbH, St&amp;Yuml;tzerbach, Germany</td><td>19138-U</td><td></td></tr><tr><td>Microinfusion pump</td><td>Univentor, Zejtun, Malta</td><td>Model 864</td><td></td></tr><tr><td>Manual microinjection pump kit</td><td>WPI</td><td>Item#: MMP-KIT</td><td>Kit allowing for micropipettes to be securely mounted to the stereotactic frame</td></tr><tr><td>Precision Drill</td><td>Proxxon</td><td>28510 MicroMot 50/E</td><td>Ball bearing drive shaft with variable speed</td></tr><tr><td>Artficial Cerebral Spinal Fluid</td><td>Tocris</td><td>3525</td><td></td></tr><tr><td>Needles 26 G Blunt and 30 G Bevel</td><td>Hamilton</td><td>26 G Blunt: 19138-U&lt;br/&gt; 30 G Bevel: 20757</td><td></td></tr><tr><td>Microtome</td><td>Leica, Germany</td><td>LEICA RM212RT</td><td></td></tr></tbody></table>

personalized needles for microinjections in the rodent brain

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to deliver gene or cell therapy products. Unfortunately, current microinjection techniques use steel or glass needles that are suboptimal for multiple reasons: in particular, steel needles may cause tissue damage, and glass needles may bend when lowered deeply into the brain, missing the target region. In this article, we describe a protocol to prepare and use quartz needles that combine a number of useful features. These needles do not produce detectable tissue damage and, being very rigid, ensure reliable delivery in the desired brain region even when using deep coordinates. Moreover, it is possible to personalize the design of the needle by making multiple holes of the desired diameter. Multiple holes facilitate the injection of large amounts of solution within a larger area, whereas large holes facilitate the injection of cells. In addition, these quartz needles can be cleaned and re-used, such that the procedure becomes cost-effective.

We compared damage induced by direct microinjection in the rat dorsal hippocampus and striatum using a quartz needle (60 &#181;M external diameter tip; one 20 &#181;M diameter side hole; type C, Figure 1) compared with two classic, 26 G blunt end and 30 G bevel edge stainless steel needles. To this aim, we injected 2 &#181;L of aCSF in the right and left dorsal hippocampus and striatum using respectively the quartz and the steel needle and, 48 h after injecti...

Watch this Scientific Journal Video about Personalized Needles for Microinjections in the Rodent Brain at JoVE.com

Personalized Needles for Microinjections in the Rodent Brain

We describe here a protocol for microinjection in the rodent brain that uses quartz needles. These needles do not produce detectable tissue damage and ensure reliable delivery even in deep regions. Moreover, they can be adapted to research needs by personalized designs and can be re-used.

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to ...

personalized-needles-for-microinjections-in-the-rodent-brain

Research

JoVE Journal

Neuroscience

11.2K Views.  University of Ferrara. Hier beschrijven we een protocol voor microinjection in de knaagdier hersenen die gebruikmaakt van kwarts naalden. Deze naalden niet produceren detecteerbare weefselschade en zorgen voor betrouwbare levering zelfs in diepe regio's. Bovendien, zij kunnen worden aangepast aan de behoefte aan onderzoek door gepersonaliseerde designs en opnieuw kunnen worden gebruikt.

Video: Personalized Needles for Microinjections in the Rodent Brain

Scalable Fluidic Injector Arrays for Viral Targeting of Intact 3-D Brain Circuits

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted in neurological and psychiatric disorders--would be greatly facilitated by a technology for rapidly targeting genes to complex 3-dimensional neural circuits, enabling fast creation of "circuit-level transgenics." We have recently developed methods in which viruses encoding for light-sensitive proteins can sensitize specific cell types to millisecond-timescale activation and silencing in the intact brain. We here present the design and implementation of an injector array capable of delivering viruses (or other fluids) to dozens of defined points within the 3-dimensional structure of the brain (Figure. 1A, 1B). The injector array comprises one or more displacement pumps that each drive a set of syringes, each of which feeds into a polyimide/fused-silica capillary via a high-pressure-tolerant connector. The capillaries are sized, and then inserted into, desired locations specified by custom-milling a stereotactic positioning board, thus allowing viruses or other reagents to be delivered to the desired set of brain regions. To use the device, the surgeon first fills the fluidic subsystem entirely with oil, backfills the capillaries with the virus, inserts the device into the brain, and infuses reagents slowly (&lt;0.1 microliters/min). The parallel nature of the injector array facilitates rapid, accurate, and robust labeling of entire neural circuits with viral payloads such as optical sensitizers to enable light-activation and silencing of defined brain circuits. Along with other technologies, such as optical fiber arrays for light delivery to desired sets of brain regions, we hope to create a toolbox that enables the systematic probing of causal neural functions in the intact brain. This technology may not only open up such systematic approaches to circuit-focused neuroscience in mammals, and facilitate labeling of brain regions in large animals such as non-human primates, but may also open up a clinical translational path for cell-specific optical control prosthetics, whose precision may enable improved treatment of intractable brain disorders. Finally, such devices as described here may facilitate precisely-timed fluidic delivery of other payloads, such as stem cells and pharmacological agents, to 3-dimensional structures, in an easily user-customizable fashion.

Controlling and analyzing neural circuits in vivo would be facilitated by a technology for delivery of viruses and other reagents to desired 3-dimensional sets of brain regions. We demonstrate customized fluidic injector array fabrication, and delivery of virally-encoded optical sensitizers, enabling optical manipulation of complex brain circuits.

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted ...

Targeting of Deep Brain Structures with Microinjections for Delivery of Drugs, Viral Vectors, or Cell Transplants

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an injecting needle produces during its trajectory is a major concern especially in the mouse brain for not only the brain is small but also sometimes multiple injections are needed.  We show here a method to produce glass capillary needles with a 50-&mu;m lumen which significantly reduces the brain damage and allows a precise targeting into the rodent brain. This method allows a delivery of small volumes (from 20 to 100 nl), reduces bleeding risks, and minimizes passive diffusion of drugs into the brain parenchyma. By using different size of capillary glass tubes, or changing the needle lumen, several types of substances and cells can be injected. Microinjections with a glass capillary tube represent a significant improvement in injection techniques and deep brain targeting with minimal collateral damage in small rodents.

In this article, we show a method to make glass capillary needles with a 50-&mu;m lumen. This technique significantly reduces the brain damage, minimizes passive diffusion of drugs and allows a precise targeting into the rodent brain.

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an ...

Rodent Brain Microinjection to Study Molecular Substrates of Motivated Behavior

Brain microinjection can aid elucidation of the molecular substrates of complex behaviors, such as motivation. For this purpose rodents can serve as appropriate models, partly because the response to behaviorally relevant stimuli and the circuitry parsing stimulus-action outcomes is astonishingly similar between humans and rodents. In studying molecular substrates of complex behaviors, the microinjection of reagents that modify, augment, or silence specific systems is an invaluable technique. However, it is crucial that the microinjection site is precisely targeted in order to aid interpretation of the results. We present a method for the manufacture of surgical implements and microinjection needles that enables accurate microinjection and unlimited customizability with minimal cost. Importantly, this technique can be successfully completed in awake rodents if conducted in conjunction with other JoVE articles that covered requisite surgical procedures. Additionally, there are many behavioral paradigms that are well suited for measuring motivation. The progressive ratio is a commonly used method that quantifies the efficacy of a reinforcer to maintain responding despite an (often exponentially) increasing work requirement. This assay is sensitive to reinforcer magnitude and pharmacological manipulations, which allows reinforcing efficacy and/ or motivation to be determined. We also present a straightforward approach to program operant software to accommodate a progressive ratio reinforcement schedule.

Rodents are an appropriate model to investigate the molecular substrates of behavior and complex psychiatric disorders. Brain microinjection in awake rodents can be used to elucidate disease substrates. An efficient and customizable brain microinjection method as well as the execution of an operant paradigm that quantifies motivation is presented.

Brain microinjection can aid elucidation of the molecular substrates of complex behaviors, such as motivation. For this purpose rodents can serve as ...

Behavior

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Syringe-injectable Mesh Electronics for Stable Chronic Rodent Electrophysiology

Implantable brain electrophysiology probes are valuable tools in neuroscience due to their ability to record neural activity with high spatiotemporal resolution from shallow and deep brain regions. Their use has been hindered, however, by mechanical and structural mismatches between the probes and brain tissue that commonly lead to micromotion and gliosis with resulting signal instability in chronic recording experiments. In contrast, following the implantation of ultraflexible mesh electronics via syringe injection, the mesh probes form a seamless, gliosis-free interface with the surrounding brain tissue that enables stable tracking of individual neurons on at least a year timescale. This protocol details the key steps in a typical mouse neural recording experiment using syringe-injectable mesh electronics, including the fabrication of mesh electronics in a standard photolithography-based process possible at many universities, loading mesh electronics into standard capillary needles, stereotaxic injection in vivo, connection of the mesh input/output to standard instrumentation interfaces, restrained or freely moving recording sessions, and histological sectioning of brain tissue containing mesh electronics. Representative neural recordings and histology data are presented. Investigators familiar with this protocol will have the knowledge necessary to incorporate mesh electronics into their own experiments and take advantage of the unique opportunities afforded by long-term stable neural interfacing, such as studies of aging processes, brain development, and the pathogenesis of brain disease.

Mesh electronics probes seamlessly integrate and provide stable, long-term, single-neuron level recording within the brain. This protocol uses mesh electronics for in vivo experiments, involving the fabrication of mesh electronics, loading into needles, stereotaxic injection, input/output interfacing, recording experiments, and histology of tissue containing mesh probes.

Implantable brain electrophysiology probes are valuable tools in neuroscience due to their ability to record neural activity with high spatiotemporal ...

Bioengineering

Manipulation of Single Neural Stem Cells and Neurons in Brain Slices using Robotic Microinjection

A central question in developmental neurobiology is how neural stem and progenitor cells form the brain. To answer this question, one needs to label, manipulate, and follow single cells in the brain tissue with high resolution over time. This task is extremely challenging due to the complexity of tissues in the brain. We have recently developed a robot, that guide a microinjection needle into brain tissue upon utilizing images acquired from a microscope to deliver femtoliter volumes of solution into single cells. The robotic operation increases resulting an overall yield that is an order of magnitude greater than manual microinjection and allows for precise labeling and flexible manipulation of single cells in living tissue. With this, one can microinject hundreds of cells within a single organotypic slice. This article demonstrates the use of the microinjection robot for automated microinjection of neural progenitor cells and neurons in the brain tissue slices. More broadly, it can be used on any epithelial tissue featuring a surface that can be reached by the pipette. Once set up, the microinjection robot can execute 15 or more microinjections per minute. The microinjection robot because of its throughput and versality will make microinjection a broadly straightforward high-performance cell manipulation technique to be used in bioengineering, biotechnology, and biophysics for performing single-cell analyses in organotypic brain slices.

This protocol demonstrates the use of a robotic platform for microinjection into single neural stem cells and neurons in brain slices. This technique is versatile and offers a method of tracking cells in tissue with high spatial resolution.

A central question in developmental neurobiology is how neural stem and progenitor cells form the brain. To answer this question, one needs to label, ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

Performing Intracerebroventricular Injection in Mice Using a Free-Hand Approach

Free-Hand Intracerebroventricular Injections in Mice

The investigation of neuroendocrine systems often requires the delivery of drugs, viruses, or other experimental agents directly into the brains of mice. An intracerebroventricular (ICV) injection allows the widespread delivery of the experimental agent throughout the brain (particularly in the structures near the ventricles). Here, methods for making free-hand ICV injections in adult mice are described. By using visual and tactile landmarks on the heads of mice, injections into the lateral ventricles can be made rapidly and reliably. The injections are made with a glass syringe held in the experimenter&#39;s hand and placed at approximate distances from the landmarks. Thus, this technique does not require a stereotaxic frame. Furthermore, this technique requires only brief isoflurane anesthesia, which permits the subsequent assessment of mouse behavior and/or physiology in awake, freely behaving mice. Free-hand ICV injection is a powerful tool for the efficient delivery of experimental agents into the brains of living mice and can be combined with other techniques such as frequent blood sampling, neural circuit manipulation, or in vivo recording to investigate neuroendocrine processes.

Author Spotlight: Insights, Innovations, and Future Avenues in Reproductive Neurocontrol 

Here, a simple and rapid approach for performing intracerebroventricular injections in mice using a free-hand approach (that is, without a stereotaxic device) is described.

The investigation of neuroendocrine systems often requires the delivery of drugs, viruses, or other experimental agents directly into the brains of mice. ...

Optimizing Stereotaxic Microinjection Accuracy in Mice

Preliminary Validation of Stereotaxic Injection Coordinates via Cryosectioning

Stereotaxic injection of a specific brain region constitutes a fundamental experimental technique in basic neuroscience. Researchers commonly base their choice of stereotaxic injection parameters on mouse brain atlases or published materials that employed various populations/ages of mice and different stereotaxic equipment, necessitating further validation of the stereotaxic coordinate parameters. The efficacy of calcium imaging, chemogenetic, and optogenetic manipulations relies on the precise expression of reporter genes within the region of interest, often requiring several weeks of effort. Thus, it is a time-consuming task if the coordinates of the target brain region are not verified in advance. Using an appropriate dye instead of a virus and implementing cryosectioning, researchers can observe the injection site immediately following dye administration. This facilitates timely adjustments to coordinate parameters in cases where discrepancies exist between the actual injection site and the theoretical position. Such adjustments significantly enhance the accuracy of viral expression within the target region in subsequent experiments.

Author Spotlight: Accelerating Cognitive Impairment Research in Mice Through Stereotaxic Injection and Cost-Effective Dyes

The present protocol describes a practical strategy to expedite the verification step of stereotaxic injection coordinates before conducting viral tracing using dyes and frozen sections.

Stereotaxic injection of a specific brain region constitutes a fundamental experimental technique in basic neuroscience. Researchers commonly base their ...

Intracerebroventricular Injection in Mouse: An Ommaya Mediated Direct Drug Administration Method to Deliver Drugs to Cerebral Ventricles of Mouse Brain

Source: Law, V. et al. <a href="https://www.jove.com/v/62033/a-murine-ommaya-xenograft-model-to-study-direct-targeted-therapy">A Murine Ommaya Xenograft Model to Study Direct-Targeted Therapy of Leptomeningeal Disease.</a> J. Vis. Exp. (2021)
This video demonstrates ommaya mediated intracerebroventricular injection in a mouse model. The method enables localized delivery of potential drug candidates and evades the blood-brain barrier.

Personalized Needles for Microinjections in the Rodent Brain

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