The author would like to acknowledge the following people. The staff of the University of Maryland Insect Transformation Facility: Channa Aluvihare, Robert Alford, and Daniel Gay. Without their dedicated work, the Insect Transformation Facility would not exist. Vanessa Meldener-Harrell for proofreading this manuscript.

NOTE: The protocol as described below uses the Sutter BV-10 microcapillary beveled. However, this protocol can be modified for use with any model microcapillary beveled.
1. Assembly of regulator, pressure gauge, and air supply tubing
<ol>
	<li>Cut a section of urethane tubing for the connection from the air supply to the base of the regulator (R; Section 1, Figure 1). The length of this section will depend on the distance from the air supply to the regulator, which will be located next to the beveler.</li>
	<li>Slide a hose clamp onto the urethane tubing, then push a hose connector into the end of the tubing. Make sure the hose connector is fully inserted by placing the hose connector against a hard surface. While holding the tubing close to the hose connector, press the tubing firmly into the hose until the hose connector is fully inserted. Together, the hose clamp and hose connector form the connection HC (Figure 1).</li>
	<li>Slide the hose clamp up to the end of the tubing where the hose connector is inserted. Rotate the hose clamp over the inserted barb end so that it forms an airtight connection.</li>
	<li>Screw the hose connector into the base of the regulator (R) by hand, then finish tightening the connection with a small wrench. Make sure the connection is airtight but not overtight.</li>
	<li>In the side port of the regulator, screw in a T connector (T) by hand, then use a small wrench to finish tightening.</li>
	<li>Cut a small piece of urethane tubing (Figure 1, Section 2), approximately 3-5 cm in length. Place two hose clamps onto the piece of tubing, then insert a hose connector into each end of the tubing and finish the connections (HC) as described in steps 1.2-1.4 for each end of the tube.</li>
	<li>Screw one end of the short tube into the base of the pressure gauge, then finish tightening with a wrench as in step 1.5.</li>
	<li>Screw the other end of the short tube into one of the ports on the T connector (T) connected to the regulator in step 1.5.</li>
	<li>Cut a section of urethane tubing (Figure 1, Section 3) for the connection between the regulator and needle holder (NH). The size of this section will depend on the distance from the beveler to the location of the regulator.</li>
	<li>Place a hose clamp onto the tubing section, then insert a hose connector as described in steps 1.2-1.4.</li>
	<li>On the other side of this section of tubing, place the female luer connector (LC) that is supplied with the needle holder (NH).</li>
	<li>Remove the retaining clamp and the nylon washer (Figure 2, f) from the manipulator. The retaining clamp and washer are designed to hold a glass microcapillary only, they are not designed to hold the additional weight of a microcapillary holder, threaded rod, and air supply tubing.</li>
	<li>Cut a rectangular piece of rubber packing sheet 1.5 cm x 2 cm to wrap the threaded rod in the bicycle fender clamp. This will help hold the threaded rod and needle holder more securely in the bicycle fender clamp. Using two needle nose pliers, open the bicycle fender clip, fold the rectangular piece of rubber packing sheet over the threaded rod 3.5 cm from one end of the rod so that it forms a U over the rod. Insert the rubber sheet-covered threaded rod into the opened bicycle fender clamp and use the pliers to close the clamp around the rod and rubber sheet. Install the bicycle clamp and threaded rod assembly onto the manipulator bolt, replace the retaining clamp without the nylon washer, and tighten the retaining clamp until it securely holds the threaded rod assembly.</li>
	<li>Thread the needle holder onto the threaded rod. Make sure that the luer connector ends in a position that will not bind the urethane tubing when it is connected (Figure 2, g).</li>
	<li>Connect the female luer connector to the male luer connector (Figure 2, g) of the needle holder (Figure 2, d).</li>
	<li>Connect the free end of Figure 1, Section 1 tube to the air supply (this connection will vary based on the air supply used). The air supply should be clean, dry air, and free of oil residue. The air supply can be from a house air source or gas cylinder, either compressed air or Nitrogen.</li>
</ol>
2. Beveling borosilicate needles
<ol>
	<li>Pull microinjection needles using borosilicate glass micro capillaries with the following settings on the instrument: Heat: 305, Fil: 4, Vel: 70, Del: 235, Pul: 160, with Loop time of 12.24.</li>
	<li>Assemble the grinding assembly according to the manufacturer&#39;s instructions, consisting of the abrasive plate and the retaining ring with magnets12. 
	NOTE: The abrasive plate may need to be wrapped with a thin strip of transparent film to prevent premature leakage of Photo-Flo (wetting agent) from the abrasive plate. This is only needed if the wetting agent solution prematurely leaks down into the pedestal oil, causing the grinding plate to stop rotating. The wetting agent reduces the risk of drying marks on the glass needles after beveling. The use of a wetting agent is not critical to the beveling process, and water may be substituted.</li>
	<li>Place 10 drops of pedestal oil onto the pedestal&#39;s optically flat surface and place the grinding assembly on top. Start the grinding assembly.</li>
	<li>Turn on the light source to illuminate the surface. Ensure the light source is positioned behind the beveled and shines at an angle of 45&#176; to the abrasive plate and needle. The illumination angle is necessary so that a shadow of the needle is easily seen. At 90x magnification, rotate the microscope head in place. Briefly stop the abrasive plate spinning and focus the microscope on the surface of the abrasive plate.</li>
	<li>Add 1% wetting agent to the wick until the wick is completely wet. Add 1% wetting agent to the surface of the abrasive plate. Ensure the wetting agent covers the abrasive surface but does not leak on to the black retaining ring.</li>
	<li>Place the pre-wet wick onto the surface of the abrasive plate as it is spinning. Ensure the wick is on the left side of the abrasive plate (as you look down from the top) and stretches from 11 o&#39;clock to 6 o&#39;clock (with the plate as a clock face). Ensure the wick does not ride on the black portion of the retaining ring.</li>
	<li>Insert a needle into the needle holder (Figure 2, d) and tighten the retaining ring to hold the needle in place. Open the regulator (Figure 1, R) and increase the pressure to 24 psi.</li>
	<li>Raise the needle holder by rotating the course adjustment knob (Figure 2, a) Make sure the needle is raised high enough that it is higher than the surface of the rotating abrasive plate, then rotate the entire manipulator so that the needle swings into place above the rotating abrasive plate. The needle to be beveled should be placed onto the rotating abrasive plate oriented such that the rotation of the plate moves away from the tip of the needle (Figure 3A)</li>
	<li>Watching from the side, use the coarse adjustment knob (Figure 2, a) to lower the needle toward the abrasive plate surface. Stop when the needle is almost touching the surface of the liquid.</li>
	<li>Use the zoom to lower the magnification of the microscope, then move the microscope so that the needle is in the center of the field of view. Once in the center of view, increase the magnification, adjusting the manipulator&#39;s position so that the needle tip stays in the center of view. Once at maximum magnification, stop the grinding plate and focus the microscope on the surface of the abrasive plate, then immediately restart the rotation of the plate once the surface is in focus. The needle may not be in view at this point.</li>
	<li>Using the manipulator coarse adjustment knob, lower the needle toward the abrasive plate. In the field of view, an image of the needle and a shadow(s) of the needle will be visible. When the needle and the shadow(s) of the needle are close to touching, switch to the manipulator fine adjustment knob and continue to lower the needle until the needle and its shadow(s) appear to touch. At this point, read the caliper (Figure 2, c) and note the reading. The surface of the abrasive plate is at or below this caliper reading. 
	NOTE: It is difficult to see when the needle touches the surface of the rotating abrasive plate, so the needle may not actually be touching the abrasive plate at this point.</li>
	<li>Allow the needle to stay at this level of caliper reading for 5-10 s.</li>
	<li>Using the manipulator fine adjustment knob, raise the needle, making sure it stays underneath the surface of the wetting agent. Stop the rotation of the abrasive plate for a few seconds and observe whether bubbles escape from the needle tip. If bubbles are present, proceed to step 2.15. If bubbles are not present, proceed to step 2.14.</li>
	<li>Move the needle back to the caliper reading using the manipulator fine adjustment knob, then move it a little lower and take a new caliper reading, then repeat steps 2.12-2.13.</li>
	<li>Start the abrasive plate again to see if bubble formation is observable while the plate is rotating. If evidence of bubble formation is visible during plate rotation, the opening of the needle tip is likely too large for sensitive embryo microinjections, such as mosquito embryo injections. If bubble formation is not visible during abrasive plate rotation, then the needle opening is ideal for use in procedures requiring a sharp and small opening needle.</li>
	<li>Use the manipulator course adjustment knob to raise the needle above the abrasive plate to a position that is high enough above the plate that the needle will not hit anything as the entire manipulator is rotated to move the needle away from the abrasive plate and microscope.</li>
	<li>Once the needle is in a position where it can be removed without hitting anything, lower the air pressure to zero, remove the needle, and place it in a needle storage box (a Petri dish with either double stick tape or modeling clay to hold the beveled needles) until use.</li>
</ol>
3. Determining the relative opening size of the beveled needle
NOTE: Determining the beveled needle&#39;s relative opening sizes is done in step 2.13. The steps below describe this process further.
<ol>
	<li>Once bubbles are observed in step 2.13, with the abrasive plate not rotating, slowly decrease the air pressure by rotating the regulator adjustment knob (Figure 1, R) until bubbles stop flowing from the tip of the needle. Note the pressure at which bubbles stop flowing from the tip.</li>
	<li>Increase the air pressure until bubbles are flowing again from the needle tip. The higher the pressure, the smaller the opening of the needle.</li>
	<li>Proceed with moving the needle to a position where it can be safely removed steps 2.16-2.17.</li>
</ol>

Protocol for Wet Beveling of Microinjection Needles Under Constant Air Pressure

<ol>
	<li>Rubin, G. M., Spradling, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+transformation+of+Drosophila+with+transposable+Element+Vectors.">Genetic transformation of Drosophila with transposable Element Vectors.</a> Science. 218 (4570), 348-353 (1982).</li><li>O'Brochta, D. A., Atkinson, P. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transformation+systems+in+insects.">Transformation systems in insects.</a> Methods Mol Biol. 260, 227-254 (2004).</li><li>Handler, A. M., James, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Insect Transgenesis: Methods and Applications. , (2000).</li><li>Harrell, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosquito+embryo+microinjection.">Mosquito embryo microinjection.</a> Cold Spring Harbor Protocols. , (2023).</li><li>Allen, M. L., O'Brochta, D. A., Atkinson, P. W., Levesque, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable,+germ-line+transformation+of+Culex+Quinquefasciatus+(Diptera:+Culicidae).">Stable, germ-line transformation of Culex Quinquefasciatus (Diptera: Culicidae).</a> J Med Entomol. 38 (5), 701-710 (2001).</li><li>Grossman, G. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Germline+transformation+of+the+malaria+vector,+Anopheles+gambiae,+with+the+piggyBac+transposable+element.">Germline transformation of the malaria vector, Anopheles gambiae, with the piggyBac transposable element.</a> Insect Mol Biol. 10 (6), 597-604 (2001).</li><li>Adelman, Z. N., Jasinskiene, N., James, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+applications+of+transgenesis+in+the+yellow+fever+mosquito,+Aedes+aegypti.">Development and applications of transgenesis in the yellow fever mosquito, Aedes aegypti.</a> Mol Biochem Parasitol. 121 (1), 1-10 (2002).</li><li>Perera, O. P., Harrell, R. A., Handler, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Germ-line+transformation+of+the+South+American+malaria+vector,+Anopheles+albimanus,+with+a+piggyBac/EGFP+transposon+vector+is+routine+and+highly+efficient.">Germ-line transformation of the South American malaria vector, Anopheles albimanus, with a piggyBac/EGFP transposon vector is routine and highly efficient.</a> Insect Mol Biol. 11 (4), 291-297 (2002).</li><li>Harrell, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosquito+embryo+microinjection+under+halocarbon+oil+or+in+aqueous+solution.">Mosquito embryo microinjection under halocarbon oil or in aqueous solution.</a> Cold Spring Harb Protoc. , (2023).</li><li>Louradour, I., Ghosh, K., Inbar, E., Sacks, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9+mutagenesis+in+Phlebotomus+papatasi:+The+immune+deficiency+pathway+impacts+vector+competence+for+Leishmania+major.">CRISPR/Cas9 mutagenesis in Phlebotomus papatasi: The immune deficiency pathway impacts vector competence for Leishmania major.</a> mBio. 10 (4), e01941 (2019).</li><li>Tamura, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Germline+transformation+of+the+silkworm+Bombyx+mori+L.+using+a+piggyBac+transposon-derived+vector.">Germline transformation of the silkworm Bombyx mori L. using a piggyBac transposon-derived vector.</a> Nat Biotechnol. 18 (1), 81-84 (2000).</li><li>. BV-10 Micropipette Beveler Operation Manual Rev. 3.00 Available from: <a target="_blank" href="https://www.manualslib.com/manual/2073788/Sutter-Instrument-Bv-10.html">https://www.manualslib.com/manual/2073788/Sutter-Instrument-Bv-10.html</a> (2018)</li></ol>

The author has nothing to disclose.

Genetic modification of mosquitoes relies on precise microinjection of the modification materials (plasmids, guide RNAs, or proteins) into pre-blastoderm embryos3,4,5,6,7,8. Crucial to this process are sharp needles that easily pierce the embryo during injection2,4. A...

Insect genetic modification is a process originally developed in Drosophila by Rubin and Spradling, and over the years, this process has been modified to create genetic modifications in other species1. The process relies on the precise delivery of modification components microinjected into embryos at a specific window of time and location within the developing embryo2,3,4. Sharp microinjection needles are a critical tool in the process of genetic modification of some insects, such as mosquitoes4,5,6,7,8,9 and sand flies10 while not as critical for other insects, such as silkworms11. Sharp needles are often a key factor between success and failure when trying to create a genetically modified insect2,4. Typically, microcapillary glass needles are pulled by heating glass to the point where the glass becomes elastic, allowing the capillary to be pulled into a tapering closed tip needle. Before the needle can be used, it needs to be opened in a manner that creates a sharp tip for injection. Traditionally, needles are opened by brushing the needle tip gently against something (the edge of a slide/coverslip, or the embryo, etc.) that causes a small amount of glass to break off from the tip, randomly creating a sharp tip2,3. A slightly less random process is dry beveling, where the needle is quickly lowered onto an optically flat, spinning abrasive plate for a short period of time, causing a small amount of glass to be abraded from the needle tip, creating a sharp tip. Dry beveling is a little less random than brushing the needle tip against something. The protocol described below takes the beveling process a step further by supplying compressed air to the needle being beveled and beveling it under a liquid layer so that bubbles are visible as soon as the needle has been opened. This protocol details a method for producing reliably sharp microinjection needles. Beveling under a liquid layer is an improvement over randomly breaking the needle as described above, and dry beveling because the user receives feedback on the beveling process, allowing the person beveling to know when the needle is open and relatively how large the needle opening is. Knowing the relative opening size of the needle tip can allow the person beveling to create needles with different opening sizes. Various needle opening sizes have different advantages; for instance, larger needle opening sizes can accommodate injection mixes of high viscosity, while smaller opening needles cause less damage to the embryo being injected.
Air is supplied to the needle using a regulator and a system of urethane tubing based on a modified air-pressure regulated injection system2. While air is supplied to the needle at a constant pressure, the needle is beveled under a layer of liquid. The beveling process is comprised of a repeated five-step process: 1) lowering the needle to the abrasive surface, 2) allowing the needle to bevel for a short period of time, 3) raising the needle away from the abrasive plate while keeping it beneath the surface of the liquid layer, 4) stopping the spin of the abrasive plate to check for the appearance of bubbles. If no bubbles are present, then steps 1 through 4 are repeated until bubbles appear; 5) Once bubbles are present, the air pressure can be adjusted to determine the relative opening size of the needle. The lower the pressure needed to stop bubble formation, the larger the opening at the needle tip.

<table><tbody><tr><td>1.0 mm O.D. microcapillaries</td><td>World Precision Instruments</td><td></td><td></td></tr><tr><td>Beveler pedestal oil</td><td>Sutter Instruments</td><td>008</td><td></td></tr><tr><td>Bicycle fender clip</td><td>VeloOrange</td><td>R-clip 4-pack</td><td>https://velo-orange.com/products/vo-r-clip-4-pack</td></tr><tr><td>Boom Stand Microscope</td><td>AmScope</td><td></td><td>AMScope 3.5X-90X Trinocular LED Boom Stand Stereo Microscope or equivalent</td></tr><tr><td>BV-10 Beveler</td><td>Sutter Instruments</td><td>BV-10</td><td></td></tr><tr><td>Diamond abrasive plate&amp;nbsp;</td><td>Sutter Instruments</td><td>104F</td><td>Diamond abrasive plate - extra fine (0.2 &amp;micro; to 1.0 &amp;micro; tip sizes)</td></tr><tr><td>Gasket, Buna-N</td><td>Clippard Instrument Laboratory, Inc.</td><td>11761-2-pkg</td><td>Used to seal connection on T&amp;nbsp; or L connectors, if not already included with these pieces</td></tr><tr><td>Hose Clamp</td><td>Clippard Instrument Laboratory, Inc.</td><td>5000-2-pkg</td><td></td></tr><tr><td>Hose connector</td><td>Clippard Instrument Laboratory, Inc.</td><td>CT4-pkg</td><td>Need 5 hose connectors</td></tr><tr><td>Microinjection Needle Holder</td><td>World Precision Instruments</td><td>MPH3-10</td><td>Needle holder for 1mm outer diameter microcapillaries</td></tr><tr><td>P-2000</td><td>Sutter Instruments</td><td></td><td>Any needle puller</td></tr><tr><td>Photo-Flo 200 Solution</td><td>B&amp;amp;H Photo, Video and Audio</td><td>BH #KOPF200P&amp;nbsp; MFR #1464510</td><td>wetting agent</td></tr><tr><td>Pressure Gauge</td><td>Clippard Instrument Laboratory, Inc.</td><td>PG-100</td><td>0-100 psi gauge</td></tr><tr><td>Reference wick</td><td>Sutter Instruments</td><td>X050300</td><td></td></tr><tr><td>Reference wick holder</td><td>Sutter Instruments</td><td>M100019</td><td></td></tr><tr><td>Regulator</td><td>Clippard Instrument Laboratory, Inc.</td><td>01-Mar</td><td>Need #10-32 ports for connections</td></tr><tr><td>Rubber Packing Sheet 6 inx 6 in</td><td>Danco</td><td>Model # 59849</td><td></td></tr><tr><td>T fitting</td><td>Clippard Instrument Laboratory, Inc.</td><td>15002-2-pkg</td><td></td></tr><tr><td>Threaded Bar</td><td></td><td></td><td>Either a threaded rod or bar with threaded end. Threads must be 10-32.</td></tr><tr><td>Urethane tubing</td><td>Clippard Instrument Laboratory, Inc.</td><td>URH1-0804-BLT-050</td><td></td></tr></tbody></table>

wet beveling of microinjection needles utilizing constant air pressure for feedback on needle opening

Microinjection needles are a critical tool in the delivery of genome modification reagents, CRISPR components (guide RNAs, Cas9 protein, and donor template), and transposon system components (plasmids and transposase mRNA) into developing insect embryos. Sharp microinjection needles are particularly important during the delivery of these modifying agents since they help minimize damage to the embryo being injected, thereby increasing the survival of these embryos as compared to injection with non-beveled needles. Further, the beveling of needles produces needles that are more consistent from needle to needle as compared to needles opened by randomly breaking the needle tip by brushing the tip against an object (side of a coverslip, the surface of the embryo to be injected, etc.). The process of wet beveling of microinjection needles with constant pressure air delivered to the needle allows the person beveling the needle to know when the needle is open (presence of bubbles) and also gives a relative indication of how large a needle opening has been created. The relative opening size in the needle can be determined by adjusting the air pressure delivered to the needle until an equilibrium is reached and bubbles stop flowing from the tip of the needle. The lower the pressure at which the equilibrium is reached, the larger the needle size; and conversely, the higher the pressure, the smaller the needle size.

The procedure described above produces consistently sharp microinjection needles. Sharp needles are characterized by being able to insert into soft chorion insect embryos, such as mosquito embryos, with little to no resistance from the embryo membrane. When mosquito embryos are microinjected for genetic modification, the embryo membrane is relatively elastic. Pushing a dull needle against the embryo membrane will cause it to indent (Figure 4B). When the needle is pulled back, the membrane re...

Author Spotlight: Enhancing Microinjection Needle Quality by Wet Beveling

This protocol describes the assembly of a pneumatic system for the delivery of pressurized air to a needle during the process of needle beveling. The protocol further describes the beveling process for creating sharp microinjection needles and how to gauge the relative opening size of the needle.

Wet Beveling of Microinjection Needles Utilizing Constant Air Pressure for Feedback on Needle Opening

Microinjection needles are a critical tool in the delivery of genome modification reagents, CRISPR components (guide RNAs, Cas9 protein, and donor ...

wet-beveling-microinjection-needles-utilizing-constant-air-pressure

Research

JoVE Journal

Biology

302 Views.  University of Maryland. This protocol describes the assembly of a pneumatic system for the delivery of pressurized air to a needle during the process of needle beveling. The protocol further describes the beveling process for creating sharp microinjection needles and how to gauge the relative opening size of the needle.

Video: Author Spotlight: Enhancing Microinjection Needle Quality by Wet Beveling

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

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Developmental Biology

Real-time Measurement of Epithelial Barrier Permeability in Human Intestinal Organoids

Advances in 3D culture of intestinal tissues obtained through biopsy or generated from pluripotent stem cells via directed differentiation, have resulted in sophisticated in vitro models of the intestinal mucosa. Leveraging these emerging model systems will require adaptation of tools and techniques developed for 2D culture systems and animals. Here, we describe a technique for measuring epithelial barrier permeability in human intestinal organoids in real-time. This is accomplished by microinjection of fluorescently-labeled dextran and imaging on an inverted microscope fitted with epifluorescent filters. Real-time measurement of the barrier permeability in intestinal organoids facilitates the generation of high-resolution temporal data in human intestinal epithelial tissue, although this technique can also be applied to fixed timepoint imaging approaches. This protocol is readily adaptable for the measurement of epithelial barrier permeability following exposure to pharmacologic agents, bacterial products or toxins, or live microorganisms. With minor modifications, this protocol can also serve as a general primer on microinjection of intestinal organoids and users may choose to supplement this protocol with additional or alternative downstream applications following microinjection.

This protocol describes the measurement of epithelial barrier permeability in real-time following pharmacologic treatment in human intestinal organoids using fluorescent microscopy and live cell microscopy.

Advances in 3D culture of intestinal tissues obtained through biopsy or generated from pluripotent stem cells via directed differentiation, have resulted ...

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

Microiontophoresis and Micromanipulation for Intravital Fluorescence Imaging of the Microcirculation

Microiontophoresis entails passage of current through a micropipette tip to deliver a solute at a designated site within an experimental preparation. Microiontophoresis can simulate synaptic transmission1 by delivering neurotransmitters and neuropeptides onto neurons reproducibly2. Negligible volume (fluid) displacement avoids mechanical disturbance to the experimental preparation. Adapting these techniques to the microcirculation3 has enabled mechanisms of vasodilation and vasoconstriction to be studied at the microscopic level in vivo4,5. A key advantage of such localized delivery is enabling vasomotor responses to be studied at defined sites within a microvascular network without evoking systemic or reflexive changes in blood pressure and tissue blood flow, thereby revealing intrinsic properties of microvessels.
A limitation of microiontophoresis is that the precise concentration of agent delivered to the site of interest is difficult to ascertain6. Nevertheless, its release from the micropipette tip is proportional to the intensity and duration of the ejection current2,7, such that reproducible stimulus-response relationships can be readily determined under defined experimental conditions (described below). Additional factors affecting microiontophoretic delivery include solute concentration and its ionization in solution. The internal diameter of the micropipette tip should be &tilde; 1 &mu;m or less to minimize diffusional 'leak', which can be counteracted with a retaining current. Thus an outward (positive) current is used to eject a cation and a negative current used to retain it within the micropipette.
Fabrication of micropipettes is facilitated with sophisticated electronic pullers8. Micropipettes are pulled from glass capillary tubes containing a filament that 'wicks' solution into the tip of the micropipette when filled from the back end ("backfilled"). This is done by inserting a microcapillary tube connected to a syringe containing the solution of interest and ejecting the solution into the lumen of the micropipette. Micromanipulators enable desired placement of micropipettes within the experimental preparation. Micromanipulators mounted on a movable base can be positioned around the preparation according to the topography of microvascular networks (developed below). 
The present protocol demonstrates microiontophoresis of acetylcholine (ACh+ Cl-) onto an arteriole of the mouse cremaster muscle preparation (See associated protocol: JoVE ID#2874) to produce endothelium-dependent vasodilation. Stimulus delivery is synchronized with digitized image acquisition using an electronic trigger. The use of Cx40BAC-GCaMP2 transgenic mice9 enables visualization of intracellular calcium responses underlying vasodilation in arteriolar endothelial cells in the living microcirculation.

Microiontophoresis entails movement of ions from a micropipette in response to a difference in electrical potential between the inside and outside of the micropipette. Biologically active molecules are thereby delivered in proportion to electrical current. We illustrate acetylcholine microiontophoresis in conjunction with micromanipulation to study endothelium-dependent vasodilation in the microcirculation.

Microiontophoresis entails passage of current through a micropipette tip to deliver a solute at a designated site within an experimental  preparation. ...

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 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 ...

Neuroscience

Earthworm, Lumbricus Terrestris: A Novel Microinjection Vasculature In vivo Invertebrate Model

Although vertebrates are indispensable to biomedical research, studies are often limited by factors such as cost, lengthy internal review, and ethical considerations. We present the earthworm as an alternative, low-cost, invertebrate applicable to certain preliminary vasculature studies. Due to the surgical availability of the earthworm's dorsal vessels, ventral vessels, and five pairs of pseudo hearts, earthworms are readily accessible, offer low-cost maintenance, and require administration of only small doses of a given compound. The earthworm model provides a simple closed vascular circulatory system with a hemoglobin structure similar to human blood. A protocol is provided for anaesthetizing the earthworms and performing surgical incisions to expose relevant blood vessels. Micropipettes for compound administration are formed by heating and pulling glass with a pipette puller and using a beveling system to create a micron-scale fine needle tip. The tips are then used with a micropositioner and microinjector to inject arbitrary compounds into the vascular system of an earthworm, repeatably, with the availability of large sample sizes and small compound volumes. Details on the intricacies of injection procedure are provided. The small vessel size of the earthworm is challenging, particularly in the case of the ventral vessel; however, mastery of the techniques presented offers high repeatability as a low-cost solution, making studies of very large sample size practical.

Earthworms are a novel invertebrate in vivo bench-top model for vasculature studies. We present techniques and equipment that allow efficient surgery and microinjection into the earthworm vasculature. Surgical protocols, microinjection techniques and the procedure for producing custom-made micropipettes are described.

Although vertebrates are indispensable to biomedical research, studies are often limited by factors such as cost, lengthy internal review, and ethical ...

Bioengineering

Microinjectrode System for Combined Drug Infusion and Electrophysiology

This microinjectrode system is designed for drug infusion, electrophysiology, and delivery and retrieval of experimental probes, such as microelectrodes and nanosensors, optimized for repeated use in awake, behaving animals. The microinjectrode system can be configured for multiple purposes: (1) simple arrangement of the cannula for placement of an experimental probe that would otherwise be too fragile to penetrate the dura mater, (2) microfluidic infusion of a drug, either independently or coupled to a cannula containing an experimental probe (i.e., microelectrode, nanosensor). In this protocol we explain the step by step construction of the microinjectrode, its coupling to microfluidic components, and the protocol for use of the system in vivo. The microfluidic components of this system allow for delivery of volumes on the nanoliter scale, with minimal penetration damage. Drug infusion can be performed independently or simultaneously with experimental probes such as microelectrodes or nanosensors in an awake, behaving animal. Applications of this system range from measuring the effects of a drug on cortical electrical activity and behavior, to understanding the function of a specific region of cortex in the context of behavioral performance based on probe or nanosensor measurements. To demonstrate some of the capabilities of this system, we present an example of muscimol infusion for reversible inactivation of the frontal eye field (FEF) in rhesus macaque during a working memory task.

We present a microinjectrode system designed for electrophysiology and assisted delivery of experimental probes (i.e., nanosensors, microelectrodes), with optional drug infusion. Widely available microfluidic components are coupled to a cannula containing the probe. A step-by-step protocol for microinjectrode construction is included, with results during muscimol infusion in macaque cortex.

This microinjectrode system is designed for drug infusion, electrophysiology, and delivery and retrieval of experimental probes, such as microelectrodes ...

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.