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

Summary

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.

Abstract

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.

Introduction

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 embryo^2,3,4. Sharp microinjection needles are a critical tool in the process of genetic modification of some insects, such as mosquitoes^4,5,6,7,8,9 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 insect^2,4. 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 tip^2,3. 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.

Protocol

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

1. 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.
2. 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).
3. 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.
4. 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.
5. In the side port of the regulator, screw in a T connector (T) by hand, then use a small wrench to finish tightening.
6. 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.
7. 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.
8. 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.
9. 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.
10. Place a hose clamp onto the tubing section, then insert a hose connector as described in steps 1.2-1.4.
11. On the other side of this section of tubing, place the female luer connector (LC) that is supplied with the needle holder (NH).
12. 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.
13. 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.
14. 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).
15. Connect the female luer connector to the male luer connector (Figure 2, g) of the needle holder (Figure 2, d).
16. 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.

2. Beveling borosilicate needles

1. 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.
2. Assemble the grinding assembly according to the manufacturer'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.
3. Place 10 drops of pedestal oil onto the pedestal's optically flat surface and place the grinding assembly on top. Start the grinding assembly.
4. 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° 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.
5. 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.
6. 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'clock to 6 o'clock (with the plate as a clock face). Ensure the wick does not ride on the black portion of the retaining ring.
7. 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.
8. 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)
9. 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.
10. 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'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.
11. 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.
12. Allow the needle to stay at this level of caliper reading for 5-10 s.
13. 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.
14. 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.
15. 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.
16. 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.
17. 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.

3. Determining the relative opening size of the beveled needle

NOTE: Determining the beveled needle's relative opening sizes is done in step 2.13. The steps below describe this process further.

1. 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.
2. Increase the air pressure until bubbles are flowing again from the needle tip. The higher the pressure, the smaller the opening of the needle.
3. Proceed with moving the needle to a position where it can be safely removed steps 2.16-2.17.

Results

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 regains its shape. With enough pressure, the dull needle will eventually tear through the membrane, and embryoplasm will likely leak out. The sharper the needle, the less indentation occurs as the needle is pressed against the embryo membrane (Figure 4E). Instead of tearing into the membrane, a sharp needle will slip through the membrane, causing little to no damage to the embryo. With an ideal needle, there will be almost no indentation of the embryo membrane as the needle slips into the embryo.

The University of Maryland Insect Transformation Facility (UM-ITF) is a service facility that creates genetically modified insects for clients in academia and industry. The facility uses beveled needles for the microinjection of mosquito embryos. During the period between January 2023 and December 2023, the UM-ITF injected 12,795 Aedes embryos across 56 client projects (Table 1). The use of beveled needles helped the UM-ITF maintain an average post-injection hatch of 31% across projects (Table 2). Furthermore, the UM-ITF was able to maintain an injection success rate of 96%, as determined by post-project follow-up with clients (Table 2). Injection success is a measure of whether the injection materials (CRISPR guides, Cas9 protein, transposon vector plasmids, etc.) injected were delivered into the embryos of the project in a manner where these materials were able to create a genetic modification, basically, was there evidence of CRISPR Indel creation, insertion of the clients transposon-based vector or insertion of an injection quality control vector (transposon ends with a promotor plus marker). In other words, were the injections good enough to produce genetic modification, if injected components were working properly. There are many factors that contribute to the success of a genetic modification project, and quality microinjections with sharp needles are a key factor of success. Beveling needles using the procedure described in the above protocol enables the UM-ITF to produce quality sharp microinjection needles consistently, therefore allowing the UM-ITF to give consistently successful injections.

Figure 1: Schematic diagram of air system components and connection to the microcapillary. Air is delivered to the capillary needle through a system of urethane tubes from an air supply (either house air or air cylinder). The urethane tubes are connected to the regulator (R) and pressure gauge by hose connections (HC), which consist of a hose connector and a hose clamp. The hose connection screws into a 10-32 thread female connection on the base of the regulator (R) and the pressure gauge. A T connector (T) is connected to the regulator (R) to spilt air between the pressure gauge and the needle holder (NH). The urethane tubing is connected to the needle holder by a luer connector (LC) that is supplied with the needle holder (NH). The connection to the air supply will vary depending on the air supply used. Please click here to view a larger version of this figure.

Figure 2: Manipulator parts and air set up. Manipulator parts: a. coarse adjustment knob, b. fine adjustment knob, c. Caliper, d. WPI needle holder, e. angle gauge, f. retaining clamp and bicycle fender clip, g. luer adapter and luer hose connector, and h. urethane tubing. Please click here to view a larger version of this figure.

Figure 3: Needle placement in relation to the abrasive plate rotation. (A) Proper needle placement during beveling. The needle to be beveled should be placed onto the rotating abrasive plate oriented somewhere between perpendicular to the rotation and in line with the rotation, where rotation moves away from the tip of the needle. (B) Improper needle placement. The abrasive plate rotation in this orientation has the abrasive plate surface pushing onto the needle tip rather than under and past the tip. In this orientation, beveling may not occur smoothly. Please click here to view a larger version of this figure.

Figure 4: Comparison of needle insertion by non-sharp and sharp needles. (A-C) Non-Sharp Needle and (D-F) Sharp Needle. (A) and (D) were captured before the needle made contact with the embryo; (B) and (E) are at the point just before the needle pierces the embryo membrane, and (C) and (F) are just after the needle piercing. The white arrow in (B) indicates the embryo membrane indenting because greater pressure has to be applied in order for a non-sharp needle to pierce. The black arrow in (E) indicates the lack of indentation due to the needle piercing done easily because of a sharp needle. Please click here to view a larger version of this figure.

Table 1: UM-ITF Aedes projects from January 2023 to December 2023. Listing of UM-ITF Aedes microinjection projects with the number of embryos injected for each project, the number of larvae that hatched from these injected embryos, and the percentage of larvae that hatched per injected embryo total. Please click here to download this Table.

Table 2: The UM-ITF Aedes project results. The UM-ITF is a service facility that provides microinjection services for insect genetic modification. The table lists the number of Aedes projects injected by ITF from January 2023 to December 2023. The number of clients who responded to project follow-up requests in which the response indicated success or failure of the injection project. The number of successful projects, where a successful project was one in which the injection materials were microinjected at a time and location into the developing embryo such that the injections were capable of producing a genetic modification, and the number of projects which were considered not successful due to lack of genetic modification. The average hatch rate for Aedes microinjection projects during the January 2023 to December 2023 period. The project success rate and the client response rate. Please click here to download this Table.

Discussion

Genetic modification of mosquitoes relies on precise microinjection of the modification materials (plasmids, guide RNAs, or proteins) into pre-blastoderm embryos^3,4,5,6,7,8. Crucial to this process are sharp needles that easily pierce the embryo during injection^2,4. A sharp needle is able to slip through the membrane, deliver the modification materials, and be withdrawn from the embryo without having any of the embryo-plasm leak out at the injection site post-microinjection (personal observation).

Microinjection needles are closed when they are first pulled and need to be opened before injection². Traditionally, needles for microinjection have been opened by gently brushing the needle against something such as the edge of the coverslip or glass slide that holds the embryos, against the embryo itself, or some other material². However, opening the needle by brushing it against an object provides an inconsistent outcome (personal observation). Sometimes, the needle will break in a way that makes it sharp, while other times, the needle becomes very dull, which, as previously mentioned, can damage the embryos that are being injected, causing them not to be modified or, at worst, killing the embryo. Alternatively, needles can also be opened by inserting the needle into an embryo. This process can break open the needle, making it ready for injection. However, it also produces random results that occasionally create a very sharp needle but more often clog the needle with embryo material before it can be used for injection. Although beveling needles take extra time and effort, the process produces sharp needles more consistently.

Genetic modification of mosquitoes is also a time dependent process that requires the embryos to be microinjected before the developing embryo cellularizes^4,5,6,7,8. When opening needles by brushing the needle against something or by inserting the needle into an embryo, the time taken to open and find a sharp needle is taken when the embryos are already set up for injection and are continuing to develop. In the best case, the process of opening a needle and producing a sharp needle only takes a minute or two, but other times, it may take long enough that the eggs set up for that round of injection are too old by the time a good needle is made. The advantage of beveling is that it is done ahead of time, and needles are ready to use as soon as the embryos are set for injection, ensuring that the embryos are at the perfect developmental stage for injection.

Although the above protocol was developed for and uses the Sutter BV-10 beveler, the protocol can be modified for use with any beveler. The main factor is the beveling surface must be able to hold a small volume of liquid under which the needle is beveled.

Beveling microinjection needles while an air source provides compressed air to the back of the needle has two advantages. First, the air pressure helps to produce consistent needles. Consistently sharp needles are produced when the beveling process is started at the same supplied air pressure, and beveling is stopped when bubbles first appear. It is critical that in order to observe the initial formation of bubbles, the abrasive plate must be stopped momentarily so that the spinning of the abrasive plate does not mask initial bubble formation. If evidence of bubbles is only visible when the abrasive plate is in motion, the opening of the needle may be too large. It is critical that the pressure of the air being supplied is the same each time beveling is started, that beveling is stopped as soon as bubbles start to escape (visualized when the plate is not in motion), and that bubbles are not visible when the plate is in motion The second advantage is that once the needle is opened, the opening size can be measured relatively by decreasing the pressure of the supplied air until bubbles stop escaping from the needle tip. By noting the pressure at which bubbles stop flowing from the tip, a relative opening size can be inferred. The lower the pressure at which the bubbles stop flowing, the larger the opening in the needle tip, and the higher the pressure, the smaller the tip opening size. Knowing the relative opening size can be useful when using an injection mix that is more viscous and, therefore, more likely to clog the injection needle. Using this relative opening size measurement allows for the creation of needles that can deliver higher viscosity injection mixes. Of course, there are some trade-offs made when using needles with larger opening sizes. Specifically, injected embryos may be more likely to leak after injection, and survival rates for injected embryos may be lower.

Materials

List of materials used in this article

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