Inner space – a review of microinjection

The manipulation of genetic material is perhaps the single greatest biological advance of the last century. Here we examine the past, present and future of the most important tool of the genetic revolution – microinjection

Single-cell microinjection is a powerful and versatile technique for introducing exogenous material into cells, or transferring cellular components between cells. It is normally performed using a fine glass capillary, under a specialised optical microscope setup incorporating a micromanipulator. Microinjection is frequently used in genetic engineering and transgenetics to insert genetic material into single cells. It is also used in basic genetics research, in the cloning of organisms, and in the study of cell biology and viruses. Common clinical applications include In Vitro Fertilisation (IVF) and Intracytoplasmic Sperm Injection (ICSI) to help achieve fertilisation for couples with severe male factor infertility. Other areas include ophthalmology procedures such as Retinal Pigment Epithelial (RPE) and Intra Occular (IO) injection

The technique of needle microinjection was first developed in the early 1900s. It has become an increasingly popular biological research method for studying living cell systems, being one of just a few viable ways to introduce non-genetic, large molecules into living cells. Microinjection is now routinely used to transpose cellular organelles, DNA and RNA, enzymes, structural proteins, metabolites, ions and antibodies, from test tubes into living cells. Other techniques such as vesicle fusion, scrape loading and electroporation have been introduced but using needles for cellular injection provides the most versatile option and the ability to be quantitative.

In the 1960s key biological studies focused on microinjection of organisms such as amoebae and mouse embryos. Following this, research expanded to investigate the use of microinjection to deliver organelles and molecules to other cells such as Paramecium, frog and mouse oocytes and eggs and mammalian cells of somatic origin, forming the basis of the future use of microinjection for the production of “transgenic” animals.

Early microinjection experiments with Xenopus oocytes studied gene expression relating to the molecular recognition pathways of oocyte maturation. Further exploitation of the technique identified receptors and ion channels of cloned genes from mammalian cells. In fact, oocytes as microinjection vehicles were instrumental in enabling expression cloning of numerous important neuronal receptors. Isolation or identification of desired clones became achievable by measuring newly expressed receptor activity, following injection of large pools of cDNAs from mammalian cells.

Today, microinjection is widely used for the study of different cell responses in a variety of systems. These include individual cell signalling responses in the regulation of metabolic pathways, second function of second messengers, the fate of injected proteins and cytoskeletal regulation, definition of apoptosis and survival signals and transformation of whole organisms such as nematodes.

Perhaps the most powerful aspect of microinjection is the ability to introduce several types of reagents into cells simultaneously, including DNA constructs, labelled dextran to mark the injected cells, antibodies, short interfering RNAs (siRNAs), and peptides. No other techniques currently available provide these capabilities.

Microinjection is a relatively complex and technically challenging method to perform, requiring specialist equipment and trained experimental technique. The work is done under a powerful microscope with a pipette holding the target cell in the field of view. The tip of a glass micropipette/injection needle (about 0.5 mm diameter), is injected through the membrane of the cell. The contents of the needle are delivered into the cytoplasm and the empty needle is gently removed.

Xenopus eggs and embryos have been widely used for microinjection studies in the fields of cell biology and developmental biology. Their use is cited in thousands of publications. Due to their extremely large size, microinjection into Xenopus cells is somewhat less demanding, requiring little practice before the researcher becomes proficient with the technique.

Due to the delicate nature of the technique, limiting the amount of vibration to the workstation is vital for effective, reproducible microinjection. It requires a suitable quiet location, away from sources of vibration (slamming doors, fridges, centrifuges etc.), fluctuating temperature, draughts and bright light. Select a sturdy table or bench with plenty of room around it, and ensure that the workstation components, including trailing wires and tubes, are not in contact with the floor or walls. If vibration is still detected when a micropipette is placed into the microscope field under high magnification, use of an anti-vibration table should be considered.

Microinjection applications are typically performed using an inverted microscope - the objective lens turret is located below a fixed stage, with the transmitted light source located above. The working distance of the condenser, measured from its bottom-most part to the stage, after the condenser has been adjusted correctly, is particularly important. To enable access and unobstructed movement of the micromanipulator headstages a “long” or “ultralong” working distance condenser should be used. Stage attachments, specimen holders etc. may need to be removed so they are not in the way. The microscope should have sufficiently good optics to be used for several hours without causing eye strain. A halogen light source with flexible fibre-optic arms is non-intrusive and provides a cold light that does not heat up the media in the culture dish. The addition of a camera and TV monitor should be considered and is essential if the system is to be used for teaching or demonstration purposes.

A micromanipulator is required to ensure precise positioning of the injection needle and to facilitate the delicate movements during the injection procedure. It is used to position and move the micropipette in proximity to the tissue to be manipulated or injected. The micromanipulator is securely mounted to the microscope using an appropriate adapter. Ultimately the success of any microinjection will depend significantly upon the stability of the micromanipulator mounting. For greatest precision, the micromanipulator should position the injector at approx a 45 degree angle to the injection dish.

A microinjector device provides the pressure needed to deliver the sample solution from the micropipette into cells. Depending upon the application being employed, different pressures may be required. Xenopus embryos are typically injected with volumes ranging from 5-50nl so this needs to be done very accurately and precisely. The microinjector can be a mechanically or electronically regulated air pressure device or a simple glass syringe where the plunger is screw controlled for precise adjustments of the pressure. There are various manufacturers of such equipment but one of the leading suppliers is Drummond Scientific.

The manual oocyte microinjection pipette has a non-rotating plunger that eliminates tip wobble, to allow more precise deposition and enables injection of 30nl or more with reproducible accuracy. It utilises capillary tips that must be heated until the glass becomes partially liquefied and then quickly stretched to form a very fine tip at the heated end. The tip of the pipette is about 0.5mm diameter and resembles an injection needle.

Using a manual microinjector clearly needs a very steady hand and considerable skill and patience. To facilitate complete control, automated microprocessor controlled instruments such as the Drummond Nanoject II Autoinjector have been specifically designed to perform ultra-delicate nanolitre injection procedures into cells, including oocytes and embryos.

The configuration of any microinjection system will depend upon the application for which it is used with the key differences being the needle size and pressures applied.

Needles are often prepared from borosilicate glass capillary tubing. To make an injection needle, the glass capillary tube is heated until molten in the middle and is stretched by weights to create a very thin, hour-glass shaped taper, about 30mm in length. The finely drawn out capillary is bent using fine forceps to identify the point where the needle resists bending. The forceps are used to very gently break the capillary at an angle at approximately the point of flexure. The ID at the tip of the needle should be around 10-30 microns. It is then completely filled with mineral oil to act as a non-compressible displacement medium. By using a hydraulic fluid instead of air in the syringe and tubing, an even greater level of control is attained.

For certain applications such as the injection of single cells into late-stage embryos, it may be desirable to inject volumes less than 2.3nl. In this case, it is necessary to use one of the gas-pressure regulated microinjectors that allow injection of extremely small volumes and permit continuous variation in injection volume. With these devices, it is necessary to calibrate the injection volume for different needles and for different pressure conditions, Suitable devices are supplied by Narishige Picospritzer, Harvard Apparatus, Sutter and others.

When using an automated microinjector, a foot pedal injection control is a valuable addition helping to keep the hands free and allow the user to pay constant attention to the cells.

Microinjection Applications There are many documented applications for the technique but some of the most popular uses are listed here.

Zygote pronuclear DNA microinjection The microinjection of DNA into the pronucleus of a newly-fertilized mammalian egg is now a common and highly efficient method for creating transgenic offspring. Pronuclear microinjection was first described in the mouse, but now many different transgenic animals have been created in this way.

Embryonic stem cell transfer into blastocyst Animals, usually mice, can be engineered with a specific gene function reduced or knocked out altogether by introducing genetically altered embryonic stem cells into the cavity of a blastocyst so that the stem cells contribute to the embryo. The resulting live animal is a chimera of both genotypes. Subsequent selective interbreeding of manipulated animals results in pure-bred gene“knock-outs” or “knock-downs” and can be used for subsequent gene function studies.

Somatic cell nuclear transfer The enucleation of an oocyte followed by the transplantation of a somatic cell is a method of producing genetically identical copies (clones) of the animal from which the donor cell was taken.

Intracytoplasmic Sperm Injection Intracytoplasmic sperm injection (ICSI) can be employed for veterinary in-vitro fertilization during rare species preservation or for any veterinary assisted conception. ICSI may also be used as a gene transfer technique when sperm are co-injected with exogenous DNA.

There is no doubt that microinjection is a valuable research procedure and the availability of affordable instrumentation makes it more accessible now than ever before. The development of tools and microanalytical techniques is likely to continue to make the process simpler and even more versatile. When combined with methods such as PCR the scope of possible microinjection experiments will further expand.

Author: Susan Mayfield is Product Manager for Liquid Handling at Alpha Laboratories t: 02380 483000  w:www.alphalabs.co.uk