Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

The overall goal of this immunostaining protocol is to detect protein expression in the aphid embryos. Critical conditions are identified that increase tissue permeability and decrease background staining, both of which are long-standing problems when staining embryonic tissues of aphids. The main advantage of this technique is that it provides specific conditions for effective penetration of antibodies and reduction of background staining, which usually are not described in a standard immunostaining protocol.

We first had this idea of this measure when we were studying how germ cells are specified in a pea aphid a variety of insect model, for genomics and environmental studies. The laboratory strain of the pea aphid, a. pisum, was originally collected in central Taiwan, and has been reared on host plants for more than 300 generations.

To germinate, soak the seeds of the host plants in tap water for three to five days at room temperature. Refill the fresh water once per day. Grow ten germinating seeds in a small pot with soil in the growth chamber under a photo period of 16 hours light, eight hours dark, at 20 degrees Celsius.

About 10 days after growth begins, the height of the plants is usually more than eight centimeters. Keep each pot of plants within a one liter glass beaker and transfer adult aphids onto the plants using a paintbrush. Then, seal the beaker with an air-permeable cover, such as gauze mesh, to prevent aphids from escaping.

Incubate the aphids in growth chamber under a photo-period of 16 hours light, eight hours dark, at 20 degrees Celsius, watering each pot of plants once every day. Fill one well of a spot plate with about 500 microliters of 4 percent paraformaldehyde. Place the plate under a stereomicroscope at low magnification, and submerge an adult aphid into the paraformaldehyde for dissection.

Dissect the ovaries by holding the head and abdomen with one set of forceps, cutting open the dorsal cuticle of the abdomen and dragging the ovaries away from the abdominal cavity. Next, fix three pairs of ovaries in 1.5 milliliter tube containing one milliliter of paraformaldehyde at room temperature for 20 minutes, with mild shaking on a mixer. Discard the fixation buffer with a glass dropper, and then wash the ovaries with PBST three times for 10 minutes, with mild shaking.

To treat the ovaries, first incubate them with Proteinase K for 10 minutes with mild shaking to increase the permeability of the embryonic tissues. Following incubation, discard the Proteinase K solution, and then wash the ovaries with 700 microliters of glycine three times for five minutes. Next, wash the ovaries with PBST twice for 10 minutes.

Fix the ovaries again with fixation buffer for 15 minutes at room temperature with mild shaking. To suppress the engoenous peroxidase activity, serially dehydrate the ovaries with different percentages of methanol in 0.2 percent PBST with volume ratios of one to three, one to one, and three to one, and then further incubate the ovaries with 100 percent methanol for one hour at room temperature with mild shaking. Then, serially rehydrate the ovaries, as instructed in the text protocol.

For three pairs of ovaries in a 1.5 milliliter tube, 200 microliters is the minimum volume for sample blocking and antibody staining. Incubate the ovaries with the 1x DIG-B blocking solution for four hours at room temperature, or overnight at four degrees Celsius, with mild shaking. Following incubation, discard the supernatant and replace with fresh 1x DIG-B blocking solution, containing the primary antibody at the appropriate dilution ratio.

Stain the ovaries for four hours at room temperature, or overnight at four degrees Celsius with mild shaking. After washing the ovaries as described in the text protocol, discard the supernatant and replace with fresh 1x DIG-B blocking solution containing the secondary antibody at the appropriate dilution ratio, and stain the ovaries as before. For immunoflourescent staining, carry out staining in the dark, because the secondary antibody is light sensitive.

The thickness of aphid embryos varies between stages of development. Mounting strategies are thus modified to fit early, mid, and late embryos. Transfer the ovaries, together with mounting medium, to the cell tray with a plastic dropper, and observe samples under a stereomicroscope at low magnification.

Cut the calluses associated with the lateral oviduct using insect pins, and then transfer an isolated ovariole to an empty glass well with a dropper. Make up the final volume to between 50 and 100 microliters using mounting medium. Then, transfer an ovariole onto the slide with a glass dropper.

Dissect egg chambers using insect pins. For embryos older than stage six of development, separation of egg chambers is suggested. For germaria and the first two egg chambers, separation is optional.

Relocate a dissected egg chamber to a clean slide using a glass dropper. For embryos at stages one to 10 of development, slowly place a cover slip over the dissected germaria or egg chambers to avoid bubbles. Alternatively, for embryos at stage 11 to 18 of development, mount egg chambers on a slide with a one-sided cover slip bridge, and place another cover slip on top of the sample.

For embryos older than stage 19 of development, mount the egg chambers on a slide with a double-sided cover slip bridge, and place another cover slip on top of the sample. Next, fill the space beneath the top cover slip with mounting media to avoid drying the sample. Mildly roll the top cover slip to obtain the right orientation for observation.

Finally, seal around the edge of the cover slips, including the bridge cover slips, with nail polish before performing image analysis, as described in the text protocol. Shown here is a pair of ovaries dissected from a female adult. An ovariole is composed of one germarium, one to two O-sites, plus five to seven embryos, all of which are accommodated within egg chambers in an assembly line fashion.

After paraformaldehyde fixation, digestion of ovaries in Proteinase K solution at a concentration of one microgram per milliliter for 10 minutes is highly recommended. Staining results show that signal intensity is significantly increased in the Poteinase K digested embryos. Incubation of embryos in the blocking solution from a DIG-based buffer set reduces more background staining than that of embryos in normal goat serum and bovine serum albumin.

The endogenous activity of peroxidase can also result in high background staining. These studies reveal that submerging aphid embryos in methanol can suppress the peroxidase activity much more efficiently than treating embryos with hydrogen peroxide. The critical conditions revealed in this study can be applied for signal detection in both germ cells and somatic cells.

Once mastered, this technique can be done in two days if it is performed properly. After watching this video, you should have a good understanding of how to carry out successful immunostaining on aphid embryos, including elevation of signal intensity, and reduction of background staining. While attempting this procedure, it's important to remember to manipulate every embryo gently to get a good shape of embryos.

Don't forget that working with paraformaldehyde and DAPI can be extremely hazardous and precautions, such as gloves, should always be taken while performing this procedure. Following this procedure, other measures like in situ hybridization can be performed in order to answer additional questions, like whether messanger RNA is localized with its protein product in the embryonic cells. This method can help answer key questions in the field of insect environmental biology, such as how germ cells are specified, and how egg cells are determined in distinct reproductive phases in aphids.