Doubled Haploid Breeding in Cereals

Abstract

Doubled haploid (DH) production has become an important tool in plant breeding largely due to its capacity to produce completely homozygous plants in one generation. Not only are traits fixed for selection but the multiple generations of inbreeding required using traditional breeding methods are circumvented. A major concern of implementing doubled haploid breeding is production costs, which can be divided into two major categories: operational limitations, such as methods of labeling plants and biological limitations, such as the proportion of germinating embryos. Operational efficiencies have been improved and biological impediments reduced to make DH breeding cost effective. However, prior to implementing a DH breeding program, the breeder should consider factors such as the potential for linkage drag, types of crosses to be used and whether production resources are sufficient to produce the DH populations necessary for success. Doubled haploid technology can be integrated with marker-assisted breeding for greater efficiency and to craft the DH population for particular traits. The technology can also be used to accelerate development of germplasm with new genes of interest and to generate cytogenetic stocks. To date, hundreds of DH-derived cultivars have been developed worldwide. In Canada, as much as 30 % of the spring wheat hectarage has been sown to cultivars developed using DH technology. The future for DH breeding is promising because robust DH protocols are available for an ever-growing number of crops and future applications will see a closer integration with molecular-marker and gene-splicing technologies.

Appendix: Wheat Maize-Pollen Doubled Haploid Manual of Agriculture and Agri-Food Canada

1.1 Maize Pollen Donor Plants

Maize is planted each week to ensure a constant supply of pollen. Approximately 25–30 seeds are placed in a 10 cm paper towel lined plastic pot containing a 6:1 mixture of soil and turface (a soil amendment similar to cat litter) (Fig. 9.3a). From newly planted seeds, plants will take approximately 6–8 weeks before tasseling, so the planting of maize must be started before the planting of spring wheat parent plants. Once the seedlings are approximately 7–10 cm tall, they are transplanted individually into 15 cm clay pots containing the same soil mixture (Fig. 9.3b). The number of seedlings that are transplanted is dependent upon (1) the number of parental wheat plants, and (2) the number of spikes being emasculated. If 15 plants are transplanted per week, then at any given time there will be approximately 120 pots of maize at different stages of development. Once the maize is in clay pots the plants should be fertilized every week with a fertilizer solution of 20-20-20 (approx. 10 gL⁻¹) and 2 gL⁻¹ of ammonium sulphate (21-0-0). The plants at all stages are grown at 22 °C light/20 °C dark with a photoperiod of 16 h. It is important to have the lights turn on around 7:00 a.m. as pollen shed is affected by light and temperature. The maize pollen starts to shed between 1 and 2 h after the lights are illuminated. Before or shortly after the lights turn on, strip all the old anthers off the tassels. By removing all the old anthers, any new anthers that start to protrude are easily noticed and use of fresh pollen is ensured. As soon as the new anthers start to emerge, either shake the pollen from the tassels onto a large Petri dish or move the maize plants to the area where the pollination will be performed. Once the pollen starts to shed, the maize plants should not be moved as the pollen is easily lost from the anthers. When the maize plants start to silk, the period of pollen shed is nearly complete and the plants should be discarded although, some plants may be kept for seed production. In which case, tap the tassels of one plant onto the silks of the others. Repeat with several plants so that there is a high level of inter-crossing. Isolate the maize plants for seed production in a separate chamber because they tend to harbor mites and aphids which are difficult to control on maize and undesirable on pollen donor plants.

Fig. 9.3

(a). Maize 1 week after seeding 25–30 seeds in a 10 cm pot. (b) One-week-old maize seedlings transplanted into 15 cm clay pots

1.2 Wheat Parental Plants

Seed of wheat plants to be used to produce doubled haploid lines are started in 8 cm plastic pots containing Sunshine 5 (a soilless mix containing peat moss, perlite and vermiculite). Seed can be planted directly or germinated in dampened filter paper lined Petri dishes. Transplanting from the Petri dish occurs when the shoot is approximately 3–5 cm long. Once the potted plants have reached the two leaf stage they are transplanted into a 25 cm plastic pot (8 L pots commonly used by nurseries) containing a 6:1 mixture of soil and turface (Fig. 9.4). At this stage, the plants are watered with 20-20-20 and ammonium sulphate (21-0-0). It is important to grow large, healthy parent plants with many tillers. The plants are grown at a 16 °C light/15 °C dark with 16 h of light. They are fertilized daily with a siphon mixer of a ratio of 16:1 water to fertilizer (fertilizer 20-20-20, 200 g/20 L). Depending on the genotype of the wheat parent plant, 6–8 weeks is required for spike emergence. If aphids appear they should be spot controlled with a pyrethroid such as Raid especially in the later stages to minimize pesticide exposure of technical staff as the plants will be handled often. Other pesticides (e.g. Pirliss, Cygon, Malathion) and fungicides (e.g. Tilt) do not appear to interfere with doubled haploid production, but there are handling and safety issues associated with their use.

Fig. 9.4

Wheat doubled haploid parental plants with segregating gametes (e.g. F₁) grown in 8 L plastic containers

1.3 Crossing

1.3.1 Emasculation

The timing of emasculation is critical when pollinating wheat with maize . A floret approximately 2/3 of the way up the spike should be checked to determine if the spike is ready. If the anthers are starting to change from a light green/yellow color to a bright yellow color and the stigma is fluffy, the spike should be emasculated (Fig. 9.5), which is often the day before natural pollen shed would occur. If the spike has awns, they should be clipped fairly close to the glumes before starting to emasculate. Starting at the bottom of the spike, using fine tipped forceps, remove the central florets of each spikelet leaving only the two lateral florets (Fig. 9.5a). Remove the anthers from the lateral florets being careful not to damage the female structures (Fig. 9.5b). Continue this procedure up to the top of the spike. Remove the very top spikelet and if any of the spikelets near the top or bottom of the spike are too small to work with, they can also be removed. Finish emasculating the other side of the spike. Once the emasculation of the spike is complete, it should be bagged with a glassine bag and closed with a paper clip (Fig. 9.4). Tag the spike with a crossing tag marked with the cross and the date of emasculation. Place the marked tag around the stem at the first internode. A second tag of a different color is placed on the tiller at the second internode. This is an indicator tag and nothing is to be marked on it. It merely alerts the person crossing that something must be done to that particular spike, i.e. pollination or spraying with dicamba.

Fig. 9.5

Spike preparation and emasculation. (a) Removal of center floret with forceps. (b) Removal of wheat anthers with forceps. (c) The yellow color of the anthers with a slight green tinge indicating the appropriate time for emasculation

1.3.2 Pollination

Being careful not to disturb other anthers, select anthers from the maize plants when they are starting to emerge (Fig. 9.6). The end of the anther should look like it is beginning to open. Remove several anthers from the maize plant. Remove the glassine bag from the emasculated wheat spike to be pollinated (the day after emasculation) and beginning with the bottom of the spike, carefully open the floret. Using tweezers, take a maize anther at its bottom end (opposite of the open end), tip the anther over the open floret and the pollen should flow easily from the anther to the stigma in the floret. One maize anther may be used for several florets. Alternatively, if maize pollen was collected in a Petri dish, use a soft brush to transfer the pollen from the dish to stigmas of the wheat floret. Once the entire spike is pollinated, mark the date of pollination on the crossing tag and replace the bag over the spike. The crossing bag may be left on the spike until the time of excision. The pollinated wheat plants are returned the growth cabinet. Ensure the lights in the growth cabinet are at a sufficient distance from the wheat spikes so the temperature is not elevated in the crossing bags.

Fig. 9.6

Maize anthers ready for pollination

1.4 Hormone Spray

The day following pollination, the spikes should be sprayed with a dicamba solution (100 mgL⁻¹). Remove the bag from the spike, and using a small hand-held sprayer, (a pump or a pressurized sprayer) apply the dicamba liberally to the spike, until a run off of liquid is seen (about 2 passes with the pressurized sprayer) (Fig. 9.7). Replace the crossing bag. Mark the tag to indicate the spike was treated. Remove the indicator tag as nothing else needs to be done to the spike until the time of excision.

Fig. 9.7

Spray application of dicamba

It is important to keep the parent plants with the crossed spikes under the same conditions as each other (i.e. 15 °C dark/16 °C light, 16 h light) as differing temperatures will affect the timing of excision. If warmer conditions were used the spike would be ready to excise at an earlier date.

The crossing procedure and the hormone spray is a three day process. In order to maximize production, lab work could be scheduled for 7 days a week. If this is not possible there are some modifications to the procedure to allow for a 5-day work week schedule. These include:

• 1. Reduce the temperature of the cabinet to 9 °C day and night on Friday. This will slow down the development of the plants over the weekend. Spikes which were emasculated on Friday are pollinated on Monday. Spikes that were pollinated on Friday receive the dicamba treatment on Monday.

• 2. Return the cabinet to normal operating temperature on Monday morning.

• 3. Slight adjustment of the timing of excision is required. Spikes that were pollinated on Monday and Tuesday are excised at 18 days post pollination.

The timing of the application of dicamba is critical to this procedure. Dicamba treatment approximately 24 h after pollination produces the best results in terms of the number of embryos recovered per spike.

1.5 Excision and Embryo Rescue

Culturing of the embryos should be done under sterile conditions in a laminar flow hood. Equipment needed for excision in the laminar flow hood include:

• 1. Dissecting microscope and light source

• 2. Sterilized distilled water (for rinsing after bleach/ethanol)

• 3. 70 % ethanol solution (for sterilization)

• 4. 50 % bleach solution (for sterilization)

• 5. 95 % ethanol solution (for wiping down surfaces in the hood)

• 6. 90 % alcohol for instrument sterilization

• 7. 3 vials for sterilization of caryopses (1 vial for sterilized water, 1 for 50 % bleach, 1 for 70 % ethanol solution) and 1 vial for 90 % ethanol solution to disinfect scalpel and forceps during excision.

• 8. Beaker for waste from sterilizing caryopses

• 9. Pair of forceps

• 10. Scalpel

• 11. Cheesecloth for wiping down surfaces with alcohol

• 12. Sterilized containers of media

• 13. 10 mL syringes

At 16 days after pollination, cut off the appropriate spikes retaining approximately 15–20 cm of the stem with the spike and place the stems in a container of water until they are ready to be excised. Carefully spread the lemma and palea open with forceps and remove the caryopsis and place into the barrel of a 10 mL syringe. Repeat for all caryopses on the spike. Be careful not to squeeze the caryopsis as they are fluid filled and lack endosperm. Identify the cross in the syringe by affixing the crossing tag to the syringe with tape or by writing the corresponding number of the syringe onto the applicable crossing tag. If more than one spike from a plant has been harvested, they can be combined, however, be sure to keep different genotypes as well as those that have been treated differently, separate. Surface sterilize the caryopses by filling the syringe with the 70 % ethanol solution for 40 s (Fig. 9.8a). Discard the ethanol and rinse the caryopses with sterilized distilled water. Next, draw in the 50 % bleach solution (Javex, approx. 6 % sodium hypochlorite), hold for 60 s, discard and rinse twice with sterilized distilled water.

Fig. 9.8

(a) Sterilization of caryopses. (b) Wheat caryopsis with developing embryo

Place caryopses onto viewing field of dissecting microscope. Dip the forceps and the scalpel in the 90 % ethanol to sterilize and allow to dry momentarily. Using forceps and a scalpel, carefully open each caryopsis. This can be achieved in one of two ways: (1) At the embryo end, hold the caryopsis with the forceps, using the scalpel cut the brush end of the caryopsis. Tilt the open end of the caryopsis towards the microscope with the forceps; (2) Steady the caryopsis with the forceps or scalpel and use another set of forceps to grab and tear the seed coat about two-thirds of the way from the embryo end and pull back exposing the embryo. The embryo is a small white or somewhat translucent structure in the caryopsis. Because there is no solid endosperm present, the embryo is easy to see (Figs. 9.8b and 9.9a). Once the embryo is found, carefully touch the forceps to the embryo. The embryo will stick to the forceps and then it can be placed on the sterilized growth media. Open a vial of media in the flow hood, and touch the embryo to the media surface about half way up the slant. Keep the vials containing embryos from the different parental crosses separated. Only one embryo is placed in each vial (Fig. 9.9b).

Fig. 9.9

(a) Selfed wheat caryopsis with endosperm development (left) compared to a caryopsis with a haploid embryo and no endosperm development (right). (b) Vial containing developing haploid plantlet on culture media

Once excision is complete, the embryos are given a 3 day cold and darkness treatment in a fridge at 4 °C. After the cold treatment, the embryos are placed in the dark at room temperature for 2 days then placed under light banks for a 16-h light/8 h dark cycle at room temperature. A combination of fluorescent and incandescent light is used. The temperature under the lights is usually around 23–25 °C. Each week, all the vials under the light banks are checked to see if the embryos are growing and are ready for transplanting.

1.5.1 Second Cold Treatment

If the embryos fail to grow after 2 weeks under the light banks, they are given a second cold treatment. The dormant 2-week-old embryos are placed back in the fridge for 2 weeks. At this time the vials are placed back under the light banks (a separate set of light banks are used to regulate the flow of vials from the first light treatment to the fridge and back under the lights after a second cold treatment.) These vials are checked for growth periodically and returned to the main light banks when significant shoots and roots are visible. After 4 months, those embryos which have received a second cold treatment and were returned to under the light banks and still show no signs of growth are discarded.

1.6 Medium

The medium constituents are agar, sucrose and Gamborg’s B5 medium. From one liter of media, 120 one-oz Quorpak vials are filled with approximately 8 mL of medium. The medium is autoclaved in the vials and cooled on a slant.

1.7 Transplanting Haploid Plants

Once the embryos grow to a size of 2–4 cm, they are ready to be transplanted. Haploids are transplanted into 14 cm long plastic Conetainers (like those used in the forestry industry) containing Sunshine 5 soilless mix which has been loosely packed and watered.

To remove the haploid from the vial, use a blunt spoonula and dig around the root, freeing it completely, and gently tip the vial over and withdraw the plantlet on the instrument. Dip the root into water to completely remove the agar from the plantlet. A hole is punched in the Sunshine 5 medium and the plantlet is placed so that the crown is covered. Firm the mix around the plantlet.

The tray of Conetainers is subsequently set into a tray of water and covered with a plastic lid to provide a humid environment. The lid is left on for 3 days or until the haploids are established. These haploid plantlets are kept at 15 °C with a 16 h light/8 h dark cycle. It is important to keep the developing haploid plantlets cool so that tillering is encouraged.

1.8 Colchicine Treatment – Doubling

1.8.1 Plant Preparation

When the haploid plantlets are between the three- and five-leaf stage, they are ready for the colchicine treatment. In order to identify the plants when they are removed from the cones, the pot stake has a hole punched, and an elastic string (approximately 10 cm long) is fed through this hole and tied in a knot at the ends. The elastic is pulled around the base of the plant, the stake is then fed through the loop of the elastic pulled through to ensure a snug fit. This step can be done the day before the plants are to be colchicine treated.

When preparing the plants, about 1/3 of the leaf tissue is trimmed off (Fig. 9.10a). Next, the plants are removed from the cones and the soil is stripped from the roots. The roots are cut back leaving about 5 mm and given a quick rinse with tap water (Fig. 9.10b). It is important to prepare the plants first thing in the morning because the colchicine treatment process requires steps throughout the day.

Fig. 9.10

(a) Trimming leaves of haploids. (b) Trimming roots of haploids

1.8.2 Colchicine Treatment and Rinse

Once the plant’s leaves and roots have been trimmed they are placed in a beaker (a 250 ml beaker will hold about 100 plants) and are ready for the colchicine treatment (Fig. 9.11).

Fig. 9.11

Plants in container for colchicine treatment

When working with colchicine, it must be handled very carefully as it is highly toxic. It is recommended to wear two pairs of nitrile gloves. Six drops (from an eye dropper) of DMSO (dimethyl sulfoxide) is added to 60 ml of colchicine (1 drop DMSO/10 ml of colchicine). This solution is transferred to the beaker containing the trimmed haploids (the crowns of the plants should be submerged). An aerator (a fish tank air pump will do) with a rubber tube leading from it, is placed in the beaker. The beaker is placed in a plastic pail and covered with a lid. A hole is made in the lid to allow the air tube to pass through. The covered pail is placed in a fume hood with the sash ajar to allow only for the air tube to run from the beaker to the air pump. This step ensures that the procedure is exposed to as little light as possible as well as containment of any chemical which may percolate from the beaker. The plants are treated for 3½ h at room temperature. Pour the used colchicine from the beaker into a labelled container for use in a subsequent day’s treatment (may be used twice, no additional DMSO is required). Once the colchicine solution has been used twice it is discarded into a container designated for toxic waste disposal.

To rinse residual colchicine from the plants, place the aeration tube back into the beaker (in sink) along with a rubber hose leading from the faucet (a backflow valve should be installed on the faucet). Have the water running at a flow that is strong enough to allow the water to trickle out of the beaker. Adjust the cold and hot water taps so that the temperature of the rinse water is between 15–18 °C. Rinse for a minimum of 2 h.

1.8.3 Replanting Colchicine Treated Haploids

After rinsing, the colchicine treated haploid plants are replanted into 20 cm long Conetainers filled with Sunshine 5 medium and then moistened. This procedure is similar to the initial planting of the young haploids where the cones are watered, holes are punched into the surface of the soil medium to allow for replanting up to the crown and the soilless mix is gently firmed around the plant. The stakes are slid off the plant before transplanting and stuck into the edge of the cone. The trays are then covered with a plastic lid and set in tubs filled with water overnight to allow for sufficient water uptake by the Sunshine 5. The tops are taken off the trays 3 days after colchicine treatment. The colchicine treated plants are placed in a growth cabinet set at 15 °C with a cycle of 16 h of light and 8 h of darkness. They are fertilized daily with a 20-20-20 fertilizer solution (approx. 10 g L⁻¹ stock solution) siphoned into the watering line (diluted 16:1).

1.9 Solution Preparation

1.9.1 Dicamba 100 mgL⁻¹ (100 ppm)

Items Required

250 ml labelled brown glass bottle (with cap)	10 mL graduated cylinder
50 ml beaker	500 mL graduated cylinder
2 cm stir bar	Small weigh boat
25 mg dicamba	Stir plate
15 mL 95 % ethanol	Analytical balance
235 mL double distilled water

Preparation Instructions

Into a 50 mL beaker add the stir bar, 15 mL 95 % ethanol and 25 mg dicamba, stir to dissolve. Pour the dicamba solution into the 250 mL brown bottle, triple rinse the 50 mL beaker and add the rinseate and the remaining water to the 250 bottle.

1.9.2 Colchicine 0.2 % (w/v)

Items Required

500 mL labelled brown glass bottle (with lid)	500 mL distilled water
2 cm stir bar	500 mL graduated cylinder
1 g colchicine	Stir plate

Preparation Instructions

Prepare the colchicine in a fume hood being sure to wear a lab coat, eye protection and two pairs of nitrile gloves. Pour approximately 200 mL of distilled water into the 500 mL bottle along with the stir bar. Carefully pour the pre-measured 1 g container of colchicine into the 500 mL bottle, using the water in the beaker to triple rinse out the container. Pour the remaining water into the 500 mL bottle and cap. The colchicine can be stirred with the magnetic stirrer outside of the fume hood.

1.9.3 Embryo Culture Medium

Items Required

2 L beaker	0.1 M HCl
40 g sucrose	Weigh boats
2 L distilled water	Analytical balance
20 g agar	240 1-oz Quorpak vials
6.4 g Gamborg’s B5	Autoclavable boxes
5 cm stir bar	Pipette
Hot plate	500 mL graduated cylinder
pH meter	Thermometer
0.1 M NaOH

Preparation Instructions

Into a 2 L beaker place the stir bar and add 2 L of distilled water. Place the beaker on the hot plate and begin heating. Measure out the agar, sucrose and Gamborg’s B5 solids. Turn on the stir bar. Add the agar to the 2 L beaker of water. Continue heating until the temperature reaches 85–90 °C. Turn off the hot plate and stirrer and remove the beaker to cool it down. Once the temperature drops below 80 °C place the beaker back onto the stirrer and continue stirring while adding the sucrose and Gamborg’s B5 solids. Place the pH electrode in the medium and adjust the pH to 6.10 using the NaOH or HCl. Dispense the medium into vials. This can be done by pouring medium into a small beaker (400 mL size) and pouring into vials by hand (approximately 8 mL per vial). A faster and more accurate way is to use an automatic dispenser. Cap the vials, but not too tightly and place them in an autoclavable container. Autoclave the vials on the liquid cycle (15 psi, 121 °C) for 15 min. Cool the vials in the autoclavable containers on a slant overnight, then store under refrigeration.