Maize inbred B104: Seed can be obtained from the USDA Agricultural Research Service seed repository.¹

Plant material: Greenhouse-grown B104 embryo donor ears are harvested 10–12 days after pollination (depending upon the weather and day length) when immature zygotic embryos are 1.6–2.0 mm long. After harvest, maize ears (in their husks and inside their pollination bag) can be stored in a laboratory refrigerator (4°C) for 1–3 days prior to use.

Binary vector pKL2013 (McCaw et al., 2021; Figure 1A): The T-DNA carries (1) 2X CaMV 35S promoter (P35S) driving the bialaphos resistant (bar) gene as a selectable marker, (2) a single P35S driving mCherry gene as a visual marker; (3) maize ubiquitin promoter driving a maize-codon-optimized Cas9, and (4) rice U3 promoter driving a guide RNA (gRNA) targeting maize glossy2 (Gl2) gene (McCaw et al., 2021).

Ternary helper plasmid pKL2299 (this work; Figure 1B): It carries the majority suite of Agrobacterium virulence genes from Ti plasmid pTiBo542 (Komari et al., 1986). The additional copies of vir genes have been shown to markedly enhance maize transformation frequencies when using Agrobacterium strain LBA4404 (Anand et al., 2018). To ensure vector compatibility, pKL2299 has the RK2 origin of replication. Therefore, this plasmid can be used as a ternary vector pairing with any binary vector plasmid that has a pVS1 origin of replication.

Agrobacterium tumefaciens strain LBA4404Thy-: This is an auxotrophic version of LBA4404 with the thymidylate synthase gene (thyA) deleted from the chromosome (Ranch et al., 2012; Anand et al., 2018).

Agrobacterium strain carrying the constructs: LBA4404Thy- harboring both pKL2013 and pKL2299 is called KL2013vir/thy and used in this study.

Metromix 360 (Sungro, Agawam, MA, United States).

Pro-Mix BRK20 (Pro-Mix, Quakertown, PA, United States).

1801 deep inserts (T.O. Plastic, Clearwater, MN, United States).

Standard Flat with drain holes, STF-1020-OPEN (T.O. Plastic, Clearwater, MN, United States).

Standard 2.5-gallon (9.4 L) nursery pots (Nursery Supplies Inc., Chambersburg, PA, United States).

Humi-dome (Hummert International, Earth City, MO, United States).

Tomato maker 4-2-6 (Organic labs, Fort Pierce, FL, United States).

Osmocote 15-9-12 (Everris, Geldermalsen, Netherlands).

Sprint 330 Iron chelate (BASF, Ludwigshafen, Germany).

Pollination apron.

Scissors.

70% Ethanol to sterilize scissors.

Shoot bags (Lawson Pollinating Bags, Northfield, IL, United States).

Tassel bags (Lawson Pollinating Bags, Northfield, IL, United States).

Permanent marker–Chisel tip for broad strokes, black, standard (NOT ethanol resistant) recommended to avoid fading in the sun.

Stapler and staples–(e.g., Ace Clipper Model no. 702).

Non-skid paperclips.

Greenhouse with tables and lighting–28°C, 16 h day/25°C, 8 h night.

Stirring hot plate and stir bars.

Beakers.

Glass flasks and sterile pouring beaker or 1 L lidded bottles for autoclaving media.

Water bath, 55°C.

Autoclave.

Petri dishes–100 mm × 15 mm and 100 mm × 25 mm, sterile.

Large, sterile Petri plate (e.g., 120 mm × 120 mm × 15 mm square plate or 150 mm × 15 mm round plate).

0.22 μm syringe filter.

0.22 μm Stericup^® vacuum filtration system.

pH meter.

Laminar flow hood.

Sterile culture loops.

Genesys 10S UV-VIS spectrophotometer (Thermo scientific, Waltham, MA, United States).

Semi-micro cuvettes (Cat # 14955127, Fisher scientific, Pittsburgh, PA, United States).

Bleach (8.25% sodium hypochlorite, e.g., Clorox).

70% ethanol in a spray bottle for disinfecting.

Bead sterilizer.

Forceps.

Scalpel and blades.

Handles for ears–recommend cheap, stamped sheet metal forks with all but one tine broken off.

Embryo dissection tool (e.g., Fisher brand Handi-Hold Microspatula or Hu-Friedy plastic filling instrument #8A).

Parafilm.

Micropore tape.

Plastic culture box to hold multiple plates, e.g., square plastic container with lid (31 cm × 23 cm × 10 cm, 12-5/16″ × 9-1/16″ × 4″, W × L × H, Cat # 295C), or square hinged plastic container with snap closure (33.3 cm × 9.2 cm × 5 cm, 13-1/8″ × 9-3/16″ × 2″, W × L × H, Cat # 700C) (Pioneer plastic, Dixon, KY, United States).

26–28°C plant tissue culture incubator (dark).

26–28°C plant tissue culture incubator (lighted 16 h day/8 h night, 120–150 μmol/m²/s).

DNA extraction buffer (200 mM Tris-Cl, 250 mM NaCl, 25 mM EDTA, 0.5% sodium dodecyl sulfate).

RNase A (DNase and protease-free).

Chloroform.

Isopropanol.

80% ethanol.

DNase-free water.

ExoSAP-IT™ (Applied Biosystems, Waltham, MA, United States).

1.5 mL centrifuge tubes.

Scalpel and blades.

Forceps.

Maize B104 plants are grown in a greenhouse equipped with artificial lighting and reverse osmosis (RO) water systems in the Donald Danforth Plant Science Center (St. Louis, MO, United States). The greenhouse conditions are as follows: photoperiod, 14/10 h (day/night); temperatures, 28/22°C (day/night); relative humidity, 40/50% (day/night); Supplemental light intensity, 150 μmol/m²/s at 1.5 m (5 ft) above ground. Lighting is a mixture of 50% of 1000 W Metal Halide and 50% of 1000 W High-Pressure Sodium Fixtures mounted 4 m (13 ft) above ground.

Maize seeds are usually sown in small containers filled with commercial potting mix. Plastic inserts (1801 deep inserts) are filled with Metromix 360 potting mix and placed onto STF-1020-OPEN flat with drain holes (Figure 2A). Before sowing, the potting mix must be wetted thoroughly but not excessively.

B104 seeds (one per cell) are placed in the potting mix approximately 2.5 cm (1 inch) deep from the surface. The tray is covered with a plastic humi-dome (Figure 2B) and placed on a bench in the greenhouse with the conditions described above. In the case of needing year-long donor plant production, seeds can be sown every 3–4 days, depending on growth space availability.

Soil moisture is monitored on daily basis. Sprouting seeds can be expected about 4 days after the sowing. Keep the humi-dome on the flat until the sixth day after the planting. Seedlings are watered only if the potting mix appears to be dry.

Seedlings with ∼4–5 leaves can be transplanted into large pots approximately 2 weeks after the sowing (Figure 2C).

Potting mix Pro-Mix BRK20 is used to fill 2.5 gallons (9.4 L) nursery pots. Each pot is top-dressed with 2/3 tsp tomato maker, 2/3 tsp Osmocote, and 1/2 tsp Sprint Iron chelate. Then RO water is used to water the pot until the potting mix becomes saturated.

Seedling, with soil adhering to the roots, is carefully transplanted from a small plastic insert into the large pot with properly prepared and wetted potting mix (Figure 2D) and placed on a 1 m tall bench (about 2.7 m or 9 ft below the lights) for about 2 weeks (Figure 2E). Plants can be moved to the floor 2–3 weeks after the transplanting (Figure 2F) and spaced at least 12.5–15 cm (5–6 inches) apart between pots.

Plants are watered as needed. Overwatering of maize plants will result in poor development of the root system. Maize plants grow differently under different growth conditions. In the greenhouse described in this work, the B104 plants are watered daily with fertilizer solution (Jacks 15-5-15 at 400 ppm, pH 6.0). Every third watering is with RO water. Watering frequency is 1–2 times daily as needed.

Female flower (silk) emergence (Figure 2G) occurs approximately 60 days after the sowing. To ensure controlled pollination, emerging ears are covered with a wax-treated paper shoot bag prior to silk emergence (Figure 2H). To avoid pollen cross-contamination, plants and shoot bags should be closely monitored. Any uncovered or improperly covered female flowers should be discarded.

Male flower (tassel) emergence (Figure 2I) should occur around the same period, although not always synchronized with the female flowers. The tassel should be covered by a brown pollination bag (Figure 2J) once it starts to produce pollen.

To produce immature embryos, silks are pollinated using pollen from the same plant or a sibling B104 plant. To encourage silks growth, the ear shoot tip can be cut (Figure 2K) one day prior to the pollination. Using a pair of clean scissors, cut the female flower ∼2.5 cm (1 inch) from the top of the shoot. The cut ear is immediately covered by a shoot bag and allowed to grow for another day to ensure freshly grown, evenly distributed silks are used for pollinations. Before being used to cut the next ear shoot, scissors should be sanitized with 70% ethanol to prevent fungal or bacteria contamination carryover between ears.

Pollination should take place in the mid-morning the next day. Pollen is collected from a B104 plant with a bagged tassel. To release pollen from anthers, the tassel bag, and the tassel should be shaken by hand gently, the bag with the pollen is then removed from the tassel for pollination (Figure 2L). The tassel is immediately covered by another new tassel bag if the tassel is to be reused in next day. Otherwise, the tassel should be removed from the plant (detassel) to avoid any unwanted pollen shedding and cross-contamination.

Identify a bagged ear shoot that was cut the previous morning. Remove the shoot bag, quickly but carefully sprinkle the pollen evenly onto the freshly grown silks (Figure 2M). The pollinated ear is immediately covered with a brown pollination bag that is labeled with the cross information such as plant ID, date, and nature of the cross (Figure 2F). Pollen collected from one tassel with 50–70% opened anthers can be used to pollinate multiple ear shoots. Pollen from freshly opened anthers is most desirable, although B104 pollen of up to 4 days post-anther emergence can still be used.

Immature embryos of 1.8–2.0 mm (up to 2.5 mm) in size are ideal for maize transformation. To obtain the embryos of the appropriate sizes, ears should be checked for embryo development while they are still on the plant (Figure 2N) around 10 days after the pollination. Embryo sizes can be measured by a caliper (Figure 2O) or estimated by a small ruler (Figure 2P). Typically, ears can be harvested between 10 and 14 days depending on the temperature of the greenhouse, which may be impacted by exterior weather conditions. In general, embryos reach desired sizes faster in the summer season than in the wintertime.

At least three days before the infection or earlier, prepare a “mother” plate (Table 2) of Agrobacterium strain KL2013vir/thy from a glycerol stock that was originally prepared from a single colony. Streak Agrobacterium on the “mother” plate using a sterile loop.

Incubate the mother plate at 28°C in the dark for 36∼48 h. The “mother” plate with fully grown Agrobacterium colonies (Figure 3A) can be stored at 4°C in the dark for up to 7 days.

Day 0: One day before the embryo infection experiment, prepare a “working” plate (Table 2) of KL2013vir/thy. Collect 7–8 colonies (2 mm diameter) from the “mother” plate (Figure 3A) and streak them evenly on the “working” plate surface. The purpose of collecting at least 7–8 colonies from the mother plate is to ensure that sufficient Agrobacterium cells are inoculated on the “working” plate. An L-shaped spreader can be used to evenly spread the cells on the plate. To ensure enough Agrobacterium culture for infection, prepare one or two additional “working” plates as back-ups.

Incubate the “working” plates at 28°C in the dark for 16–20 h (Figure 3B). Avoid using “working” plates that are older than 24 h after the streaking.

Remove the husk leaves and silks from the ears harvested from the greenhouse. Insert an ethanol-sterilized handle into either the top or the base of the ear (Figure 3C). The ear can be cut into two halves horizontally to fit into a large container for sterilization.

Place the ear into a large, sterile beaker on a laminar flow bench. Pour sterilization solution [2 L of 20% commercial bleach (8.25% sodium hypochlorite) plus two drops of Tween 20] into the beaker so that the solution will cover the ear completely. Gently swirl the ears and tap them to the beaker bottom 10–20 times to remove bubbles trapped on the surface of the ears. Keep the ear in the sterilization solution for 30 min, stirring lightly every 10 min to ensure all ear surfaces are in contact with the solution.

Discard the sterilization solution into a large waste container. Add a large volume of autoclaved Millipore water into the beaker to wash the ears. Stir lightly 2–3 times and leave the ears in the water for 5 min. Repeat the wash step two more times, for 2 min each to complete the sterilization step.

Place the sterilized ear on a large, sterile Petri plate on the laminar flow bench. Using a sharp, clean scalpel, carefully remove the opaque kernel top, 2∼3 mm thickness at once until the clear endosperm is exposed (Figure 3G). Cutting too deep into the kernel may accidentally damage the embryos, especially when they are larger than 2 mm.

Insert a sterile micro-spatula (Figure 3D) perpendicular to the ear axis at the bottom of the kernel then squeeze gently toward the ear-tip side (Figures 3G,H). The embryo is located at the bottom, ear-tip side of the kernel. The embryo will emerge between the pericarp and endosperm. If the embryo does not pop up from the kernel, gently scoop out the endosperm with the spatula to isolate the embryo from the adhering tissue. Figure 3I demonstrates the use of a dental filling instrument for embryos isolation. The micro-spatula can be used for squeezing or scooping, while the dental filling instrument is more efficient in scooping the immature endosperm.

Put the embryo into a 2 mL Eppendorf tube containing 1.6 mL of liquid 700A infection medium without thymidine and acetosyringone (AS, Table 2). Collect up to 100 embryos in one tube (Figure 3G, Step 2). The embryos can be stored in the tube on the bench for 2–3 h.

In a 50 mL conical tube containing 10 mL of 700A infection medium (Table 2), add 20 μL of 25 mg/mL thymidine (final concentration, 50 mg/L) and 10 μL of 100 mM AS (final concentration, 100 μM). Thymidine and AS should be added freshly before each experiment. Divide 700A infection medium into two 50 mL tubes (5 mL each).

Harvest Agrobacterium cells from the working plate (Figure 3B) using a sterile loop. Inoculate the bacteria in the 5 mL 700A infection medium. Vortex the tube for 4–5 s to fully resuspend the bacteria.

Measure optical density (OD) of Agrobacterium suspension at 550 nm on a spectrophotometer and adjust the OD₅₅₀ of the suspension to 0.50 ± 0.05. This Agrobacterium suspension should be used immediately as it can readily form aggregates within 30 min after inoculation.

Before infection, wash the isolated embryos once with a freshly prepared 700A infection medium. Carefully remove the 700A infection medium using a pipet, then add 1 mL of freshly made Agrobacterium suspension to the embryos in the tube. Cap the tube and gently invert it 3–4 times. Incubate the tube at room temperature (22–24°C) for 5 min, by placing it sideways on the bench.

Tap the tube 4–5 times to dislodge the embryos from the sidewalls of the Eppendorf tube. Then pour the bacteria suspension containing the embryos onto a co-cultivation plate (710I, Table 2). Tilt the plate to collect and remove as much bacterial suspension as possible with a pipette.

Under a dissecting microscope in the laminar flow bench, carefully orient the embryos by placing them scutellum side (smooth and round side) up (Figure 3G, Step 3). Use the micro spatula or the dental filling instrument scaler (Figure 3D) to gently flip the embryos; avoid damaging the embryos (Figure 3E).

Wrap the plates with a micropore tape or place the unwrapped plates in a clean plastic culture box (Figure 3F). Incubate the plates at 20°C in the dark for 16–20 h.

Transfer and place the embryos, scutellum side up, to a resting medium (605G, Table 2). Place no more than 25–30 embryos per plate. Incubate the plates at 28°C in the dark for 7 days. Transient RFP expression is visible 3 days post-infection.

Eight days after infection, transfer the embryos to a maturation medium supplemented with 3 mg/L bialaphos (13329B3, Table 2). Carefully remove growing hypocotyls with forceps and scalpel. This is to prevent the hypocotyl from lifting or re-orienting the embryo away from the medium. Place no more than 8–10 embryos per plate. Incubate the plates at 28°C in the dark for 10 days. Growing callus tissue emerges in this stage.

Subculture the callus pieces to a new maturation medium containing 6 mg/L bialaphos (13329B6, Table 2). If the calli grow vigorously (over 2 cm²), place 4–5 calli per plate. The base of calli with growing shoots may break naturally. If it breaks apart, spread the growing shoots onto the medium. Incubate the plates at 28°C in the dark for 14 days.

The RFP gene mCherry expression in the callus or developing shoots can be examined by using either a fluorescent microscope (Olympus SZH10 stereo microscope with Texas red filter, excitation wavelength 535–585 nm, emission wavelength 605–690 nm) or using a NIGHTSEA dual fluorescent protein flashlight and filter glasses (NIGHTSEA LLC, Lexington, MA, United States). Tiffen 29 dark red filter can be used for DSLR camera for imaging.

Transfer callus pieces with fully developed shoots onto a rooting medium containing 2 mg/L bialaphos (13158B2, Table 2). Place one or two callus pieces in one plate to ensure enough space for shoot growth. Remove excessive callus materials around the shoots. Incubate the plates at 26°C in a light chamber, with a photoperiod of 16 h light and 8 h dark (80 μmol/m²/s) for 14 days.

Plantlet (>4 cm) with developed roots (>2–3 cm) (Figures 4A,B) can be transplanted to soil and moved to the growth chamber for acclimatization. Green mature shoots without roots may be subcultured for an additional 14 days under the same condition. The excessive callus materials associated with the shoots should be removed during the subculture.

If the T0 plantlets are to be brought to maturity for seeds, pollen donor plants will need to be planted at this time or 60 days prior to the projected date of pollination. This step needs to be taken to ensure access to high-quality pollen. Because female and male flowers of in vitro regenerated maize plants are often not synchronized, wild-type B104 plants can be used as pollen donors to pollinate transgenic T0 female flowers. When the root of the T0 plant is visible in the Petri dish, germinate B104 seeds every 5–7 days until all transgenic plantlets are transplanted from Petri dishes to the soil.

When T0 plantlets have established in tissue culture rooting media, they are ready to be transplanted to soil. Transplant plantlets with healthy roots (longer than 2–3 cm, as shown by red arrows in Figure 4B). Smaller plants can be subcultured to fresh medium for continued growth.

Fill 1801 deep inserts (2/3 of the height) with autoclaved potting mix such as Metromix 360 and place them into STF-1020-OPEN flat with drain holes (Figure 2A). Make a hole for the plant in the middle of each cell using a spoon. Autoclaved potting mix is recommended to prevent fungal contamination. The autoclaved moisten potting mix (60∼70% saturated) can be stored in a cool place for 2 weeks.

Using a pair of sterile forceps, first break up the media around the roots on the plate, then hold the base of the plantlet, gently pull it from the gel media without damaging the roots.

Hold the plantlet with a gloved hand and gently swirl it in sterile water 3–4 times to wash off the plant medium. It is important to remove excess medium associated with the roots as the presence of gel may bring fungal growth, which can kill the young plant. However, it is also important to avoid any physical damage to the plants during the process. If the roots are coated with plant medium, do not scrape it off, but rather gently rinse to remove the remaining media clumps.

Carefully remove any callus and shoots without roots using a pair of sterile forceps. Non-regenerated tissue will naturally come off from the regenerated plantlets. If the tissue is tightly bound with the plantlet, do not use a scalpel to dislodge them from the plantlet. For bushy plants (Figure 4C) or multiple shoot clusters, carefully separate individual plants from the cluster as much as possible but avoid tearing or damaging roots or shoots in the process. The cleaned individual plantlet (Figure 4D) can be examined for the red fluorescent protein expression on the root at this stage.

Place one plantlet per cell. Bury the roots and base of stem 3–5 cm deep in the potting mix (Figure 4E). Cover the flat with a plastic humi-dome (Figure 4F). Keep the plantlet upright to avoid touching the side of the humi-dome. Place the tray in a growth facility with 14/10 h (day/night) photoperiod, and 28/22°C (day/night) temperature. Plantlets transferred from the tissue culture media to potting mix tend to desiccate quickly due to low humidity. Therefore, the use of a humi-dome is critical to maintaining the moderate humidity level for the optimal acclimation to greenhouse growth conditions.

Check the plantlets and soil moisture daily. Water only when the soil becomes dry.

Four to six days after transplanting, remove the plastic humi-dome and continue watering as needed.

Phenotyping and genotyping of the T0 plantlets can be performed approximately 10–12 days after the transplantation (see below sub-section).

After approximately 2 weeks, healthy and robust-looking plantlets can be transplanted into a large pot as described in steps 6–8 in the above section of “Growing donor plants for the production of the immature embryo for transformation.” Both 1 gallon (3.8 L) and 2.5 gallon (9.4 L) pots can be used for growing maize plants to full maturity. It is critical not to overwater the plants, especially when they are young.

When ears start to emerge (50–100 days post-transplant), T0 plants are pollinated to produce seed for subsequent generation testing and analysis. Male and female flower development can occur at separate times on an individual T0 plant, which will necessitate crossing B104 wild-type or other transgenic B104 plants to generate seeds.

To avoid pollen cross-contamination, transgenic plants should not be grown with any non-transgenic plants in the same growth room. All emerging ears are covered with a wax-treated paper shoot bag to prevent pollen contamination of the silks before pollination.

Cover the tassel with a bag. The bag should be folded to secure it around the stem with a paper clip to prevent pollen from escaping (Figure 2J).

If the greenhouse space is crowded, detasseling the T0 plants will be necessary to prevent transgenic pollen cross-contamination. When the tassel is extended from the whorl, pull it out gently for detasseling. The detasseling may stimulate shoot growth.

Follow steps 9–13 described in the section “Growing donor plants for the production of the immature embryo for transformation” for T0 plant pollination.

Immediately cover the pollinated ear with the pollination bag. Record the ear donor, pollen donor, silking date, and pollination date on the tassel bag. Staple the back of the tassel bag.

Ten days post-pollination, remove the pollination bag and slide it into the node between the ear and the plant’s stalk. Peel back the husk to observe the seed set and allow fresh air to the ear to prevent fungal contamination.

Thirty days post-pollination, stop watering to allow dry down and senescence of plants with mature ears. Keep the tassel bag on the stalk to keep the record.

Forty days post-pollination, harvest mature dried T0 ear and place in pollination bag, record the harvest date on the bag.

Dry the ears at room temperature inside the pollination bag on a benchtop for 2–3 weeks.

Remove kernels from ear and place in a seed packet, label the packet with appropriate experimental information, harvest date, and seed weight.

Seeds can be stored in a cool and dry place (15°C, Relative Humidity < 30%). For long-term storage, seeds can be kept at −20°C with an additional desiccant such as silica gel.

Knockout mutants of maize glossy2 (Gl2) gene (Bianchi et al., 1975) can be evaluated on T0 plantlets. Although the gene product of Gl2 is still unknown, it is responsible for the formation of a hydrophobic waxy cuticle layer in juvenile leaf tissues. The loss-of-function mutants can be readily identified by misting water on the young leaf surface (Bianchi et al., 1975). Water will roll off from wild-type leaf, whereas gl2 null mutations affect the deposition of extracellular cuticular lipids allowing water droplets to adhere to the leaf surface. This is most obvious when working with homozygous or biallelic mutant plants.

Note that misting on T0 plants can only be used as a preliminary evaluation. It should not be used to make conclusion on whether the plant is a loss-of-function mutant event. Sequencing analysis is necessary to verify and confirm the phenotypes observed on the T0 plants.

Plants that have stayed out of the humi-dome at least for 4–6 days (typically 10 days post-acclimatization) can be evaluated for gl2 phenotype. Use a spray bottle to mist the leaf surface with tap water.

The gl2 loss-of-function mutants would show water droplet adherence on both sides of the leaf blades while the surface of wild-type leaf blades would repel water droplets due to the hydrophobic cuticle layer.

Total genomic DNA can be isolated from the transgenic maize plants using a modified version of the protocol described by Edwards et al. (1991). Sample 3∼4 cm² of fresh leaf tissue in a 1.5 mL tube. Add 500 μL DNA extraction buffer containing 100 μg/mL RNase A into each tube and grind the leaf tissue using a polypropylene homogenizing pestle attached to an electric drill. Incubate the tubes for 5–10 min at room temperature. If the leaf tissue is smaller than 2 cm², reduce the DNA extraction buffer to 250 μL.

Add 500 μL (equal volume to the DNA extraction buffer used in step 1) of chloroform and mix gently by inverting the tubes for 3 min, and centrifuge for 5 min at a maximum speed using a benchtop centrifuge (21,130 × g).

Carefully transfer about 300 μL of the top aqueous phase to a new tube and add 240 μL (80% of aqueous phase, v/v) of isopropanol. If 250 μL of extraction buffer was used, then transfer 150 μL of the aqueous phase into a new tube and add 120 μL of isopropanol. Be careful not to take any interphase or organic phase into the new tube. Mix well by inverting 4–5 times and centrifuge for 5 min at a maximum speed (21,130 × g).

Remove the supernatant using a 1 mL pipette and rinse the genomic DNA pellet once with 500 μL of ice-cold 80% ethanol.

Centrifuge for 2 min at a maximum speed on a benchtop centrifuge and carefully remove the ethanol using a 1 mL pipette. Briefly spin down the tubes to collect the remaining ethanol on the tubes and remove it using a 200 μL pipette. Air-dry the genomic DNA pellets for 10 min on a flow bench.

To redissolve the genomic DNA pellet, add 50 μL of nuclease-free water and finger tap the tubes 4–5 times. Measure the genomic DNA concentration using a Nanodrop spectrophotometer. Typically, 20–200 ng/μL of genomic DNA is obtained using this method (1–10 μg of total genomic DNA), and 1 μL of genomic DNA is used for a 20 μL PCR reaction.

Perform PCR screening to identify transgenic plants and to genotype the target gene, Gl2. PCR amplifies ∼1 kb fragments from the gl2 locus and the Cas9 gene using the primer sets and PCR conditions previously reported (Lee et al., 2019).

Run 1% agarose gel electrophoresis to resolve the PCR products and identify Cas9-positive T0 plants.

For the Cas9-positive plants, treat the PCR products from the gl2 locus with the ExoSAP-IT™ reagent for Sanger sequencing analysis. Mix 5 μL of gl2 PCR product with 2 μL ExoSAP-IT™ reagent and incubate 30 min at 37°C followed by incubation for 15 min at 80°C for enzyme inactivation.

Perform Sanger sequencing using a preferred service provider.

Analyze the sequence trace files from the transgenic plants and wild-type B104 using publicly available web-based CRISPR editing analysis software; Tracking of Indels by Decomposition (TIDE; Brinkman et al., 2014), Inference of CRISPR Edits (ICE; Hsiau et al., 2019), and Degenerate Sequence Decode (DSDecodeM, Xie et al., 2017). Short indel mutations can be readily identified by these programs and it is recommended to use at least two software to cross-check the outcomes.

Note: Direct Sanger sequencing of the PCR products and trace file analyses using online decode tools demonstrated here are recommended for rapid screening of mutant events. For precise/final genotyping analysis, cloning of the PCR products into a plasmid vector and then Sanger sequencing of multiple clones are recommended.