Accelerated In Vitro Propagation of Sweetpotato Clones (Ipomoea batatas L.)

Abstract

Fast and reliable propagation of plant material is an important need in different stages of breeding programs and production systems. In vitro propagation ensures that pathogen- and virus-free plants stay phytosanitary clean over time while providing high multiplication rates. Using liquid instead of solid culture medium can reduce the interval of individual propagation cycles and contributes to speeding up of the process (1.5–2.5 times), especially during the initial growth phase of the plants. Sophisticated immersion systems have been developed for many plant species, but they are difficult to apply when hundreds or thousands of different genotypes are propagated simultaneously. Additionally, these systems require a high input of technical equipment, know-how and experience to avoid bacterial or fungal contamination during the propagation process. The following protocol describes a low-input suspension technique that combines the use of liquid and solid medium, and permits the successful propagation of genetically diverse sweetpotato genotypes [Ipomoea batatas (L.) Lam.] with a high multiplication rate. As sweetpotato is an important staple crop in low-income/technology countries of Africa, Asia, and South America, the described method may find valuable application for the breeding programs in these regions.

7.1 Introduction

The efficient large-scale propagation of genetic resources of important staple crops such as sweetpotato is a crucial component for industrial production, breeding programs, and research projects. Plant material can be propagated clonally in the greenhouse (cuttings), field, or laboratory, but due to practical and economic reasons, propagation in the greenhouse is frequently preferred over in vitro propagation in the laboratory. However, in vitro propagation has the distinct advantage that it is performed under controlled environmental and sterile conditions within glass test tubes or other vessels, which allows a space-efficient and continuous propagation that guarantees that pathogen-free material stays clean over time. The use of virus-free material not only contributes to higher yields and thus increased quality of storage roots produced in the field, but also satisfies phytosanitary criteria required for international exchange of plant genetic resources (PGR). Further, the use of phytopathogen-free in vitro material eliminates the effect of virus infection as a source of variation in field, greenhouse, and laboratory experiments as virus-infected plants may show a different pattern of growth, development, and crossing ability. Although plant tissue culture methods were developed in the first half of the twentieth century, based on pioneering work of Haberlandt, Kotte, Robbins, Went, White, Skoog, Miller, and others (Haberlandt 1902; Kotte 1922; Robbins 1922; Went 1926; White 1939; Skoog and Miller 1957), a major general breakthrough was achieved in the early 1960s, through the development and publication of the probably most famous plant culture medium, the Murashige & Skoog medium (MS) (Murashige and Skoog 1962). Over the following decades, it was shown that thousands of species grow well on solid MS medium, generally with full salt strength concentration, but sometimes also with a reduced (½ or ¾) salt strength. Haberlandt already used suspension culture in liquid medium for his pioneer experiments (Haberlandt 1902), but only after White published the use of liquid culture media for the culture of tomato roots tips in 1933/1934 (White 1933, 1934), did liquid medium become established as an alternative to solid culture medium. Later, in the 1980s and 1990s, more sophisticated systems were developed, both for complete and temporary immersion of plant explants (Harris and Mason 1983; Tisserat and Vandercook 1985; Aitken-Christie and Davies 1988; Simonton et al. 1991; Alvard et al. 1993), principally with the objective to increase oxygenation in the culture medium, remove phytotoxic compounds, maintain the nutrient balance (renewal of medium), and avoid hyperhydration. All these are excellent propagation methods, but they involve a high level of technical knowledge and experience, as well as specialized equipment, i.e. inputs develo** economies and economies in transition do not always have easy access to obtain. Additionally, the risk of contamination increases when liquids are continuously pumped, moved from one vessel to another, or completely renewed. Based on our experience with hundreds of genetically diverse sweetpotato accessions, the response was uniform and favorable in classic suspension liquid medium under rotation (Fig. 7.1), with only a very limited number of genotypes showing signs of hyperhydration.

Fig. 7.1

(a) Five to 8-week-old in vitro sweetpotato plants are used as starting material. Plants should be completely developed, have minimum 5–6 nodes (leaves), and show a normal aspect (no signs of yellowing, deformation, hyperhydration, etc.) (b) (I) In vitro plants placed on sterile paper sheet. Roots, leaves, and tip are cut off using forceps for plant tissue culture and scalpel No. 10. (II) Cutting of uni-nodal stem segments of ~0.5–1.5 cm. Do not cut too close to the axillary bud to avoid damaging it. (III) Stem segments ready to be transferred to liquid culture medium. (c) Uni-nodal stem segments placed in liquid culture medium (50 mL Erlenmeyer flask, ~22 mL of culture medium). Place 5–15 stem segments per flask, close flask with sterile aluminum paper and seal with saran wrap or parafilm^®. (d) Incubate Erlenmeyer flasks under rotation (80–90 rpm) at 25–29 °C with a 16 h/8 h photoperiod. Higher temperatures (28–29 °C) further speed up plant growth. (e) In vitro plants coming out from liquid medium, 12–18 days after propagation. Plants should be fully rooted, and minimum have 2–4 leaves for transfer to solid medium. (f) Young plants that were transferred from liquid to solid culture medium. Assess growth and contamination on a weekly basis. After 3–6 weeks on solid culture medium, plants are ready for a new cycle

The simple suspension method described here permits laboratories to propagate, with a limited input of resources up to 80–100 accessions in a physical space of ~0.7 m². The cross-contamination risk is minimal (as explants are always contained within the same sealed container), and the use of uni-nodal explants results in a high multiplication rate. The described protocol was originally developed to speed up the propagation process of the initial phase of cryopreservation at CIP (years 2017–2018), and subsequently underwent additional improvements in the framework of a phytosanitary project (years 2019–2020).

7.2 General Considerations, Equipment, Materials, and Preparation of Culture Media

All culture media should be prepared with distilled deionized water and commercial, pre-prepared, full-strength Murashige & Skoog medium (MS). Analytical grade reagents and calibrated/validated equipment should be used during all protocol steps (e.g. pH-meter, autoclave, laminar flow chamber, electronic dispensing pump, etc.). Chemicals are stored at room temperature (22 ± 3 °C), under refrigeration (5 ± 3 °C) or in the freezer (−15 ± 5 °C), according to specifications in the Material Safety Data Sheet of each product. Reagents should be dissolved using a magnetic stirrer at 600–1000 rpm, depending on the volume of culture media that are being prepared. Label stock solutions with the following minimum information: type and concentration of solution, name of person who prepared the solution, known hazards (included in a NFPA diamond), and date of preparation.

7.2.1 Equipment and Materials

• 1. Equipment: laminar flow chamber; analytical balance; autoclave; pH-meter; magnetic stirrer; microwave oven; water distiller; orbital shaker; dispensing pump; heat sealing machine; vacuum pump (for filter sterilization system); refrigerator/freezer; incubation chamber/room.

• 2. Materials: aluminum foil; autoclave indicator tape; scalpel holder; forceps for tissue culture (23 cm, fine point); scalpel blades (#10); Sterifil^® Aseptic System (Merck Millipore); prefilter (Type: AP15, Ø 47 mm); filter (Type: GSWP, pore size: 0.22 μm, Ø 47 mm); glass beaker (100 mL); glass measuring cylinder (100 mL); glass test tubes (25 × 150 mm) with plastic caps; weight spatulas; pipette pump or bulb (for 10 mL pipette); plastic beakers (1 and 2 L); plastic racks for 25 × 150 mm test tubes (7-way tray); plastic weighing dishes (small and large size); Pyrex glass bottles (autoclavable, 1 and 2 L); saran wrap; parafilm; sterile Erlenmeyer glass flasks (50 mL); screw cap plastic flasks (20 mL); sterile paper sheets (A5); sterile plastic pipettes (10 mL); sterile tool holder; sterilization sleeves (width: 30 cm); stir bars (1/8″ and 3/8″); wash bottle with alcohol (70%).

7.2.2 Media Preparation

7.2.2.1 Preparation of Solid Culture Medium (1 L)

• 1. Components: 4.33 g/L of MS salts; 5 mL/L vitamin stock solution (see Note 1); 1 mL/L thiamine stock solution (at 100 ppm) (see Note 2); 1 mL/L gibberellic acid stock solution (at 1000 ppm) (see Note 3); 30 g/L sucrose; 3.0 g/L phytagel.

• 2. Pour ~600 mL distilled-deionized water into a plastic beaker (1 or 2 L). Place beaker on a magnetic stirrer and let stir at medium speed.

• 3. Dissolve 4.33 g MS salts without vitamins.

• 4. Dissolve 30 g sucrose.

• 5. Add 5 mL previously thawed vitamin stock solution and 1 mL gibberellic acid stock solution.

• 6. Add 1 mL previously thawed thiamine stock solution.

• 7. Pour culture medium into a plastic measuring cylinder (1 L) and bring it up to a final volume of 1000 mL with distilled-deionized water. Pour culture medium back into the beaker.

• 8. Measure pH and adjust with HCl (1 M) or NaOH (1 M) as needed to a value of 5.70 ± 0.02 (see Note 4).

• 9. Place 25 × 150 mm test tubes in autoclavable plastic racks (36 tubes per rack; 3 racks per liter of culture medium [= 108 tubes]).

• 10. Add 3.0 g of phytagel to the culture medium.

• 11. Dissolve phytagel in microwave at 100% intensity for 10 min, pause after 7 min and mix the medium on a magnetic stirrer (see Note 5).

• 12. Using an electronic dispensing pump, dispense 9 mL of culture medium per 25 × 150 mm test tube. Cap all test tubes. Label one of the tests tubes with type of media, date, and initials of the person that prepared it. Place an autoclave indicator tape onto one of the tube caps of each rack.

• 13. Autoclave the culture medium for 20 min (at 121 °C, 15 psi). Let the culture medium cool down to room temperature (23 ± 3 °C).

• 14. Store culture medium at 5 ± 3 °C for a maximum period of 2 weeks.

7.2.2.2 Preparation of Liquid Culture Medium (1 L)

• 1. Components: 4.33 g/L MS salts; 5 mL/L vitamin stock solution (see Note 1); 1 mL/L thiamine stock solution (at 100 ppm) (see Note 2); 1 mL/L sterile gibberellic acid stock solution (at 1000 ppm) (see Note 3); 30 g/L sucrose.

• 2. Pour ~600 mL of distilled-deionized water into a plastic beaker (1 or 2 L). Place beaker on a magnetic stirrer and let stir at medium speed.

• 3. Dissolve 4.33 g MS salts without vitamins.

• 4. Dissolve 30 g sucrose.

• 5. Add 5 mL previously thawed vitamin stock solution.

• 6. Add 1 mL previously thawed thiamine stock solution.

• 7. Pour culture medium into a plastic measuring cylinder (1 L) and bring the solution up to a final volume of 1 L with distilled-deionized water. Pour culture medium back into the beaker.

• 8. Measure pH and adjust with HCl (1 M) or NaOH (1 M) as needed to a value of 5.70 ± 0.02 (see Note 4).

• 9. Pour liquid medium into a Pyrex glass bottle (1 L), close the bottle loosely to allow pressure to escape during the autoclaving process. Label bottle (type of medium, date and preparer’s initials). Place autoclave indicator tape on the bottle cap.

• 10. Autoclave the culture medium for 20 min (at 121 °C, 15 psi). Let the culture medium cool down to a temperature of ~35–40 °C (in a laminar flow chamber, LFC).

• 11. Using a sterile disposable pipette (10 mL) (see Note 6), add 1 mL of filter-sterilized gibberellic acid (GA3) stock solution. Shake gently.

• 12. Place sterile 50 mL Erlenmeyer flasks in LFC (see Note 7). Carefully remove the aluminum foil cap from the Erlenmeyer flask top and place it onto a sterile paper sheet (see Note 8). Using a separate sterile disposable pipette (10 mL), dispense 22 mL of culture medium per Erlenmeyer flask.

• 13. Cover flasks with sterile aluminum foil cap. Take care that the internal surface of the cap does not come in contact with any nonsterile object. Seal with Parafilm between aluminum foil cap and flask.

• 14. Randomly select 2–3 flasks from each batch and incubate at 25 ± 2 °C. Assess the flasks after 4–5 days for fungal/bacterial contamination. Place the rest of the sealed flasks in plastic containers (~20 flasks/container) and store at 5 ± 3 °C for a maximum period of 2 weeks.

7.3 Methods

7.3.1 In Vitro Propagation in Liquid and Solid Culture Medium

• 1. Turn on LFC and a glass bead sterilizer. Clean workspace of LFC (see Note 9). Set up tool holder and tissue culture forceps (23 cm) (see Note 10). Attach scalpel blades to scalpel holders (see Note 11). Apply best practices when working in the LFC (see Note 12).

• 2. Place sterile Erlenmeyer flasks in LFC (see Note 7). Set up sterile paper sheets (A5) in LFC (Fig. 7.2) (see Note 8). Place test tubes containing the in vitro mother plants in a test tube rack on a lab cart next to the LFC.

• 3. Read barcode labels of test tubes and print out new labels for Erlenmeyer flasks (see Note 13).

• 4. Using forceps, place a small pile of 3–4 sterile A5 paper sheets in the central part of the LFC workspace. The pile serves as a cutting surface during propagation (see Note 14).

• 5. Remove saran wrap from test tube cap and open cap. Using plant tissue culture forceps (23 cm) remove plants from the tube and place them in the central section of the paper sheet pile (see Note 15).

• 6. Using sterilized scalpel with a No. 10 blade and tissue culture forceps, remove leaves and roots of the plantlets and cut stem in several segments (or explants) of approximately 10–15 mm, with one single bud per segment (Fig. 7.3). If the distance between nodes is very short (<3 mm), cut stem segments with two buds. The internodal distance varies depending on genotype and plant age.

• 7. Uncap an Erlenmeyer flask (see Note 16). Transfer up to 15 stem segments per Erlenmeyer flask (size: 50 mL) (see Note 17). Close flask with aluminum foil cap (Fig. 7.4). Seal and label (see Notes 18 and 19).

• 8. Incubate Erlenmeyer flasks on an orbital shaker at 80–90 rpm at 25–29 °C (equipped with clamps or a rack) [see Note 20], light intensity of 100 ± 20 μmol/m²/s and photoperiod of 16 h light/8 h darkness (Fig. 7.5) [see Note 21].

• 9. After 10–18 days in liquid medium (genotype-dependent), transfer rooted plantlets from Erlenmeyer flasks to glass test tubes (25 × 150 mm) with sterile solid culture medium (see Note 22). Use long tissue culture forceps for plant transfer. Place 2 plants per test tube (Fig 7.6) [see Note 23]. Explants that did not develop into complete and rooted in vitro plants are transferred to Erlenmeyer flasks with fresh liquid medium and incubated for 10–18 additional days. Occasionally, although rare, accessions may not grow well in liquid medium, in which case it is recommended to propagate them in deep petri dishes on solid culture medium, previous transfer to test tubes (Fig. 7.7) [see Note 24].

• 10. Close test tubes with sterile cap. Seal cap with saran wrap (2 layers) and label with barcode label (see Note 19).

• 11. Incubate test tubes for 4–5 weeks (genotype-dependent) at 24 ± 1 °C, light intensity of 100 ± 20 μmol/m²/s and photoperiod of 16 h light/8 h darkness. Adult in vitro plants can be submitted to a new propagation cycle (Fig. 7.8) (see Note 25).

Fig. 7.2

Set up of supplies in Laminar Flow Chamber (LFC), required for in vitro propagation. (a) Glass bead sterilizer (at 250 °C). (b) Forceps for tissue culture (23 cm) and scalpel holders with blade No. 10, supported on sterile tool holder (stainless steel). It’s recommended to work with various sets of forceps and scalpels, and to ensure that tools have cooled before use. (c) Pile of sterile paper sheets. For each propagation step locate a pile of 2–3 sheets in the central section of the LFC’s working table and replace them by a new sheet after each accession. (d) In vitro sweetpotato plants used as starting material for propagation in liquid culture medium. (e) Sterile liquid culture medium (22 mL) contained in Erlenmeyer flasks (50 mL). (f) Roll of saran wrap used for sealing the joint between cap and vessel (Erlenmeyer flask or test tube). (g) Antibacterial wipe moistened with alcohol (70%). After each propagation cycle clean table surface with wipe. (h) Alcohol spray (disinfection of hands and working surface). (i) Barcode labels which are sticked on the vessels to identify the propagated accession

Fig. 7.3

Stem segments of in vitro sweetpotato plants prior to transfer to liquid culture medium. Leaves and roots were removed carefully, and the stem was cut in uni-nodal segments of ~0.5–1.5 cm (one bud per segment)

Fig. 7.4

(a) Transfer of uni-nodal stem segments to Erlenmeyer flask. Transfer 5–15 segments per flask (50 mL). (b) Erlenmeyer flask containing stem segments. The flask is closed with a sterile aluminum foil cap that was previously sterilized together with the empty vessel (the liquid culture medium was dispensed afterwards in the laminar flow chamber). The vessel is labeled with a bi-dimensional barcode and sealed with saran wrap

Fig. 7.5

(a) Vertical view of orbital shaker equipped with extended aluminum platform and special stainless-steel rack for Erlenmeyer support. The use of a rack (instead of clamps) permits to place more flasks per area. The shown model has a capacity of 81 Erlenmeyer flasks (50 mL). (b) Horizontal view of modified orbital shaker

Fig. 7.6

(a) Aspect of in vitro sweetpotato plants after ~14 days in liquid culture medium, before its transfer to solid culture medium. In vitro plants were grown from uni-nodal stem segments. (b) Removal of in vitro plants from Erlenmeyer flask (liquid culture medium) before its transfer to solid medium. During transfer, ensure that roots, plants, and tools do not get in contact with unsterile surfaces (e.g. external surfaces of Erlenmeyer flask or test tubes)

Fig. 7.7

(a) An alternative propagation method for uni-nodal stem segments of sweeetpoato in vitro plants in deep petri dishes (depth: 25 mm, Ø 90–100 mm) on solid culture medium. Stick label on the upper part of the lid to ensure plants grow under uniform light conditions. (b) Twelve to 20-day-old in vitro sweetpotato plants grown in deep petri dishes on solid culture medium. Plants are ready for its transfer to test tubes, when they have rooted, and the plant’s tip has reached the dish’s lid

Fig. 7.8

(a) Sweetpotato in vitro plants, ~1 week after its transfer from liquid to solid culture medium. (b) Aspect of sweetpotato in vitro plants after 12–18 days in liquid culture medium, followed by 3–4 weeks on solid medium. After this period, plants can be submitted to a new propagation cycle in liquid medium

7.4 Notes

• 1. Components: 0.2 g of calcium pantothenate; 10 g of calcium nitrate; 10 g of l-Arginine; 20 g of ascorbic acid; 2 g of putrescine-HCl. Preparation: Pour ~300 mL of distilled-deionized water into a glass beaker (500 mL). Place the beaker on a magnetic stirrer and let the solution stir at medium speed. Dissolve calcium pantothenate, calcium nitrate, l-Arginine, ascorbic acid and putrescine-HCl. Pour stock solution into a glass measuring cylinder (500 mL) and bring it to a final volume of 500 mL with distilled-deionized water. Pour the solution back into the beaker and thoroughly mix with a magnetic stirrer. Using a disposable pipette (10 mL), dispense 20 mL of vitamin stock solution per 20 mL plastic vial. Cap vials and label. Store vitamin stock solution at −20 ± 5 °C for a maximum period of 2 months.

• 2. Component: 10 mg of Thiamine-HCl. Preparation: Pour ~50 mL of distilled-deionized water into a glass beaker (100 mL). Place beaker on a magnetic stirrer and let stir at medium speed. Dissolve Thiamine-HCl. Pour stock solution into a glass measuring cylinder (100 mL) and bring it to a final volume of 100 mL with distilled-deionized water. Pour solution back into the beaker and homogenize it on a magnetic stirrer. Using a disposable pipette (10 mL), dispense 20 mL of thiamine stock solution per 20 mL plastic vial. Cap vials and label. Store Thiamine-HCl stock solution at −20 ± 5 °C for a maximum period of 2 months.

• 3. Component: 0.5 g of gibberellic acid (GA3). Preparation: Dissolve gibberellic acid in 2–3 drops of NaOH (1 M) on a small plastic weighing dish. Use a laboratory spoon for dissolving and mixing the solution. Empty dissolved GA3 into a 500 mL glass beaker. Rinse off weighing dish with distilled-deionized water over the glass beaker (all dissolved GA3 should end up in the beaker). Add ~200 mL of distilled-deionized water. Place beaker on magnetic stirrer and let the solution stir at medium speed. Pour solution into a glass measuring cylinder (500 mL) and bring it to a final volume of 500 mL with distilled-deionized water. Pour stock solution back into the beaker. Measure pH and adjust to 5.70 ± 0.02 (see Note 4). Filter sterilize GA3 through the Sterifil^® Aseptic System in the LFC. Using a sterile disposable pipette (10 mL), dispense 10–30 mL of GA3 stock solution per sterile Eppendorf flask (30 mL) in LFC. Close container with sterile cap and seal with parafilm. Store GA3 stock solution at 5 ± 3 °C for a maximum period of 2 weeks.

• 4. To increase and decrease the pH of any solution, add drop(s) of NaOH and HCl to a solution, respectively, slowly while stirring the solution and monitoring the change in pH. Solutions which have a final volume of less than 500 mL, it is recommended to use more diluted solutions of NaOH and HCl for pH adjustment (e.g. 0.5 M). Start the pH adjustment by adding three to five drops of HCl or NaOH at once, while stirring the solution. Wait 2–3 min for the pH to stabilize and then continue adding smaller volumes as needed. When the pH adjustment gets close to the required value (e.g. 5.70), dilute some drops of HCl (1 M) or NaOH (1 M) in distilled water and use this solution for the final pH adjustment. Patience is key aspect in changing the pH of a solution.

• 5. Use a microwave oven to dissolve the gelling agents for solid culture media that do not contain any thermolabile compounds. When the gelling agent has started to dissolve (before boiling), take the beaker out of the microwave and thoroughly mix the solution by stirring with a magnetic stirrer to avoid formation of agglomerates. Next, place it back in the microwave and heat until boiling and mix. The gelling agent is dissolved completely when the solution changes from cloudy (milky) to a transparent/clear solution. For instance, to prepare 1 L of culture media, heat the culture medium for 7–8 min in the microwave, homogenize the solution on a magnetic stirrer, and then continue heating for 3–4 min more. The heating time will be approximately doubled for a 2 L solution. When a dispensing pump is used for pouring the culture media, the beaker containing the media is placed on a magnetic stirrer (to ensure a homogeneous composition in all vessels).

• 6. Before opening the sterile sleeve of the disposable pipette, check that neither the wrapper nor pipette have damages or have been opened. Open the wrapper and remove the pipette aseptically and insert the top into a pipette-aid (pump or bulb). Fill the pipette a bit above the desired volume line and then slowly lower the meniscus to that line. Remove the pipette from the vessel, allowing the outside of the pipette to gently touch the inner lip of the vessel to remove any adherent liquid. Aseptically move the pipette to the receiving vessel and empty the pipette content. If you are emptying the entire content of the pipette you will have to “blow-out” the remaining liquid in the tip with a firm puff of air from the pipette-aid. If the tip or tip-close part of the pipette gets in touch with any nonsterile solution or object, change the pipette by a new sterile.

• 7. Set up of Erlenmeyer flasks: Fill a sink ¾ full of water. Add neutral detergent. Preferably, use a biodegradable, phosphate-free and nonionic detergent (e.g. Scienceware^® Aquet^®). Submerge 50 mL Erlenmeyer flasks in detergent solution and let them soak for about 15 min. Brush internal surfaces of the flask using a test tube brush (Ø 20 mm). Clean the external surfaces with a scour pad. Rinse flasks 3 times with deionized water to remove remaining detergent. Let flasks dry upside down on a plastic tray overnight (min. 18 h). Rinse flask internally with alcohol (70%) and remove remaining label glue from the external surface (using alcohol [70%] and scour pad). Let flask dry for ~15 min until the alcohol has completely evaporated. Only use completely dry flasks. Using scissors or paper cutter, cut aluminum foil squares of ~6 × 6 cm. Place two centrally aligned squares onto the neck of each flask (one single layer could break). Pull down the four sides of the aluminum foil, forming a uniform flask cap. Firmly press aluminum foil along the opening of the flask and the neck. Place a maximum of 12 Erlenmeyer flasks per Kraft paper bag (#8). Seal bag with masking tape. Stick a piece of autoclave indicator tape on the bag. Dry heat-sterilize flasks for 20 min at 121 °C.

• 8. Cut A4 Bond paper in half using scissors or paper cutter. Pile up the paper sheets. Place a pile of ~1 cm of A5 paper sheets per sterilization sleeve. Seal sleeve with a heat-sealing machine. The sleeves are equipped with an internal autoclave control strip that changes color after successful autoclaving (no additional autoclave tape is required). Alternatively, a pile of A5 paper sheets (1–1.5 cm) can be placed in a kraft paper bag (#8). Seal bag with masking tape and stick a piece of autoclave indicator tape on the bag. Place 2 bags per autoclavable plastic bag and seal the plastic bag with masking tape. Autoclave sleeves or bags at 121 °C for 15 min. Dry sleeves/bags in a glassware dryer to ensure the paper sheets are completely dry. Before opening the sleeve in the LFC, check that the autoclave control strip has changed its color. Also verify that the sleeve has no signs of damage (cracks or holes). The sleeve is opened in the LFC by carefully pulling the two sides of the sleeve in opposite direction (at the sealed extreme of the sleeve). Kraft paper bags are opened in the LFC with sterile scalpel or scissors (at the extreme that was sealed with masking tape). Remove the pile of paper sheets using sterile forceps and place it in the upper left or right corner of your workspace in the LFC. The work surface of the LFC was previously disinfected with alcohol (70%). Do not to get in contact with nonsterile surfaces during the setup of the sterile paper sheets (i.e. opening of sleeve/bag, removal and colocation of paper sheets with sterile forceps). Cutting of stem segments for in vitro propagation is performed on a pile of 3–4 sterile paper sheets. Also, sterile items like aluminum foil caps of Erlenmeyer vessels are supported on a pile of 2–3 sterile papers to avoid bacterial or fungal contaminations.

• 9. After turning on the LFC check for stable air flow. Modern LFCs are equipped with a manometer to measure the velocity of the sterile air flow. A minimum air flow velocity of 0.35 m/s is required to ensure adequate sterile conditions within the chamber. Before sterilizing tools, check the glass bead sterilizer has reached its operative temperature of 250 °C. In the case no sterilizer is available, tools can also be flame sterilized using an alcohol burner. Disinfect working surface with alcohol (70%) using antibacterial wipes.

• 10. Tool holders are generally made of stainless steel or glass (= petri dish). Heat-sterilize tool holders in sterilization sleeves for 20 min at 121 °C. Alternatively, sterilize tool holders in sealed autoclavable bags. Before removing tool holder from the sleeve/bag, disinfect hands or gloves with alcohol (70%). Take care not to touch those surfaces of the holder that will support the tools. Place tool holder in the upper right/left corner of the workspace, next to the glass bead sterilizer. Distance between holder and back wall should be ~10 cm. Clean forceps and scalpel holders using an antibacterial wipe moistened with alcohol (70%). It is recommended to work with a set of 4–5 forceps and 4–5 scalpel holders. Heat-sterilize forceps for 20–30 s. Insert forceps deeply into the sterilizer’s glass beat vessel, as only those surfaces are adequately sterilized that get into contact with the glass beads. Place forceps on tool holder.

• 11. Open wrap of scalpel blade. Using forceps carefully remove blade from wrap. “Grip blade with forceps, or similar, avoiding contact with cutting edge. Hold handle in left hand with tool’s bayonet fitting uppermost. Place blade partway over handle fitting and engage slots. Slide blade until it clicks into position. To improve assembly, flex blade slightly upwards when sliding onto the handle. For removal or change of blades, grip blade with forceps making sure that the cutting edge is facing away from hand and body. Ensure the blade is pointing downwards and towards the trolley and NOT towards you or another member of your team. Whilst holding the handle firmly lift the back edge of the blade with the forceps or needle holders and slide away the handle” (Swann-Morton 2021).

• 12. Maintenance of sterile conditions within the LFC requires some basic rules and guidelines for best practice: (a) Minimize clutter, (b) Wash your hands and arms properly and thoroughly before working in the LFC, (c) Arrange objects in such a way that the work area is directly bathed with clean air from the HEPA filter, (d) Avoid spraying or squirting solutions onto the HEPA filter, (e) Remove outer pouches and wraps on the edge of the work area, (f) Keep large objects away from the back of the hood to not disrupt the laminar airflow, (g) Avoid coughing, quick movements, and talking, (h) Remove jewelry around your hands and wrists when working in the LFC, (i) Place only sterile materials inside the clean work area, and (j) Propagate plants in the central part of the work space, and not close to the outer edge of the LFC.

• 13. Use of barcode labels for reading and printing reduces the risk of human mistakes and mix-up of accessions. Barcode labels should contain basic information identifying the accession and the process stage, such as accession identifier, phytosanitary status, propagation date, lot of culture medium and name (or initials) of the person who has propagated the plants. Bidimensional or Quick Response (QR) barcodes can contain larger quantity of alphanumeric data than unidimensional barcodes.

• 14. The pile should be thick enough to avoid scalpels coming in contact with the LFC’s workspace surface during cutting. If any nonsterile object gets in contact with the upper surface of the paper pile, replace it by a new pile of three to four sterile sheets (A5). Replace the paper pile by a new one after each processed accession.

• 15. Cut sealing saran wrap vertically, using a discarded scalpel blade. Cut smoothly and away from the fingers holding the tube. Opening of test tube and removal of plants is best performed inclining the test tube to a nearly horizontal position, which avoids potential contaminants (bacteria, fungi, remnants of sealing tape) falling into the test tube. Additionally, it is easier to remove plants from the tube in a close to horizontal position. Firmly grip the plants at the basal zone of the stem during removal and carefully remove them from the tube, together or without the block of solid culture medium. Take care that long roots or culture medium do not get into contact with external surfaces of the test tube.

• 16. Loosen the aluminum paper around the neck of the flask without touching the internal part of the aluminum foil nor the external surface of the flask which was covered by the foil. Grip the base of the foil cap, carefully remove the cap from the flask and place it upside up on a pile of 2–3 sterile paper sheets.

• 17. Stem segments coming from different test tubes should be transferred into separate flasks and well labeled. When required during handling, hold the Erlenmeyer flask in its base, but never touch the sterile neck zone of the flask. During transfer, grip stem segments smoothly with forceps without damaging the bud. Segments can be transferred in groups, grip** 2–5 segments at once.

• 18. Grip aluminum foil cap in its base during closing. After placing the cap on the flask, grasp around the neck-covering section of the foil and pull it smoothly downward until the aluminum foil has semihermetically closed with the mouth section of the flask. Press the aluminum foil around the neck-zone of the flask to ensure the cap is tightly closed. Using saran wrap or parafilm seal junction between cap and flask, as well as the flask’s neck and mouth section. Wrap bottom up, and then bottom down (two layers).

• 19. When labeling test tubes or Erlenmeyer flask, first stick the label onto a piece of saran wrap and then label the container by wrap** the plastic wrap around the neck of the Erlenmeyer flask or test tube. This prevents the label from sticking directly on the container and its subsequent removal and washing is simpler. Place the label from the original container onto the new Erlenmeyer flask or test tube to which the material is transferred to, as a control label, along with a new printed label. This helps to track correct labeling in each multiplication cycle.

• 20. Check maximum capacity (weight) of orbital shaker and avoid overloading. Distribute flasks equidistantly on the shaker platform to ensure smooth rotation without misbalancing the shaker. Periodically, e.g. every 6 months, perform preventive maintenance of the shaker, which should include replacement and/or adjustment of motor belt, assessment of motor performance, check soldering points and components of mainboard and lubrication of axis when required. At CIP, we have manufactured and installed a larger and lighter aluminum platform to the orbital shaker (60 × 60 cm), which permits placing up to 80–100 Erlenmeyer flasks (50 mL) per shaker. High incubation temperatures of 28–29 °C speed up growth and development of in vitro plants in liquid culture medium.

• 21. Temperature, light intensity, photoperiod, and relative humidity can be controlled and recorded using an environmental measuring/control system or device. An easy, efficient, and cheap way to measure environmental variables is using mobile data loggers with external temperature, humidity, and light intensity probes. The interval for recording the data points can be adjusted individually and downloaded remotely (wireless) or through direct cable connection with a desktop PC. Advanced systems can send automatic alarms by email or text message, when the predefined maximum and minimum values of the environmental conditions are overpassed.

• 22. When in vitro plants have developed minimum one root and three to four new nodes (or leaves), they can be transferred to solid culture medium. Stem should show robusticity and not be excessively elongated (avoid long distances between nodes). Transfer plants as soon as possible to solid medium (genotype-dependent).

• 23. Before removing the aluminum foil cap from the Erlenmeyer flask, first remove the sealing, and then carefully loosen the basal and central part of the cap to facilitate easy opening. Grab the foil cap at its basal part during opening, and don’t touch the neck or opening of the flask. During transfer, hold roots and stem together and avoid loose roots and leaves that could come into contact with external vessel surfaces. Introduce basal section of the stem ~1.5 cm into solid culture medium (roots should completely be submerged in the medium). Plants should be located equidistantly in the tube, i.e. the distance between the two plants and the test tube wall should be the same.

• 24. Before transferring explants to fresh liquid medium, perform 2–3 small cuts in the base of the explant to stimulate root formation. Sweetpotato accessions that do not grow well in liquid medium can be successfully propagated in deep petri dishes (Ø: 90–100 mm, height: 25 mm) on solid culture medium (50 mL/petri dish). Place 10–36 uni-nodal explants per deep petri dish and incubate for 10–18 days (genotype-dependent) at 24 ± 1 °C, photoperiod of 16 h/8 h of light/darkness and a light intensity of 100 ± 20 μmol/m²/s. After 14–20 days, transfer rooted plants to 25 × 150 mm glass test tubes (2 plants/tube) and incubate at the same environmental conditions.

  Ensure in vitro plants submitted to a new propagation cycle in liquid culture medium are fully rooted and have developed a robust stem and minimum 5–6 nodes or leaves. Do not use in vitro plants with anormal aspect (yellowing, bad growth, deformation, hyperhydration, etc.) for a new propagation cycle.