Manual for TC Cassava Sweet Potato

Procedures for Recovering Cassava and Sweet potato Germplasm Distributed In vitro

Roca, W.M., Rodriguez. J.A and Roa. J

Genetic Resource Unit CIAT Apartado Aéreo 6713 Call, Colombia S.A

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Table of Contents

Introduction 1Recovery of the Cultures 1MicropropagationPottingField TransplantationGreen house transplantation

Appendix 1. Preparation of the Culture Medium

Appendix 1a. Preparation of Murashige and Skoog Stock Solutions

Appendix 1b. Preparation of Growth Regulator Stock Solutions

References

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1.0 Introduction

Invitro methods have been developed at CIAT for the international exchange of cassava (Manihot esculenta Crantz) clones. Sterile cultures in artificial nutritive media are established from disease-free mother plants produced by means of thermotherapy and meristem-tip culture and tested for known cassava viral, bacterial, and fungal pathogens. The cultures for shipment consist of well-rooted plantlets in an agar medium contained in properly capped1 6 x 125 mm test tubes. The test tubes, in turn are packed within polystyrene boxes.The tubes are labeled with the clone’s name or number, a phytosanitary certificate, issued by the Colombian authorities, and a phytosanitary statement which provides supplementary information regarding the health status of the cultures, are included in the package.

Several countries have accepted the invitro system as a means to improve the phytosanitry aspects of the international exchange of cassava germplasm. In the last few years, the technique has been used not only to distribute selected germplasm from CIAT to national programs, but also to introduce into CIAT large numbers of new germplasm collected in the crop’s major centers of variability. Furthermore, because of their small size and their disease-free condition and high propagation potential, meristem-derived cultures have been used at CIAT to develop an in vitro gene bank of cassava.

The in vitro system is only effective as a tool for germplasm exchange. If it is managed by well-trained personnel, these guidelines are provided to acquaint the personnel of national programs and other collaborators with the procedures to recover, propagate, and transplant materials from the cultures after arrival at the recipient institution.

1.1 Recovery of the Cultures

It is important to make the necessary arrangements to expedite clearance of the package through customs as soon as it has arrived. Recipients will be advised on the approximate arrival date of the shipment.

1. Upon their arrival, unpack the test tubes carefully and place them on test tube supports, in an upright position.

2. After short trips top to 1 week, the planlets can be potted directly (section 4) or can be micropropagated prior to potting (section 3) if you have the facilities and personnel

3. If the shipment has taken 1-2 weeks to arrive at its destination, you should expose the cultures to an Illumination of 2000 in two 40 W fluorescent lamps at 0.50 m above the cultures and a temperature of 26-28°C for 1 week. This will allow the plants to recover from the detrimental effects of darkness. After this recovery treatment, the cultures are ready for direct potting (section 3) or propagation and potting (section 2)

4. After longer trips (more than 3 weeks) and depending on genotype, etiolation and chlorosis, and sometimes tissue necrosis due to phenolic oxidation, can occur with variable intensity due to the lack of light. You can save extremely deteriorated cultures. If any available green bud (terminal or auxiliary) is aseptically excised and cultured on a fresh medium (step B). Immediately upon unpacking, expose less damaged cultures especially those with little or no shoot browning to temperatures of 22-24°C and 1000 lox of illumination for 2 weeks. Even a slight growth of auxiliary buds, with or without leaf growth, is sufficient to proceed to the micropropagation step.

1.2 Micropropagation

Micropropagation is carried out within a laminar flow cabinet or in a transfer room for aseptic work. Make sure to have on hand:

two No.10 scalpels, with their handles two forceps: one short (12cm) and one long (25 cm) several petri dishes containing 3-4 sterilized filter papers 2-3 flasks containing sterile distilled water an alcohol burner cotton wetted with 70% alcohol 2-3 flaps (10 x 20 cm each) of sterile filter paper a container with 95% ethanol

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The culture medium includes the mineral salts of Murashige and Skoog + 2% sucrose + 1 mg/1 thiamine HCl = 100 mg/l Inositol + 0.05 mg/I benzyl aminopurine + 0.05 mg/I gibberellic acid + 0.02 mg/l naphathalene acetic acid; Ph 5.7-5.8; agar = 0.6%. The preparation of this medium is described in Appendix 1

Figure 1 micropropagation of cassava through nodal cuttings.

1.2.1 Procedures for micropropagation

1. Wash hands and arms with soap and running water and wipe with 70% alcohol. If possible, cover the hair with a microbial cap and the mouth and nose with a microbial mask

2. Immerse the tools in 95% alcohol for 2-3 minutes; flame quickly, and place them under the sterile flaps of paper.

3. Open one petri dish and wet the filter papers with the sterile water taking care to flame the flask. One at a time, uncap 2-3 test tubes (Figure 1-a), and using short forceps, gently pull the plantlet from the tube. Place all planlets within the petri dish (figure 1-b)

4. Using a scalped and the short forceps cut off all expanded leaves and roots. Then, cut the stem into segments, each one comprising one node (Figure 1-e). Every segment or “nodal cutting” will then include one auxiliary bud and a portion of stem (1-2 mm above and 4-5 mm below the bud, respectively). In addition, cut the terminal bud from every shoot, leaving 4-5 mm of storm below the apex. Make sure to make clean, horizontal cuts with the scalpel.

5. Using the larger forceps, gently pick up each “nodal cutting” and each terminal bud, and “ plant” each one in agar solidified medium (Figure 1-d) in an 18-25 = X 150 MM TEST TUBE. Quickly flame the tube and cap it immediatelyDuring the operation, make sure to frequently wet the tools with 95% alcohol and flame them. Keep the petri dish closed as much as possible.

6. Incubate the “node cutting’ cultures at 28°C under an illumination of 2000 lux until root initiation and, thereafter, under 4000 lux, and maintain a 14 hour photoperiod throughout.After about 3 weeks, every modal cutting will give rise to a complete planlet (Fig 1-e). Once, the plantlet has grown to 6-8 cm tall and rooting is proportional to the amount of shoots, the cultures can be subjected to a hardening treatment prior to potting. Hardening is not absolutely necessary, but it will improve the potting.

7. For hardening, increase the illumination to 8,000-10,000 lux and, if possible, decrease the temperature to 24-25°C during 1-2 weeks

8. Three to four days prior to potting, take the paraffinated paper off the caps, whether or not you have hardened the cultures.

1.3 Potting

For potting, the following items are needed: Pots, Optimal potting is achieved using 3 to 4 inch jiffy-type pots, but clay or plastic bags 96.5 x 8

inches) can also be used Substratum, which consists of a mixture of one part [soil with three parts of fine sand. The sand must

be thoroughly washed with soft water. A fair amount of deionized water

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Long (15-20 cm) forceps Pot labels A water –soluble fertilizer rich in P; e.g. 10-52-10 (N-P-K) Plastic, clay or cement trays with sufficient draining holes, containing damped sterilized soil. A 2.0 X 10- X 0.3 –m transparent plastic frame to cover the trays A 2.0 X 1.0- X 0.8-m greenhouse table to hold the trays and the plastic frame The two or three tables of similar dimensions to carry out the potting and to hold the pots prior to field

transplanting Styrofoam cup You should do the potting in a fresh, but not too cold location that is protected from direct sun and

from insects.

1. A few hours prior to potting, damp with water the soil contained in the trays, place the trays on the greenhouse table and cover them with the plastic frame (Fig 2). A high relative humidity environment will form within the plastic chamber

2. Sterilize the substratum mixture using steam and fill in the pots. Wet the substratum to about one half water saturation. Make a hole in the center of each pot (Fig 3)

3. Wash your hands thoroughly with soap and water. Uncap the test tubes, one at a time, and, with the aid of the forceps, gently pull out the planlet (Fig 4). Using the fingers, pull the plant from the tube mouth.

4. Holding the plant on the palm of your hand, wash off as much agar as possible from the roots (Fig 5), and place the roots and the lower part of the shoot within the hole in the pot ( Fig 6). Press the substratum and immediately apply deionized water around the plant (Fig 7)

5. After potting label the posts and water with the high P fertilizer 10-52-10. Use 2g fertilizer in 1 litre of water and apply 80-100 ml solution per pot. Thereafter, when needed and until the time of field transplanting, water with only the high P solution.

6. Place the pots in the high humidity chamber (Fig 8). Five or six days after potting and thereafter raise the plastic frame slightly during the fresher hours of the day, close the frame at night. By day 10 or 12, the plastic frame can be taken off completely.

7. After the 12th day, move the pots (still on the trays) to a warmer and much more illuminated part of the greenhouse. In about 2 more weeks, the plants are ready for field transplanting.

The total time taken from receipt to this stage usually is 1 or 2 months depending on whether direct potting or micropopagation and potting were carried out respectively.

1.4 Transplanting in the greenhouse

MATERIALS

Black plastic bags for planting Sterile planting media sand/soil ratio of 3/1 Transparent plastic cups Labels A greenhouse with temperature between 26 and 28 degrees for keeping only clean material In vitro plants in 17N media (enraizamiento) that are 6-8 cm long with maximum of 30 days after

sub-culturing1. Prepare planting media a week before transplanting in the following way: - wash the sand 3 times

until it is totally clean, put to dry in the sun, sieve soil, mix and sterilize for 3 hours.2. The plants should be in the screen-house for a minimum of 2 days before transplanting for them to

adopt.3. Fill the bags for planting one day before4. On planting day add to the planting media a solution of Banrot 0.75gr/lt, or a fungicide containing

active ingredients against mushrooms (Phytophthora, Phytium) bacteria. This should be prepared with deionized or sterile distilled water.

5. A lot of care should be taken not to damage the roots, add little water to the flash to help loosen the agar for the plants to be removed easily. Make a hole in the planting media and insert the plant.

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6. A flask contains 5 plants, each is placed in a different bag, with its respective label, and a cup is placed over it to increase humidity to protect it for the next 8 days. This cup should preferable be transparent to allow in light.

7. After transplanting the material should be inspected daily, without removing the plastic cups to avoid dehydration, and watered only when necessary.

8. Remove the cups after 8 days and water if necessary using deionized water and remove dead leaves and initiate fertilizer application.

1.4.1 FERTILIZER APPLICATION

Fertilizer application is very important because it gives to the plants nutrients that it needs for its growth, due to this the fertilizers are in two groups according to its functionality.

1. Fertilizers with micro nutrients that the plant needs in its initial stage: Plan/pro (plantex), cosmocel, coljap development and coljap production each one of them is used in a concentration of 0.5%, The major micro nutrients the fertilizers have to have are Calcium, Zinc, Magnesium, Sodium, Manganese and Cobalt.

2. The fertilizers with macro nutrients that the plant needs at latter stage: N/P/K and the urea (little it is used) one of them is used to a concentration of 1% with Nitrogen as a major component for foliar growth

Fertilizer application is carried out every 8 days, alternating the fertilizer and precious amount of 50ml should be done I the morning 8-10.00 am or later in the day 5-6.00 pm when it is cool so that the plants can assimilate most of the fertilizer.

First application should be done 15 days after transplanting and should be applied to the soil. After apply every 8 days. Plantex/Plan-pro (apply in the soil) Then coljap development (apply in the foliage) Coljap production (apply in the foliage) Cosmocel (apply in the soil) Triple 15 (N/P/K) (apply in the soil) Urea (apply in the soil)

NOTE;

Sometimes it is necessary to apply fungicides and pesticides to prevent fungal and/or pests. Apply 0.7 ml/l fungicide or 1 ml/l of acaricide. These should not be done simultaneous with the fertilizations as the plant may get burn because of excess chemical

1.5 Field Transplantation

Successful transplanting can be achieved under cloudy days or during the late afternoon. The soil should be in field capacity and the plants 10-15 cm in height. Use jiffy-type pots or plastics bags and keep root disturbance minimum

1. Carry the plants in their pots, along with the trays, to the field and cut off the largest leaves2. Remove the pots from the tray and place them within the hole large enough to hold the pot and up to

the loved two nodes of the shoot3. Press the soil around the plant and water immediately4. Maintain high soil humidity for 10-15 after transplanting, and watch out for ant, worm or cricket

attacks.After planting, cover each plant with a Styrofoam cup (Make sure to make some openings on the upper side of the cup). Leave the plants covered or 1 week, then uncover them once continue with stopAt the fourth week, the plants should be completely established in the field and can be handled conventionally. However, it is recommended that you pay special attention to protecting the plants from insects or disease

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throughout the crop. Keep in mind that these plants are elite stocks for the multiplication of clean planting material and that their production shipment, and handling after arrival have been done with great care and effortFor news regarding the condition of the cultures upon arrival, any progress developments in handling or additional information, please write to

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APPENDIX 1

Preparation of the Culture Medium

A) Basal Medium. Can be prepared in either of two forms:

1. Using stock solutions of mineral salts, vitamin and growth regulators. For the preparations of the Murashige and Skoog stock solutions see (Table 1)

To prepare 1 liter of medium, to 500 ml of double distilled water add:20.0 ml of stock solution No.11.0 ml of stock solution No.21.0 ml of stock solution No.32.9 ml of stock solution No.45.0 ml of stock solution No.5

2. Using the pre-made Murashige and Skoog medium in powder form (without sucrose and without vitamins and agar). Each bag contains 4.3g of powder, which serves to prepare 1 litre of basal medium. The powder can be stored at 8-10°C or under desiccation, for up to 2 years. To prepare the basal medium dissolve the entire contents of one bag in 500 ml double distilled water. Add the same volumes of stock solutions No.1 through No.5 as in Table 1.

B). Supplements. Once either of the basal media is ready, proceed as follows:

Add 5.0 ml of stock solution No.6 and 6.25 ml of stock solution No.7 Dissolve 20.0 g of sucrose Add 5.0 ml of the benzyl aminopurine (BAP) stock solution and 2.0 ml of Naphthalene acetic acid

(NAA) stock solution Complete to 700ml with double distilled water Adjust the pΗ to 5.7-5.8 Dissolve by heating 6.0 g of agar in 300ml of double distilled water Mix well the medium with the agar solution.

C. STERILIZATION

Quickly distribute the prepared medium in 18-X.150-mm test tubes (5ml/tubes); let cool slightly and cap the tubes

Autoclave the tubes containing medium; 15 pounds (121°C) per square inch during 15 minutes; decompress slowlyPlace the tubes in a fresh place until the agar solidify, then store them in darkness at 6-8°C until used.

Appendix 1a. Preparation of Murashige and Skoog Stock SolutionsTo prepare the stock solutions, dissolve one by one, all the ingredients presented in Table 1, in the volumes of double distilled water shown.


Benzyl aminopurine (BAP)(10 ppm):

Dissolve 20 mg in a small volume of 1.0 N HCl; complete to 200 ml with double distilled water (This is a 100-ppm solution of the growth regulator): Take 20 ml of the 100-ppm solution and complete to 200 ml (This is the 10-ppm stock solutions).

Gibberellic acid (GA3) (10 ppm): Dissolve 22 mg (90% gibberellic acid) in a small volume of 1.0 N KOH; complete to 200 ml with water; take 20 ml of this solution and complete to 200 ml with water.Naphathalene acetic acid (NAA) (10 ppm):

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Dissolve 20 mg in a small volume of 1.0 N KOH, complete to 200 ml with water; take 20 ml of this solution and complete to 200 ml with water

Table 1. Murashige and Skoog Stock and Medium Preparation.

Stock solution no.a

ConstituentsSubstance Amount

Volume of stock per 1 liter basalmedium

1 NH₄NO₃KNO₃MgSO₄. 7H₂OKH₂PO₄Dissolve in 1000 ml water

82.5 g95.0 g18.5 g8.5 g

20.0 ml

2 H₂BO₃MnSO₄ . H₂OZnSO₄ . 7H₂ONa₂MoO₄ . 2H₂OCuSO₄ . 5H₂OCoCl₂ . 6H₂ODissolve in 100 ml water

0.62 g2.176g0.86 g0.025 g0.0025 g0.0021 g

1.0 ml

3 KlDissolve in 100 ml water

0.075 g 1.0 ml

4 CaCl . 2H₂ODissolve in 100 ml water

15 g 2.9 ml

5b a) Na₂EDTAb) FeSO₄ . 7H₂O

Dissolve in 200 ml water

1.492 mg1.114 mg

5.0 ml

6 Thiamine-HClDissolve in 200 ml water

0.8 g 6.25 ml

7 m-inositolDissolve in 200 ml water

0.8 g 6.25 ml

a. Stocks 2 and 6 should be kept frozen; all the others at 8-10°C. Keep stock 5 protected from lightb. Separately dissolve ‘a’ and ‘b’ in 50 ml with water; take 20 ml of this solution and complete to 200 ml with

water.

Addition of Growth Regulator Stocks to the Medium: To determine the volume (Appendix 1) of each growth regulator stock solution necessary to obtain the prescribed concentrations (step B), apply the following formulation:

C₁V₁ = C₂V₂C₁ = Concentration of stock – 10 mg/l

C₂ = Final concentration of growth regulator in the mediumBenzyl aminopurine = 0.05 mg/lGibberellic acid = 0.05 mg/lNaphthalene acetic acid = 0.02 mg/l

V₁ = Volume (in ml) of stick solutions needed = X

V₂ = Final volume of medium = 1000 ml

X= 0.05mg/l x 1000ml = 5.0 ml of either BAP/GA3 10mg/l

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References

1. CIAT (Centro International de Agricultura Tropical). 1983. Elite Cassava germplasm from CIAT. Cali, Colombia.20 p

2. Hewitt, W.B and Chiarappa, L (eds) 1977. Plant health and quarantine in the international transfer of genetic resources. CRS Press, Cleveland, Ohio.

3. IBPGR (I international Board for Plant Genetic Resources) 1983a. Genetic resources of cassava and wild relatives. IBPGR Secretariat, Rome, 56 p.

4. 1983b. Practical constraints affecting the collection and exchange of wild species and primitive cultivars. IBPGR Secretariat, Rome, 11 p.

5. Lozano, J.C; Belloti, A. Reyes, J.A; Howeler, R; Leihner, D; and Doll, J. 1981. Field problems in cassava. Centro Internacional de Agricultura Tropical, Cali, Colombia. 192 p.

6. Murashige, T and Skoog F.1962. A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol. Plant. 15:473-497.

7. Roca, W.M. In press. Cassava. In: D.A Evans; W.R. Sharp; P.V. Ammirato; and Y. Yamada (eds), Handbook of plant cell culture. Vol. 2: Crop species. MacMillan, New York

8. Schilde, L and Roca, W.M.In Press. Pathogen elimination in potato and cassava. In: J. Cock (ed), Propagation of tuner root crops. Centro Internacional de Agricultura Tropical, Cali, Colombia.

9. Tery, E.R 1982. A review of cassava and sweet potato diseases and their relation to germplasm exchange. Research Briefs, Vol.3 International Institue of Tropical AGRICULTURE, Ibadan, Nigeria, 4 p

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Micropropagation and Maintenance

Rolando Lizárraga, Ana Panta, Nelson Espinoza,John Dodds

(Reproduced from CIP Research Guide 32)

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Table of Contents

Introduction 1Advantages of tissue culture 1Introduction of in vivo material into InvitroMeristem isolation culturesThermotherapyField TransplantationGreen house transplantation

Appendix 1. Preparation of the Culture Medium

Appendix 1a. Preparation of Murashige and Skoog Stock Solutions


References

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Tissue culture allows the rapid clonal propagation of a large number of plantlets over a short period, as well as the maintenance of germplasm under controlled conditions in small spaces and with reduced labor

requirements.

This document complies the advances, methodologies and materials used for tissue culture at the various institution including International Potato Center (CIP), Mikocheni Agricultural research Institute etc. It analyzes isolation, micropropagation, and long term storage techniques.

Advantages of tissue culture

CIP maintains a sweet potato germplasm collection of over 5,000 accessions. Their clonal maintenance on the field is expensive and involves the risk of loss due to infectious diseases or unfavorable climatic conditions. Thus, in vitro maintenance presents the following advantages:

Lower labor costs, Absence of field infections,

Protection against unfavorable climatic conditions,

Timely access to material under maintenance,

Timely access to material for pathogen clean-up,

Permanent availability of (when pathogen tested) material for propagation and exportation.

Introduction of in vivo material into in vitro

The explants from the mother plant must come from the greenhouse. Alternatively mature plants directly from the fields normally are very dirty and are difficulty to thoroughly cleaned them, read for initiation. They can be re-established in the green house for 2-3 month with frequent spraying with insecticides and acaricide to eliminate the ectoparacites such as aphides white flies or mites. Plantlets of 2-3 months old, which are in excellent health conditions, and free of lateral buds that are, too sprout are best for initiation. The stems are excised from the mother plant and the leaves removed. Part of the petiole should be left to cover the buds. Stem cuttings 2 to 3 cm long each must include an axillar bud and a portion of the internodes behind the bud for ease of manipulation.

Presurface sterilization

Before sending them to the in vitro laboratory, the stem cuttings are treated with a wide spectrum acaricide to destroy the mites at any stage of development. This can be achieved by any commercial acaricide for few minutes. Then acaricide is rinsed away by washing the stems in running water for 30-45 minutes. They are then placed in a clean bottle/beaker covered with a Petri dish/aluminum foil until starting surface disinfection.

To start surface disinfection of the stem cuttings, remove the water from the container, add 96% ethanol and let stand for two seconds. Discard the alcohol and immediately add a 2.5% solution of calcium hypochlorite (brought to pH 8 by addition of HCl). Sodium hypochlorite or chlorine (bleach) may also be used. If possible, add a few drops of a dispersing-adhering agent such as Tween 20 or 80 (4 drops/l of solution). The bottle is then placed in the laminar flow transfer chamber. After 15 minutes under sterile conditions, the hypochlorite is eliminated, by washing three times with sterile water. To reduce phenolization of the explants, after the last

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rinse they are left in a sterile 100 ppm solution of ascorbic acid before proceeding to excision. Blot the explants into a filter paper to dry off the hypochlorite solution.

Under these conditions, proceed to excise the buds by eliminating the largest possible number of leaflets and leaf primordia. The excised portions should be as small as possible. The optimum size is 0.6 mm but the explant may be bigger if no adequate excision instruments, such as a stereoscopic microscope, are available.

The buds/shoot tips are initiate in initiation media and sealed with parafilm and incubate in the growth room with required environment for 4 weeks. In the first propagation stage, 16 x 125 mm culture vessels are used. In the second stage, either 18 x 150 mm or 25 x 150 mm tubes may be utilized.

In a situation where cultures are initiated using meristem, Due to the fact that meristematic portions larger than 0.6 mm are used, saprophytic bacteria mainly may contaminate some material. If so, proceed in either of two ways.

1. If a stereoscopic microscope is available, excise meristems between 0.4 and 0.6 mm long, plant them in the introduction medium (MMB-I) and transfer them weekly to the MMB-II medium.

2. Eradication of bacteria or yeasts may be attempted by adding antibiotics to the medium. For bacteria it is recommended to use Rifampicine (Rimactan 300 CIBA) at 400 ppm.

A concentrated (12,000 ppm), filter-sterilized solution of Rifampicine is used to soak small (5 x 5 mm) squares of filter paper, which are allowed to dry in the laminar flow chamber. Approximately 0.03 cm3 of concentrate solution is used for every square of filter paper. They are best if used before seven days for they progressively lose they effectiveness. This process must be carried out under aseptic conditions. The paper squares are introduced into the culture medium with the planted bud, which must be transferred to a fresh medium with another antibiotic paper every 3 to 5 days. Other antibiotics such as Cefotixine (Mefoxin Merck) in doses of 500 ppm may also be used.

For contamination by yeast it is advisable to use 0.25 to 0.5 ppm doses of Amphotericine B. The filter paper procedure described above will be used in all instances.

1. Meristem isolation and culture

The meristem is a tissue made of cells under division and is the active growth point of the bud. The dome of the bud contains the meristematic cells and is surrounded by foliar primordia and primary leaves. The meristematic cells divide and form new tissue. Nutrition of the dissected section is provided by the artificial medium. The isolation of the meristematic zone under aseptic conditions and its culture in an adequate nutritive medium allow plantlet development with a differentiation pattern similar to that of a normal plant.The aseptic dissection of the meristem for virus eradication is a delicate process requiring skill.

Photo 1 shows a photographic sequence of the dissection procedure.

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1.2.3. The plant stems are cut in segments including a node and the corresponding axillar bud. The material

is deeped into a disinfectant and sodium hypochlorite as for the initiation of invitro cultures. 4. After rinsing in distilled sterile water, place the material under the dissection microscope and use a

needle or surgical knife to remove the leaves around the growth point until only the cupule and two or three foliar primordia are left.

5. The cupule and foliar primordia are dissected with the surgical blade and transferred to culture medium MMB-I. The dissected meristem is transferred weekly to a fresh MMB-II medium. After 6 to 8 weeks the meristem will develop into a plantlet. The plantlets are now ready for subculture in the propagation medium (see section on Culture Media).

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QuickTime™ and a decompressor

are needed to see this picture.

2. Thermotherapy

At CIP before excision of the meristems, the plants undergo a month thermotherapy at 38 C for 16 hours and at 32 C for eight hours under constant light conditions. This high temperature treatment has increased the efficiency of the production process of virus free material. After thermotherapy either axillar or apical meristems may be used indistinctly.

Photo 1. Photographic sequence of meristem dissection:

A: Isolated and disinfected apical budB: Dissection after removal of primary leavesC: Meristem with two foliar primordia

Micropropagation

The purpose of micropropagation is to obtain a large number of clonal plants in a short time. The following methods are used at CIP:

Propagation by nodes

This is based on the principle that the node of an in vitro plantlet placed in an appropriate culture medium will induce the development of the axillar bud resulting in a new in vitro plantlet. It must be noted that this type of propagation is based on the development of a pre-existing morphological structure. The nutritional-hormonal condition of the medium plays a simple role in breaking the dormancy of the axillar bud and promoting its rapid development. The propagation medium described in the section on Culture Media is used.

Callus formation and plant regeneration must be carefully avoided because they tend to affect the genetic stability of the genotype.

The plantlets grow under long day conditions (16 hours of light at 45 m E/m2/seg2 or 3,000 lux) and at temperatures ranging from 25° C to 28° C. Under these conditions micropropagation is fast. Each node will develop into a plantlet occupying the full length of the test tube. The plantlets will be ready for subculture after six weeks (Photos 2 and 3).

The resulting sweetpotato in vitro plantlets may be easily transplanted to in vivo conditions either in small pots or directly to field beds (Photo 4).

Propagation by stem cuttings in liquid medium

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http://www.cipotato.org/training/materials/sweetpotato/images/032.jpg

As with the potato, it is possible to micropropagate sweetpotato in bottles containing a liquid medium (Photo 5). Stem cuttings with 5 to 8 nodes are prepared by removing both the apex and root of the plantlet to be propagated. The cuttings are placed in a liquid medium containing gibberellic acid to break the dormancy of the stem cutting’s axillar buds. The nodes will sprout and new plantlets develop over a period of 3 to 4 weeks. The plantlets may be used as initial material for simple node propagation or once again for propagation with stem cuttings in liquid medium, depending on program needs. Shaking of cultures may accelerate and promote the development of new plantlets. However, this is not essential.

Photo 2. Plantlet growing from a node

Photo 3. Sequence of in vitro development

Photo 4. Plantlets after transfer to peat-moss pot

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a: Homemade peat-moss pot (newsprint)

b: Commercial peat-moss pot (jiffy 7)

Photo 5. Liquid medium culture for rapid propagation

Long term maintenance

Long term maintenance is important for propagation and conservation itself. For clonally propagated cultures it is important that every propagule be free of even the slightest genetic alterations that may build up from one generation to the next and result in major changes affecting uniformity and production.

For the maintenance of germplasm clones, it is crucial to conduct a detailed analysis of the culture’s genetic stability. The clonal storage of germplasm consists on the maintenance of specific gene combinations (genotypes). If a plantlet’s genetic combination changes during storage, the validity of the storage techniques must be examined. The capacity to detect genetic changes during propagation and maintenance depends on the methods used.

In many germplasm collections the stored genotypes are routinely evaluated for the morphological characteristics of the plantlets growing under controlled conditions. If the plants show new morphological characteristics (e.g. leaf shape, or storage root color) some genetic changes are obvious. However, genetic changes such as virus resistance may not be detected through the observation of morphological changes.

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Biochemical methods are now used to study the genetic stability of both potato and sweetpotato. They are the analysis of soluble protein and isoenzyme patterns. Although these methods are highly effective in determining changes in genetic products, they do not allow directly to determine changes in the genes.

New methods such as Restricted Fragment Length Polimorphism (RFLP) are considered to be more sensitive ways of determining genetic changes. It is important that germplasm banks and seed programs use the most effective methods in determining the genetic fidelity of the propagation and maintenance systems.

Growth restriction media

After many years of research, propagation media for sweetpotato have been developed that optimize rapid in vitro growth. However, maintenance requires limiting growth to a minimum while maintaining culture viability. Use of growth restriction media allows to maximize the interval between transfers (subcultures) of in vitro plantlets. At CIP, transfer of most sweetpotato material under maintenance is carried out once a year, and in some cases once every year and a half.

Laboratory experiments aimed at limiting in vitro growth of sweetpotato include the use of hormonal growth retardants such as abscisic acid (ABA), growth inhibitors such as B995 or chloride chloride (CCC), as well as osmotic regulators with addition of low assimilation sugars such as manitol or sorbitol.

The difficulty involved in this type of study is that under these conditions genotypes react differently. Studies of germplasm collections in vitro maintenance should aim at developing maintenance media broad enough for a large variety of genotypes.

Also, the storage medium should not allow callus induction that may result in genetic alterations.

Many storage methods have been reported for sweetpotato. At present, the method described in the next section is used at Cip.

Restriction of storage temperature the growth of in vitro plantlets may be restricted reducing incubation temperature. Adequate in vitro growth of sweetpotato can be obtained with temperatures between 28° C and 30° C. At 8° C, survival time is less than one month. For genotypes studied to date, 15° C seems to be the optimum temperature. However, this has to be confirmed yet.

As with other in vitro cultures such as cassava and potato, low temperature and growth retardants may be used simultaneously. For now, the combined use of osmotic stress and low temperature (15° C) appears as the best and least costly way of maintaining sweetpotato germplasm collections.

Culture media used in the work reported here are based on Murashige-Skoog (1962) and Gamborg B-5 (1968) salts.

Medium for initiation

Calcium pantothenate 2 ppm Gibberellic acid 20 ppm

Ascorbic acid 100 ppm

Calcium nitrate 100 ppm

Putrescine HCl 20 ppm

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L-Arginine 100 ppm

Coconut milk 1%

Sucrose 5%

Agar or 0.7%

Phytagel/Gelrite 0.25%

Medium for transfer of meristems or buds (MMB-II)

Calcium pantothenate 2ppm Gibberelic acid 15 ppm


Calcium nitrate 100 ppm


L-Arginine 100 ppm

Saccharose 5%

Agar or 0.7%


Propagation medium (MPB)

Calcium pantothenate 2 ppm Gibberelic acid 10 ppm

L-Arginine 100 ppm



Sucrose 3%

Agar or 0.8%


Maintenance medium (MCB)

Glucose 2% Sorbitol 2%

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A pH 5.8 is used in all media.

Note: These culture media were prepared to attain maximum uniformity in a collection including a large number of varieties. When few varieties are involved, it is advisable to use simple culture media such as Murashige and Skoog salts with the addition of gibberellic acid and sucrose.

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Cryotherapy protocol for pathogen elimination from Sweet potato and Cassava

Introduction

Although several protocols for cryotherapy of shoot tips of sweet potato have been published

(Towill and Jarret, 1992; Hirai and Sakai, 2003; Wang and Valkonen, 2008a), encapsulation-

vitrification described by Hirai and Sakai (2003) and Wang and Valkonen (2008a) proved to be

efficient for sweet potato. Therefore, encapsulation-vitrification is described in detail as

follows:

Procedures for Cryopreservation of Sweet potato shoot tip by encapsulation vitrification

1. Initiate stock cultures of sweet potato on MS medium containing 30 g/l sucrose, 2.6 g/l

gelrite (pH, 5.8) and grow them in the growth room with 24-280C and 16 hrs light for 4

weeks to get good shoots

2. Excise shoot tips (1-1.5 mm) from 4-weeks old stock cultures and encapsulate them with

3% Na-alginate solution in 0.1 M CaCl2 solution for 20 min to form beads (4 mm in

diameter), each bead should containing one shoot tip

3. Preculture beads in liquid MS medium containing 0.3 M sucrose for 16 h on a shaker (90

rpm)

4. Transfer the beads into liquid MS medium containing 1.6 M sucrose and place the

cultures for 3 h on a shaker (90 rpm). After shaking of 3 h surface-dry the beads by place

them on sterilized filter paper inside laminar flow;

5. Transfer the beads into PVS2 solution and vitrify the beads for 60-90 min by shaking on

orbital shaker and surface dry them after 90 min.

6. Transfer the beads into 1.8-ml cryovials, each cryotube containing 10 beads, and directly

immerse the cryovials into liquid nitrogen for 1 h

7. Quickly transfer the cryovials in a water bath at 40oC for 3 min;

8. Empty the beads into liquid MS containing 1.2 M sucrose for 20 min at room

temperature and surface-dry the beads as in step 4

9. Post-culture the beads on solid NH4-free MS containing 0.5 mg/l BAP at 24-28o C in the

dark for 7-10 days

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10. Transfer the beads onto solid MS medium containing 10 mg/l GA3 and cultured under

light at 24-28 o C until the shoot tips develop shoot. Subculture is done once every 4

weeks.

Fig. 3. Flow chart of cryotherapy of shoot tips by encapsulation-vitrification (Wang and Perl, 2006)

shoot tip Encapsulation Precultureee

Loading

ThawingUnloadingSurvival

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In vitro stock shoots

Vitrification

FreezingRegeneratio

Protocol for cryotherapy of cassava shoot tips by vitrification

(Charoensub et al., 2003) is presented as the followings

1. Initiate invitro stock of cassava onto MS + 0.02 mg/l BA + 0.1 mg/l GA3 + 0.01 mg/ NAA + 30 g/l

sucrose (pH=5.6) solid media and grow them into a growth room for 8 weeks

2. Excise a nodal segments (one lateral bud, 0.5 cm long) and culture on MS + 0.02 mg/l BA + 0.1

mg/l GA3 + 0.01 mg/ NAA + 30 g/l sucrose (pH=5.6) for 12 days

3. Excise shoot tips of (1 mm with 2 leaf primordia) under the microscope and preculture in MS +

0.3 M sucrose for 16 h

4. Transfer 10 shoot tips each into 1.8 ml cryovials containing MS + 0.4 M sucrose + 2 M glycerol

for 20 min (You can pick up the ST with sterile tweezers and drop them into cryovials together with loading

buffer into vials)

NOTE: From these stage to last stage is critical important you prepare all the required matariaal for eash step just

before the end of each step)

5. Vitrify (Osmoprotection stage) the shoot tips in 1.8 ml cryotubes by adding 1 ml of PVS2

solution and incubate at RT for 45 min.

6. After 45 min decant half on the PVS2 solution and immerse cryotubes in LN for 1h each of

cryotubes contains remaining 0.5 ml PVS2

7. Remove the cryovial from LN and quickly deep them into water bath set at 45 o C to thaw them

for 1 min

8. Decant the remaining PVS2 solution and wash the ST by incubating them into unloading solution

containing MS + 1.2 M sucrose for 20 min inside laminar flow

9. Culture shoot tips on MS +0.02 mg/l BA + 0.1 mg/l GA3 + 0.01 mg/ NAA + 30 g/l sucrose solid

media overlaid with filter paper for 4 days

10. Transfer to fresh MS +0.02 mg/l BA + 0.1 mg/l GA3 + 0.01 mg/ NAA + 30 g/l sucrose solid media

for recovery and regeneration

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shoot tip Preculture Loading Vitrification

FreezingThawingUnloadingSurvivalRegeneration

25

Fig. 5. Flow chart of cryotherapy of shoot tips by droplet-vitrification (Wang and Perl, 2006)


shoot tip Preculture

Loading Vitrification

FreezingThawingUnloadingSurvivalRegeneration

Droplets

26

Fig. 6. Flow chart for cryotherapy of shoot tips by encapsulation-dehydration (Wang and Perl, 2006)


shoot tip PrecultureEncapsulation Air-drying

FreezingThawingSurvivalRegeneration

27

AppendixFacilities and equipment requiredTissue culture facilities (Laminar flow cabinet and growth room with 24-280C 16hr light)Liquid nitrogen (LN)Water bath (400C)Petri dishes (90 mm diameter)Erlenmeyer’s conical flask (100ml)Filter papersCryovials (1.8-2 ml capacity)Pipettes 1 & 5 mlPipettes tips 1& 5 mlOrbital shaker (90rpm)

Buffer and solutions Basal medium: MS (Commercial available premix without vitamins)1g/l casamino acids30g/l sucrose + 2g/l geltrite, pH =5.7

2.5% Na-alginate solution: BM 25g/l Na-alginate0.4M sucrose2M glycerolbalance the pH to 5.7 and autoclave

0.1M CaCl2 solutionMS11.1g/l CaCl20.4M sucrosepH5.8

Preculture medium: BM + 0.25M sucrose, BM + 0.5M sucrose, BM + 0.75M sucrose; pH =5.7

Osmoprotection solution: liquid BM + 1.6M sucrose + BM + 2M glycerol, pH =5.7

Plant vitrification solution 2 (PVS2): BM 30% glycerol15% ethylene glycol15% DMSO0.4M sucrose

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Balance pH to 5.7

Washing solution: BM + 1.2M sucrose, pH =5.7

Survival medium: BM + 0.5mg/l BA + 1mg/l GA3, pH =5.7

Rooting mediaMS30g/l sucrose2.6g/l gelritepH 5.7

Regrowth mediumMS10mg/l GA330g/l sucrose2.6g/l gelritepH 5.7

survival mediaNH4-freeMS0.5mg/l BA30g/l sucrose2.6g/l gelritepH 5.7

Unloading medumNH4-freeMS1.2 M sucrose30g/l sucrose2.6g/l gelritepH 5.7

Unloading solutionNH4-freeMS1.2 M sucrose2M glycerol

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Virus indexing

Virus detection in plants regenerated from cryo-treated shoot tips is a critical step because viral

conditions can be confirmed only through this step. Various detection methods for viruses have

been developed, for example, ELISA and RT-PCR, but virus-indexing using indicator plant is cost-

effective and also reliable (Wang and Valkonen, 2008b). Therefore, this simple, efficient method

is presented here:

Fig. 7. Virus indexing by micro-grafting of cultivars tested upon the indicator plant, I. setosa (Wang and Valkonen, 2008b).

Cultivars to be tested A “V” cut at basal part of scion

A “vertical cut” at top of rootstock

In vitro grafting

Indicator

Virus indexing

30

A silicon tube is used to hold grafting point

Transport, receipt, and propagation of invitro sweet potato

plantlets

Tissue culture materials consist of small, aseptic plantlets growing on a synthetic nutrient medium. The

aseptic nature of this material makes it an ideal method for international exchange of germplasm as it

minimizes the risk of transmitting fungal and bacterial diseases. This document contains information for

the recipient of invitro plantlets on the handling procedures to be followed for further micro-propagation or

transfer to non-sterile growing conditions.

Transport of tissue culture material

1. Packing the material is packed in polystyrene in a cardboard container. Each package contains several

small glass test tubes, each with three well-developed plantlets. Extra agar media is added to prevent

damage from movement during shipment (Photo 1).Photo 1

The test tubes are capped with plastic covers and sealed with parafilm to prevent entry of contaminants

into the cultures and loss of water from the medium.

Shipment.

Whenever possible, tissue cultures are hand-carried to ensure rapid transport. When this is not feasible,

the fastest possible method is usually airfreight. In vitro plantlets can survive two to three weeks without

light.

Handling after arrival.

The cultures should be cleared from customs as quickly as possible. When advance notice of the

shipment is known, alert the customs officials of its expected arrival. Carefully remove the test tubes from

the package in a laboratory or clean room. Do not open the tubes. Do not remove the plantlets. If the

plantlets have become yellow, place the test tubes under diffused light for about 1 week in a clean room.

Invitro shoot cultures are free from diseases. Work under clean conditions according to the following

description (steps 4 and 5 of the following section on "Transfer to Planting Materials") to prevent

contamination during and after unpacking.

Use of tissue culture material.

The plantlets can be used in two different ways:

Transfer to planting mix

Micropropagated

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Transfer to planting mix

1. Materials

Peat moss Fine sand (1 mm diameter) Aluminum foil Clay pots (8 to 10 cm diameter) or jiffy pots

Larger pots (20 cm diameter) Distilled water1% calcium-hypochlorite solution70% alcohol solution

Strong soap

Fertilizer with high content of phosphorus (5-50-17) or (12-12-12)

2. Mix peat moss and sand (1:2 volume).

If an autoclave is available, fill the pots with peat moss/sand mix, cover them with aluminum foil,

and sterilize for 1 hour. If an autoclave is not available wash the clay pots (jiffy pots are already

sterile) with detergent, rinse them well with running water, and sterilize the planting mix and some

additional sand separately by any other means (heat, steam, fungicides, etc.).

Take the pots and the in vitro culture to a clean bench that is protected from air currents, dust,

dirt, insects, or other contaminants.

Wash your hands with strong soap and 1% calcium-hypochlorite solution. Then rinse hands in

70% alcohol.

Irrigate the pots with a small amount of water.

Prepare the pot that is to receive the plantlet by making a hole in the center of the peat

moss/sand mix with a clean stick or pencil.

Before removing the plantlets, disinfect the outside of the test tube using a piece of cotton or cloth

moistened with 70% alcohol to reduce the risk of contamination.

Using clean fingers, remove the parafilm and the plastic cover from the test tube. Work with one

tube at a time.

3. Gently pull the plantlets with the agar out of the test tube using sterilized forceps (flamed to red

heat and cooled) (Photo 2).

Photo 2

11. Wash the agar from the roots by gently immersing them several times in sterilized water, trying

not to wet the rest of the plantlet.

12. Plant each plant individually in the holes in the potting mix with the roots plus one or two nodes

below the surface (Photo 3).

13. Place the sterilized sand around the plantlet and press lightly to keep the plantlet straight in the

pot.

14. Place the plant into a humid chamber during 48 h. Remove the humid chamber and wait until the

roots are established (about 10 days).

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15. Keep the pots in a clean location, at 25 to 27 C with 14-16 hours illumination.

16. Until the plants are well rooted irrigate lightly with tap water if it has a low salt content; otherwise

use demineralized or rain water. Do not overwater.

Photo 3

17. When roots are established, you may dissolve supplementary nutrient in the irrigation water.

Commercial peat moss often contains fertilizer, thus less additional nutrient may be required.

18. Gradually expose the plants to the normal atmosphere by removing the beakers for short periods

each day.

19. Once the plants are established, transfer to larger (e.g. 20 cm diameter) pots. Be careful not to

break the roots. When the plants are well rooted, normal fertilizer can be dissolved in the

irrigation water. At CIP we use 5 g N:P:K at 12-12-12 per liter water. Apply 50-100 cm3 per 20 cm

diameter pot. Again, do not overwater.

Micropropagation

1. Materials and equipment

Culture medium (see next two following sections)

Test tubes

Plastic caps or cotton wool

Forceps

Scalpel

Parafilm

Alcohol lamp

Alcohol 70%

Autoclave

Sterile work area ("microvoid")

2. Prepare the nutritive growth medium according to the procedure given in the section on Medium

to Sweet potato Micropropagation.

3. Dispense 4 cm3 of the medium in each test tube. Cap the tubes with plastic caps or cotton wool

plugs and autoclave them for 15 minutes. Keep the test tubes vertical while the agar sets.

4. Working under sterile conditions (sterile area or "microvoid") follow steps 8 to 10 of previous

section.

5. Transfer the plantlets from the test tuber to a sterile petri dish and make nodal cuttings using

sterile scalpel and forceps. Each nodal cutting consists of a 0.2-0.5 cm stem segment with an

axillary bud (Photo 4).

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Photo 4

6. Place each nodal cutting in a test tube. Ensure that each cutting lies on the agar surface with its

axillary bud pointing upwards (Photo 5). Place 1 or 2 cuttings per test tube.

Photo 5

7. Close the test tubes, seal with parafilm, label and place in a clean area where the room

temperature is 25-27° C. Give 45m E/m2/seg2 or 3,000 lux illumination for 14-16 hours each day.

8. The axillary bud of each nodal cutting grows into a new plantlet within 2-4 weeks and is ready for

transplanting to pots as previously described, or for further micropropagation (Photo 6).

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Documents

Manual for TC Cassava Sweet Potato