Vanilla

Crop improvement of vanilla is in progress in India at the Indian Cardamom Research Institute (ICRI), Spices Board, P.O. Kailasanadu, Idukki, Kerala; Indian Institute of Spices Research (IISR), P.O. Marikunnu, Calicut, Kerala, Kerala Agricultural University, and other institutions. These institutes maintain a good germplasm collection of vanilla as well. A minimum descriptor for vanilla is developed for characterizing the germplasm (Kuruvilla et al., 2000).

Species Diversity

V. planifolia G. Jackson is the cultivated vanilla species in India. In addition to this, the other vanilla native species available in India are Vanilla pilifera Holtt., Vanilla andamanica Rolfe., Vanilla aphylla Blume, Vanilla walkeriae Wight, and Vanilla wightiana Lindl. The Tahitian vanilla (Vanilla tahitensis J.W. Moore) is also conserved in India.

Breeding Behavior

The floral biology of V. planifolia is adapted to outcrossing, and hand pollination is resorted to pod set in India, as the natural pollinators are absent in the country (Sasikumar et al., 1992). No self-incompatibility or natural crossing is reported in V. planifolia from India as observed in Reunion Islands or Mexico (Bory et al., 2008a). However, in V. wightiana, natural fruit setting is reported (Rao et al., 1994). Stray fruit set under natural conditions is also seen in V. aphylla.

Reproductive biology of V. planifolia such as time of pollination, stigma receptivity, and effect of pollen load on the size of the beans were studied (Shadakshari et al., 1996; Bhat and Sudarshan, 1998, 2000). These authors reported that the ideal time for pollination is from 6 a.m. to 1 p.m., and stigma receptivity is up to 24 h. They also observed that complete transfer of pollen results in maximum fruit growth.

Inflorescence initiation in vanilla occurs in late January or early February and the flower opens from mid-February to April–May. Pollination is carried out manually as and when the flower opens and the process is continued for at least 40–60 days. The beans develop quickly in the initial stages and attain their full size within a period of 5–6 weeks under favorable conditions and thereafter slow down. The rate of elongation of beans is maximum during the first 30 days after fertilization (Kuruvilla et al., 1996).

The reproductive mode of V. planifolia needs to be studied, thoroughly looking into the rate of outcrossing, self/cross incompatibility, and autogamy.

Cultivar Diversity

Although vanilla was introduced to India about 200 years ago, the present-day gene-pool in India is derived from this original introduction. Being perpetually propagated vegetatively from these original germplass, a wide genetic base in the primary gene pool of vanilla is very unlikely (Sasikumar, 2004), akin to the situation in some other vanilla-producing countries (Soto Arenas, 1999; Lubinsky, 2003; Bory et al., 2008b). There is no authentic record of any subsequent introductions to the country.

Over the years of domestication and selection by farmers, some new variants (subcultivars) have been recognized in Mexico and Reunion Islands, but no such variants are reported from India, barring a variegated mutant “Marginata” (Minoo et al., 2006a, 2008b) and some accessions with branched inflorescence and varied leaf size. Somatic crossing over (Nair and Ravindran, 1994) reported in V. planifolia can give rise to new variation, apart from mutation, sexual recombination, or epi-genetic variation.

Genetic diversity analysis of V. planifolia accession in India will be worth attempting as it may help to confirm the existence of genetic variability, if any, in the germ-plasm and help in the breeding program.

Interspecific Hybridization

The variation that exists among the cultivated species of vanilla or even in some related species can be combined to produce new types through hybridization. The secondary gene pool may contain useful genes for self-pollination, root rot and virus resistance, larger fruits, reduced photosensitivity, better aroma profile, and pod indehiscence for incorporating into the cultivated vanilla. Because of incompatibility between some vanilla species, breeders may attempt to raise the progenies of interspecific crosses through in vitro seed culture. Interspecific hybrids between V. planifolia × V. aphylla (Minoo et al., 2006b, 2008a) and V. planifolia × V. wightiana (Rao et al., 1992b) are reported from India.

The xenia/metaxenia aspects too can be looked into inter alia as it is observed that pollen of some vanilla species have a positive effect on the pod size of V. planifolia (B. Sasikumar, personal observation).

Climate and Soil

Vanilla being a climbing orchid, conditions favorable for its vegetative growth as well as flowering are to be provided for the commercial production of vanilla beans. The plant requires warm and moist conditions of humid tropics for proper growth and sustainable production. It thrives well between 10°N and 20°S latitudes, having well-distributed moderate rainfall between 150 cm and 300 cm. The crop tolerates a wide range of temperature, but ideally, the mean minimum temperature during winter months should not go below 12°C and should not exceed 35°C during summer, and the optimum temperature range being of 25°–32°C with a mean relative humidity of 80%. The plant does not tolerate any prolonged duration of drought or water-logging, and exposure to hot sun or to strong winds.

According to Potty and Krishnakumar (2003), vanilla can be successfully cultivated in areas nearer to the equator, where warm and humid climate prevails throughout the year and up to an altitude of 1100 m asl.

Vanilla prefers land with gentle slope and light porous soil with adequate drainage. Soil with high humus content is preferred, although the plants can thrive well in sandy loam to even lateritic soils. The humus-rich soils of Western Ghats and the northeastern states of India are highly suited for its cultivation. Vanilla is grown successfully as an intercrop in coconut and arecanut gardens (Sasikumar et al., 1993).

Propagation

Vanilla is amenable to both sexual and asexual methods of propagation. The stem cuttings are capable of striking roots at nodes when they come in contact with soil or any other rooting media. Vegetative propagation through stem cutting is, by and large, the accepted method because it is easy and quick to establish. However, vanilla being a monopodial orchid, collection of sizeable quantities of stem cuttings from the main plantations could lead to the arrest of vegetative growth of mother plants (Ayyappan, 1990) and will be at the expense of subsequent years’ crop. Hence, production of planting materials in nurseries is resorted to under commercial farming. In areas where viral disease is rampant, virus-free certified micropropagated materials are recommended for reviving the plantations.

The primary source of cuttings for the nurseries may be collected from disease-free, healthy, and vigorous mother plants from yielding plantations. The length of the vine used for planting varies from place to place, but has profound influence on further growth and time taken to attain maturity. In situations where there is scarcity of mother vines, cuttings of three to four nodes are used. It is recommended to use such cuttings in the nurseries to raise longer vines rather than directly planting in fields because shorter stem cuttings take longer time for establishment and yielding.

Trench Multiplication

A simple and rapid multiplication procedure for planting material production was described (Kuruvilla et al., 2003). The site for the rapid multiplication nursery should be ideally located with respect to accessibility, availability of water source, gentle slope, and deep and fertile soil of loamy nature having optimum natural shade. Wherever the optimum shade is not available, it is necessary to provide adequate shade using agro-shade nets permitting 50% light intensity. Trenches of 60 cm wide and 60 cm deep are opened at convenient length, leaving 40 cm in between. The trenches are filled with topsoil, well-decomposed farmyard manure (FYM), and sand in the ratio 3:1:1. Standards of 2–2.5 m length for trailing of vines are planted at a distance of 1 m along the trenches at least two months before planting vanilla. Fast-growing trees possessing low branching habit, small leaves, and rough bark are preferred. Plumeria alba, Erythrina lithosperma, and Glyricidia maculata, and others are the common live standards used in India. One-meter vanilla vine cuttings with at least 10 nodes are to be used for planting in the rapid multiplication nursery. Three or four basal leaves of the vines are cut retaining one-third of leaf blades and dipped in 1% Bordeaux mixture for 15 min and then kept in shade for about one week for partially losing the moisture.

The basal portion of the vine is laid on the soil surface near the standards and covered with a thin layer of soil in such a way that the cut end of the vine is bent upward above the soil surface to avoid contact with soil. The rest of the vine is then tied on to the standards so that the nodes are pressed to the standard. The plant base should be mulched with partially decomposed wood debris or leaf litter. The vines are allowed to grow on the supports to a height of 1.5–2.0 m and later coiled loosely around the branches of the supports. The plants are to be irrigated at frequent intervals, as per need. Vanilla responds well to fertilizers. Application of 50 g N, 25 g P₂O₅, and 80 g K₂O per vine in six split doses in a year at bimonthly intervals enhances the growth of vines. Alternatively, foliar application of fertilizer could be given at bimonthly intervals. A well-maintained plant would produce about 5–7 m of growth per year, and therefore the size of the nursery can be planned taking into account the annual requirement of the planting materials. On an average 1 ha of nursery would be sufficient to produce about 40,000 m of planting material annually.

Polybag Nursery

An alternative method to generate planting material when there is shortage of standard vine cuttings of 1 m is to raise the shorter cuttings with two or three nodes in polythene bags and growing it to 8–10 nodded rooted cuttings of standard size. Field establishment of polybag plants is invariably better. Nurseries should be located close to the main field for early to transportation. Black polythene bags of 20 × 20 cm size and 100–150 gauge thickness are normally used for planting stem cuttings. In order to facilitate proper drainage, five to six holes are given at the lower half of each bag. The bags are filled with potting mixture prepared with jungle topsoil, decomposed FYM, and sand in the ratio of 3:1:1. Siddagangaiah et al. (1996) observed that vermicompost and decomposed coir pith are better rooting media for vanilla.

Cuttings with three nodes are planted in polybags and tied to bamboo splits/stakes inserted in the bags for support. The length of the cuttings used has been found to have profound influence on subsequent growth. When two node cuttings were used, vine growth was only 32.6 cm in six months, against 51.8 cm with three node cuttings (Krishnakumar, 1995). The exposed soil in the polybag should be given a layer of mulch, preferably decomposed leaves to protect the soil, to retain soil moisture, and to serve as a source of plant nutrients. Partial shade should be provided either by using agro-shade nets or coir nets.

ICRI, Spices Board of India has standardized a more convenient and economical method of raising polybag nurseries. Vanilla cuttings planted in polybags are allowed to trail in coir yarn as illustrated (Figure 20.1). The fully grown nursery vines along with the coir yarn would be tied to the standard during field planting ensuring least disturbances to the poly bagged seedlings. The moisture holding capacity and rough surface of the coir yarn favored better vine growth (ICRI, 2006).

FIGURE 20.1 Nursery production of vanilla on coir yarns (ICRI method).

The plants are maintained with regular watering. Foliar application of 1% diam-monium phosphate (DAP) can be given to growing vines two months after planting, which can be repeated at monthly intervals. Alternatively, vermiwash can be used for enhancing the growth of vines. Spraying vermiwash once in two weeks increased the length of vine to 66 cm in six months compared to 42 cm obtained in the control (ICRI, 1999). The cuttings so raised in the polybags will be ready for field planting in 6–7 months, by which time they should have reached about 50–70 cm of vine length.

Supports for Vines

Vanilla, being a climbing orchid, requires standards for support and shade. A number of tree species have been recommended as suitable standards for vanilla. Apart from live standards, trellis, latticework, wooden and concrete posts, and wire or bars are used as support for vanilla vines. Wooden posts are subject to decay and to the attack of termites and hence it becomes necessary to replace them at frequent intervals. In case of wire or bar supports, the tender portion of the vines may be easily broken. When nonliving supports are used, it is always necessary to provide some form of partial shade for the vanilla plants. Hence, live standards are preferred for vanilla cultivation.

In general, the support tree species with small leaves, which permit filtered light through the foliage, are useful. Species that can easily be propagated through long stem cuttings and those that grow faster and produce branches sufficiently low (from 1.5 to 2.0 m from the ground) for the vines to hang within easy reach of the workers are found to be the most ideal. The trees should be strong enough to support the heavy growth of vines and beans and should withstand strong wind and rainfall. Trees should never entirely defoliate during summer months, but should be amenable for periodic pruning for shade regulation. It is also important for the support trees to have deep penetrating roots so that they do not compete with the shallow rooted vanilla plants for nutrients. Some of the common suitable support trees used in India are Gliricidia maculate, P. alba, Morus sp. Casuarina, Liberian coffee, Coral tree, Lebeek tree, E. lithosperma, Jatropha sp., and so forth.

Planting

Vines of 1 m length either from plantation/trench nurseries/polybag rooted cuttings are generally preferred for planting. Cuttings are planted close to the base of the support tree by laying the 3–4 basal nodes, from where leaves have been removed, on to the soil surface and gently pressing these nodes to the soil or putting sufficient soil to cover the nodes. While planting, care should be taken to ensure that the basal cut end portion of the cutting is kept just above the soil surface, as otherwise chances of decay are more. The top end of the cutting is to be tied to the base of the support tree gently so that it will eventually climb on to them. Partially decomposed organic materials such as coconut husk, mulch, straw leaves, and so on should be placed over the newly planted cutting at the base of the support tree to a thickness of 10–15 cm or more. If shade is not sufficient from the support tree, palm fronds or other leaves can be used to provide shade to cuttings.

The ideal time for planting vanilla is when the weather is neither too rainy nor too dry. Planting in August or September after the southwest monsoon is recommended under Indian conditions. It takes about 4–8 weeks for the cuttings to strike roots and to show initial signs of growth from any one of the leaf axils. Under irrigated conditions, rooted polybag cuttings could be planted any time of the year.

After Care

A vanilla plantation has to be given constant attention after its establishment. It should be frequently visited to train the vines to grow at a convenient height, prune the growing vines and tree supports, and observe diseased and pest-infected plants. Cultivation of other crops near the roots is not advisable as it may disturb the roots and soil leading to damage of vanilla plants. Periodical removal of dead vines or rotting portion of vines is to be undertaken regularly. Success of the plantation would depend on preventing any serious outbreak of diseases and creating favorable conditions for flowering and beans production.

Nutrient Management

Vanilla, being a hemiterrestrial orchid, providing right soil environment at the plant base through appropriate mulching, rather than soil fertility per se, is important for its successful cultivation. The best source of nutrients is the deep layer of decomposed mulch maintained over and around the vanilla roots. The nutrient-supplying capacity of mulch depends on the source and composition of the mulch.

Nutritional studies carried out at the ICRI, Spices Board has indicated that vanilla yield can be enhanced by soil application of 20:10:30 g NPK per vine per year and foliar application of urea, single super phosphate, and muriate of potash at the rate of 1.0%, 0.5%, and 1.5%, respectively, during January, May, and September. The crop responds well to foliar application, and therefore during the initial years of vanilla growth, foliar application of inorganic manures is practiced. DAP (2%) and muriate of potash (1%) or NPK complex (2%) may be sprayed on the lower surface of the leaves when adequate humidity (>60%) is available in the plantation. Foliar application of DAP and micronutrients, particularly Zn and B boost the development of pods. Studies on the effect of nutrient uptake through aerial roots of vanilla revealed that Knop’s solution (25%) showed improvement in the growth of vanilla (ICRI, 2003).

The crop is highly amenable to organic cultivation. In addition to chemical fertilizers, various organic manures are applied to vanilla. Decomposed organic matter, bone meal, FYM, vermicompost, and fermented deoiled cakes are applied at least twice in a year, that is, during June–July and September–October. Foliar application of vermin wash (1:5 dilutions) favored the growth and development. Application of Vyrsha ayurvedic preparations such as panchakavya, fish amino extract, and egg amino extract is also being widely practiced under organic vanilla cultivation.

Mulching

Mulching with green or dry leaves is to be carried out at least twice in a year. They not only provide food in the form of potassium-rich organic matter but also conserve moisture and hold the layer of mulch in position, creating a deep and rich root zone that vanilla favors. In studies on the growth performance of vanilla with various mulch materials, coir pith favored relatively better vine elongation compared to traditionally used wood debris mulch. The advantage of coir pith is attributed to better moisture conservation, maintenance of conducive microclimate, and supply of micro- and macronutrients for vanilla growth (ICRI, 2005).

Vanilla requires a moist climate with frequent but not excessive rains. Under excessive rainfall, there is widespread occurrence of rot diseases. Under prolonged drought conditions, the plants may suffer from physiological damage and the vines may not recover. In extremely dry conditions, irrigation should be provided at least once in 4–5 days. Mulching with mango leaves (25 kg per vine per year) and watering once in four days was found to increase vanilla yield (Muraleedharan, 1975). Depending on the area under cultivation, sprinkler or hose irrigation may be practiced. Microsprinkler/fog irrigation is advocated to maintain relatively high humidity levels in the plantations to ensure high percentage of bean sets. Care should be taken to avoid water stagnation around the plant base or excessive water stress at any point of vine or pod development.

Shade Management

Careful lopping of branches of the support tree is very important to give shade to vanilla plants. Newport (1910) stated that checkered shade rather than dense shade is to be preferred. Vanilla vines require more sunlight than shade during flowering and at the time when the beans are maturing.

It is the farmers experience that the existing shade trees should be pruned to admit sunlight as uniformly as possible, 30–50%. It may be desirable to regulate shade allowing more light during the blossoming period. Allowing more light is helpful to check vegetative growth and favor flowering. Heavy shade should be avoided because the stem will become thin, leaves small, and flowering and fruiting generally reduced. Under too much sunlight, on the contrary, the leaves not only get sunscald and turn yellow but the plants become weak during a drought period and more susceptible to root rot diseases. The amount of light vanilla can tolerate depends mainly on the water supply to the roots as influenced by the atmospheric humidity and irrigation practices.

Under high rainfall and high relative humidity, vanilla can withstand more sunshine than during low humidity and drought periods. Thus, it is important that support trees maintain much of their foliage during dry periods.

Trailing and Pruning

The trailing nature of vanilla vines has an effect on flowering. The vines are coiled around the lower branches of the supporting tree or over the lattice of trellis so that they may hang down. Care is required not to tear or bruise the leaves, branches, or roots. Bending of vines appears to be an important operation for inducing flowering and fruiting beyond the bend, which may be due to accumulation of carbohydrates and possibly other flower-inducing hormones in such regions of the vine.

Arresting the vegetative growth of vanilla vine by pruning helps to induce flowering. Training and pruning are undertaken to induce flowering and bean production. Here, the hanging shoots of 1–1.2 m long are bent down around the branches of the support tree, slightly twisted in the process, with tips pruned at about 45 cm from the soil. Any vegetative shoots appearing after the bend portion of the hanging shoot are removed, but the shoots appearing on the rest of the plant before the bend portion of the hanging shoots are allowed to grow. These shoots will constitute the bearing branches of the following year. As a result, there is a decreased sap flow toward the bearing branches, which favors flower formation. After the harvesting of beans, the yielded portion of the vine is removed from the plant. The following year’s bearing branches are to be prepared by bending and pruning. Plants that have been pruned and readied for flowering require heavy manuring of leaf mold, decomposed leaves, lime, ashes, and manure. The vine architecture after 3–4 years is therefore of vine with a number of shoots hanging down over the branches of the live support, without overcrowding or overlapping. Normally, five to six bearing branches per plant can be prepared per annum, depending on the age and growth of the vine.

The yielding behavior of pruned hanging shoots under different growth stages comprising of 4, 6, 8, 10, and 12 nodes per shoot was studied. It revealed that the formation of inflorescence is irregular in the hanging shoots irrespective of its growth stage. The 10-node hanging shoots had maximum number of inflorescences per shoot. However, the first-grade quality of beans (above 15 cm) was 57% in 10-node shoots, 77% in 6-node, and 88% in 4-node hanging shoots, respectively, indicating that production of more number of beans in a bunch or in a productive vine would be at the expense of size of the beans (Hrideek et al., 2003).

When the pods are ripe and harvest is completed, the yielded portion of the vine is removed and only the new shoots of the previous year are retained. Plants that have been pruned and readied for flowering require heavy manuring of leaf molds, rotting leaves, lime, ashes, and cow dung.

Fungal and viral diseases are the main production constraints. The crop losses due to these biotic stresses vary. Fungal diseases such as Fusarium and Phytophthora rots cause severe crop losses, and occasionally total crop losses are reported. The viral diseases, although not lethal, result in varying degrees of production and productivity losses. Vanilla being an exotic plant for India, the planting material might have come from outside unknown sources where the virus-free nature of the material was never ascertained and quarantine regulation was never applied. Lack of adequate molecular diagnostic facilities, especially for viral diseases, is another major constrain that limits the detection of viruses. This is important in view of the vegetative propagation method of cultivation. The genetic variability with respect to disease/ pest resistance is comparatively low, which might be the reasons for high disease incidence and consequent crop loss.

Except for the reporting of new diseases, detailed investigations on the management of various diseases are insufficient. Integrated pest and disease management remains an important priority for crop protection of vanilla, with a greater focus on biological control to reduce pesticide inputs for vanilla cultivation.

Vanilla being a crop with poor genetic variability for disease resistance, efforts should be made to induce more variation to identify reasonable disease resistance or tolerance for the major diseases. The recent report of variable disease reaction in seedling progenies of vanilla and somaclonal variations is of considerable interest and needs to be pursued further (Minoo et al., 2006b).

The main diseases reported on vanilla in India are given in Table 20.3.

Fungal Diseases

This is one of the most destructive diseases observed in all the vanilla-growing countries and is severe in India too. It was first reported in India in 1980 (Philip, 1980). Later it was shown to be caused by Fusarium oxysporum sp. vanillae (Thomas et al., 2002).

Symptomatology and Etiology The disease is observed during warm humid conditions. The fungus colonizes the root and causes yellowing of the vines with degeneration of the root system. It also causes dark brown rot on the stem as well as on the leaves (Figure 20.2). The rotting starts at any part of the plant. All parts of the plant are prone to infection, but more often at the nodal region. Later it spreads upward causing rotting, resulting in drying up of the distal portion beyond the point of infection. Severely affected gardens appear dry from distance, typical of wilting. The affected stem exhibits vascular browning, typical of fusarial wilts (Y.R. Sarma, unpubl. data).

FIGURE 20.2 Fusarium stem and leaf infection.

Epidemiology

Although detailed epidemiological investigations have not been carried out, ambient temperature around 25–28°C favored the disease incidence and spread. The disease occurs individually and in certain cases overlaps with Phytophthora rot. Hence, mixed infections are common. Abundant sporulation is observed in the affected tissues. Disease spreads through rain splashes within the plant and across the vines. Detailed investigation on the disease progression curves in relation to environmental condition is called for. The fungus survives on dead dried-up debris and the dead and the dried-up vines left over from gardens are the perennial source of inoculum.

Disease Management

Since the available cultivars are all highly susceptible and as such, no disease resistance exists. Recent report of variability of seedling progenies of V. planifolia to stem rot is important and needs intensive field evaluation (Minoo et al., 2008b).

Cultural Practice

A regular monitoring of the disease is essential for early identification of the disease to take corrective measures. Phytosanitation involving collection and removal of infected vines from the garden is essential to reduce the pathogen inoculum in the soil.

Chemical Control

Phytosanitation followed by spray and soil drenching with 0.2% carbendazim periodically, depending on the intensity of the disease, has been suggested (Thomas et al., 2002).

The disease was first recorded in vanilla plantation of Manalaroo Estate in Nelliampathi and Arnakal Estate at Vandiperiyar during 1998 and also in a stray vine at Directorate of Arecanut and Spices Development Office, Calicut (Y.R. Sarma, unpubl. data). It was later observed in Idukki and Kottayam districts of Kerala, specially in Moovattupuzha and Koothattukulam areas (Bhai and Thomas, 2000).

Symptomatology and Etiology

The disease incidence coincides with the southwest monsoon in Kerala. The disease occurs as water-soaked spots. Observations at Manalaroo and Arnakal Estates showed that Phytophthora infection occurs on leaves, stems, and beans causing rotting (Figures 20.3 and 20.4). All parts of the vine are prone to infection. On leaves, the water-soaked spot enlarges up to 0.5–1.5 cm with a translucent advancing margin. On stems, it is typically a wet rot unlike that of fusarial rot, and affected portions become soft and rot completely. In vanilla fruit bunches, infection starts at any point of beans either from the tip end or from the stalk region. When the infection starts at the tip, it gradually spreads upward resulting in varying degrees of rotting. If infection occurs at the stalk or peduncle region, either the infected bean drops off or dries resulting in beans hanging to the peduncle region (Y.R. Sarma, unpubl. data).

FIGURE 20.3 Phytophthora stem and leaf infection.

FIGURE 20.4 Phytophthora bean rot.

The causal agent was identified as Phytophthora meadii. Under artificial inoculation, infection was recorded 5–8 days after inoculation. The fungus on carrot agar medium appears cottony white. The sporangial size of 39.6 × 20 μm with an LB ratio of 1.98 was recorded. The cultures were heterothallic. Also, intercalary chlamy-dospores in the mycelium were observed (Bhai and Thomas, 2000).

Epidemiology

No detailed epidemiological investigations have been conducted. However, the disease coincides with the southwest monsoon with continuous rainfall of 15–20 days in a month with intermittent showers during the July–August period of the year. On the infected beans, fungal growth occurs, which harbors abundant sporangia. Disease spreads through rain splashes both in the vine and across the vines.

In Kerala and Karnataka, P. meadii has been reported on rubber, small cardamom, and arecanut. It is important to investigate their pathogenic relationship at molecular level and the possibilities of crossing and development of new races/ biotypes of P. meadii.

Disease Management

Cultural practices: As mentioned earlier in the case of fusarial rot/wilts, phytosani-tation measures consisting of systematic collection and pulling out weeds of infected and dried up vines is important.

Chemical control: Spraying of the vines with 1% Bordeaux mixture and soil drenching with copper-oxy chloride (0.25%) was reported effective (Bhai and Thomas, 2000). Prophylactic spraying of the vines with 4–5% of potassium phos-phonate (Phytophos 40), a systemic chemical, at 15 days intervals did control the disease but if the wet spells continued, the disease control was poor (Y.R. Sarma, unpubl. data).

This disease, caused by Sclerotium rolfsii, was reported from the Ramamangalam area of Kerala in a survey carried out during 1999 (Thomas and Bhai, 2000).

Symptomatology and Etiology

Rotting of either one or two beans or all the beans in a bunch was observed. Whitish mycelium spreads on the bean in a fan-like fashion. The infected beans showed rotting symptoms with deep sunken areas that are reddish brown. On leaves and stems, the fungus spreads whitish thick mycelial threads and later produces cream-colored to brownish grain-like sclerotial bodies (Y.R. Sarma, unpubl. data).

This is a minor disease observed in Chempukadavu area of Kozhikode district of Kerala in a mixed cropping system of coconut, arecanut, clove, and vanilla. The disease is caused by Cylindrocladium quinquiseptatum. It starts as water-soaked spots on beans, which later enlarge into a sunken spot of 1–10 mm. The lesions coalesce forming reddish-brown sunken lesions resembling anthracnose. Either a single bean or all beans in a bunch get affected. Leaf infection with sunken lesion was also observed (Bhai et al., 2006).

This is another minor disease of vanilla observed in Kozhikode district of Kerala, caused by Colletotrichum vanillae. The disease is characterized by premature yellowing of the beans with dark-brown sunken lesions that gradually expand resulting in rotting of the beans (Figure 20.5). This leads to premature bean shedding (Bhai et al., 2006). In a survey during April–May, 2003, premature bean shedding was recorded. It was reported that high incidence was due to high temperature and low relative humidity (Bhai and Dhanesh, 2008).

FIGURE 20.5 Colletotrichum bean rot.

The disease is generally observed in vanilla plantations during the postmonsoon period of September–December. It starts as a brown patch on the petiole and lower portion of the unfolding youngest leaf. The funnel-like leaf holds rainwater and might predispose to infection. Later infection spreads downward to the tender portion of the shoot causing shoot tip rot. F. oxysporum isolated from the affected shoots was found pathogenic. Association of Colletotrichum gloesporide was also observed. Spraying the foliage with 0.2% carbendazim was found effective (Thomas et al., 2002, 2003).

Others

Rhizoctonia solani and Mucor racemosus have also been recorded as pathogens of vanilla (Bhai and Dhanesh, 2008), but their relevance need to be investigated further.

The effectiveness of biocontrol agents for the management of soil-borne plant pathogens has been well established in recent times. Its importance was clearly brought in the disease management of crop spices (Sarma, 2006a, 2006b). Reduction of pathogen inoculum in vanilla plantations by applying Trichoderma viride, Trichoderma harzianum, and Pseudomonas fluorescens was reported earlier (Thomas et al., 2002). Both Fusarium and Phytophthora overlap in the field and spatial segregation of these pathogens in a plantation under a set of ecological conditions is not possible. It is imperative and logical to identify biocontrol agents that are suppressive to both these pathogens of vanilla. Studies carried out with different isolates of P. fluore-scens and Bacillus sp. both in in vitro and in vivo conditions showed the bioefficacy of certain bacterial isolates to suppress P. meadii and F. oxysporum in vitro. They also provided growth promotion in vanilla. The suppression of fusarial wilt in large-scale field establishment with local strains of T. harzianum in Mauritius has been established (Y.R. Sarma, unpubl. data).

Viral Diseases

Importance of vanilla viruses at global level received considerable attention (Wisler et al., 1987; Pearson and Pone, 1988; Pearson, 1990; Benezet et al., 2000; Grisoni et al., 2004). Identification of virus problems of vanilla in India is recent. It was only during 2003 that incidence of viral disease was recognized in the states of Kerala and Karnataka (Bhai et al., 2003; Sudharshan et al., 2003). Systematic investigation on vanilla virus diseases started recently at IISR, Calicut, on identification, characterization, and molecular diagnostics (Bhat et al., 2004; Bhadramurthy, 2008).

Electron microscopic studies revealed the presence of three types of flexuous particles resembling Potexvirus, Potyvirus, and Closterovirus and an isometric particle (Bhat et al., 2004). Investigation on virus incidence in vanilla plots carried out recently revealed about 3–10% incidence in Karnataka and 0.13–5% in Kerala (Bhadramurthy, 2008). These studies revealed that cucumber mosaic virus (CMV, Cucumovirus), cymbidium mosaic virus (CymMV, Potexvirus), bean common mosaic virus (BCMV, Potyvirus), and bean yellow mosaic virus (BYMV, Potyvirus) are infecting vanilla in India and mixed infection have been observed. The first authentic investigation on vanilla CMV strains in India was undertaken by Madhubala et al. (2005). The affected vines appear with distorted foliage with small and leathery leaves (Figure 20.6). In artificial inoculation, the members of Chenopodiaceae, Cucurbitaceae, and Fabaciaceae were found infective with CMV strains of vanilla. Isometric virus particles with 28 μm diameter were seen based on the detailed electronic microscopic investigation. Further studies confirmed that vanilla CMV strains belong to subgroup I of CMV. Mild chlorotic mottling and streaks on leaves are the characteristic symptoms of CymMV (Figure 20.7). The investigation revealed that Indian CymMV isolates are closely related to vanilla CymMV isolates from French Polynesia (Bhat et al., 2004). Studies on the yield losses and the epidemiology are lacking and need to be intensified to develop effective disease management strategies.

FIGURE 20.6 Stem necrosis caused by cucumber mosaic virus.

FIGURE 20.7 Leaf mosaic caused by Cymbidium mosaic virus.

Vanilla vines may be symptomless carriers of viruses. The present status of multiplication and new planting would continue as long as yield reduction is not observed. From the practical view point one can go for replanting when the virus infection exhibits declining phase in the yield. It is desirable to build up information on the yield reduction in relation to virus infection to ascertain and identify the exact declining phase with respect to yield. This would guide the management strategy to know when to resort to replanting.

Since these are all sap transmissible viruses, farm operations such as pruning, looping, or pollinating have to be regulated to avoid undue virus transmission leading to disease spread.

Elimination of viruses from the planting material through meristem tip culture, production, and distribution of healthy planting material would be a priority for sustainable vanilla production in future.

In India, pest problems are generally minor on vanilla; however, several insects are recorded to damage vines, shoot tip, flower buds, or roots, by Hemipteran bugs, Lepidopteran caterpillars, and Coleopteran weevils.

Hemipteran Pests

Halyomorpha picus (Pentatomidae) (Figure 20.8) causes serious damage by sucking the sap from the shoot tip and inflorescence. Subsequently, such affected areas become necrotic and rot. Nymphs and adults suck the sap from the peduncle and flower buds. Pin-prick-like punctures at the site of feeding and subsequent necrosis and rotting are the typical symptoms of bug feeding. The affected vegetative buds drop within 3–5 days and the affected inflorescence become rotten. Incidence of the pest is higher during the inflorescence initiation period, that is, January–February.

FIGURE 20.8 Vanilla bug H. picus (a) shoot tip necrosis, (b) nymph, and (c) adult.

The bug is reported to occur in Karnataka, particularly where vanilla is inter-cropped with arecanut plantations. It is also reported from Koothattukulam in Kottayam district of Kerala.

Female bugs lay spherical eggs in clusters on the lower surface of leaf. The eggs are white when laid and turn cream within 3–4 days. Nymphs hatch in 5–6 days. First-instar nymphs are gregarious and remain on top of the egg shells and do not feed. The second- to fifth-instar nymphs are black and actively feed on the shoot tip.

The nymphal duration lasts for about 60 days. The pest causes about 40% damage by way of feeding on the inflorescence (Prakash and Sudharshan, 2002).

Management of H. picus includes monitoring of the bug during November– February and removal of egg mass and first-instar nymphs that are seen on the lower surface of leaves. Spray of Monocrotophos at 0.1% a.i. controls the nymphs, if the infestation is high.

Nezara viridula (Pentatomidae) occurs throughout the tropics; the bug lays eggs on leaves and stalks; the nymphs suck the sap of flower buds and stalks. The pest is reported to occur on vanilla in Karnataka and Kerala but its incidence is very low (Varadarasan et al., 2002a).

The bug was reported to occur on vanilla in Kerala and Karnataka as a minor pest. The nymph and adult suck the sap from beans and leaves (Varadarasan et al., 2002b).

This insect found in Idukki district, Kerala, sucks the sap from the leaves, vines, and inflorescence. Ants are found to be associated with this sucking insect and the pest incidence occurs during January–February (Varadarasan et al., 2002b). Control measures with Monocrotophos at 0.1% a.i. is warranted only if the pest is serious.

Coleopoteran Pests

Three species of Coleopteran pests are recorded in vanilla. Among them a weevil, Sipalus sp. occurring in Idukki district of Kerala, causes serious damage to young shoots, vines, and leaves.

The weevil lays eggs singly along the length of the vine and the emerging grubs feed on the inner core of the stem by boring a tunnel, the entire length of vine shreds, rots, and falls down (Varadarasan et al., 2002a).

The adult weevil is 8–10 mm long, 2–3 mm wide, and is light to dark black with two wavy white cross bands on the elytra. They feign death upon approach or touch (Thanatosis). The adult female is larger than the male. Adults mainly feed on shoots by inserting the snout; the injured area becomes necrotic within a day, leading to rotting of the shoot tip/vine. The weevil also feeds on leaves by scrapping upper or lower epidermis with mesophyll tissue, leaving a thin transparent epidermis on the lower or upper surface of leaves (Figure 20.9).

FIGURE 20.9 Vanilla vine weevil, Sipalus sp. (Left: egg deposited with necrosis; right: grub tunneling the vine).

After copulation, females lay eggs singly, 2–4 mm below the epidermis on the vine or shoot tip. The egg is capsule like 5–6 mm long, 0.8–1.0 mm wide, white and subsequently turns to yellow before hatching. The site of egg deposition develops necrosis. Eggs are laid only on tender vines and not on the leaves. The emerging grubs feed on the necrotic tissue by making tunnels in the vine.

The first-instar grub is yellow and the final-instar grub is white with brown head capsule. The grub (larva) period lasts for 35–40 days. The grubs tunnel the vine by feeding and the entire length of the affected vine becomes necrotic. The mature grub pupates inside the tunnel with fibrous material; pupation lasts for 19–21 days. The adult weevil emerges from the vine by making a small slit in the dried vine.

Extensive damage by the grubs and the adult weevil on the vine is observed mostly in open areas where shade is less. The adult weevils are seen in the field during November–January, and can easily be located. The weevils are not very active, and, hence, they may be hand picked and destroyed to reduce the damage on the crop (Varadarasan et al., 2002a).

This beetle, found in Wynad and Nilgiris in India, cuts through the leaf from the lower surface, eating the entire leaf tissue, except for the thick translucent cuticle of the upper epidermis. The damaged leaves rot as a result of fungi infection. The pest may be managed by keeping the garden weed-free and in severe cases by application of Malathion 0.1% (Rai and Nayar, 1976).

Holotrichia (Scarabaeidae): Roots of vanilla are damaged by white grubs in Idukki and Thiruvananthapuram districts of Kerala. Drenching Chlorpyriphos 0.05% a.i at the plant base during May or application of entomopathogenic nematodes controls the pest.

Lepidopteran Pests

The caterpillar of the moth occurring in Idukki, Kottayam, Ernakulam, and Thrissur, districts of Kerala, feeds on the vegetative shoots; these pale green caterpillars are seen in between the shoot bud and the first leaf, forming a web and feeding the shoot that leads to rotting of the terminal bud (Figure 20.10). Incidence of the caterpillar was recorded in January and February. The adult moth is reddish brown with a broad bright yellow band across forewings, gray hind wings, and body. The pest may be controlled by spraying Monocrotophos at 0.1% a.i. if the incidence is high (ICRI, 1996).

FIGURE 20.10 Shoot tip damage by webber caterpillar, Archips sp.

The yellowish ash-colored caterpillars with brown heads are very agile and damage the inflorescence and flower buds by feeding (Figure 20.11). The late instar caterpillar is light brown and pupates in between the damaged inflorescence. The pest was recorded in Chemannar in Idukki district of Kerala during February–March. Monocrotophos 0.075% or Lambda cyhalothrin 0.004% control the pest efficiently.

FIGURE 20.11 Left: Egg deposit on flower bud, right: damage on a flower bud by semi-looper, Nemoria sp.

Noninsect Pests

Among these, mites, snails, and avian pests cause considerable damage.

This mite is found to infest beans in storage in India (Sasikumar et al., 1992).

Snail: Achatina sp: They were found to feed on the chlorophyll tissues from the base of the vine and were recorded in Nagercoil in Tamilnadu, where coconut fronds were used for mulching on the plant base. The snails hide in the mulch during daytime, and come out in the night to feed on vanilla. The snails are also found to feed on leaves (Figure 20.12). The management of this pest includes avoiding mulching with coconut fronds or any other planting materials that do not decompose fast. The snails may be collected from the mulch and destroyed (Varadarasan et al., 2003b).

FIGURE 20.12 Damage on leaves by snail, Achatina sp.

Chicken cause much damage by scratching among the mulch and thus exposing and damaging the roots.

Vanilla Scab

Longitudinal corky formations on the beans of vanilla have been observed, which are commonly referred as “vanilla scabs.” It was suspected to be caused by insects or fungi or due to abiotic factors such as dew drops or higher light intensity. Experiments conducted at the ICRI, Kerala, indicated that scab formation in vanilla is not due to insect or fungus or due to any other abiotic factors. It was found that the mechanical injury at the early stage of the beans caused scab. Close observation on beans, leaves, and internodes showed that wherever the early stage of plant parts come in contact with hard objects, such as bamboo or dry twig or even mature leaf lamina, scab formation was observed on the point of contact. Subsequently, experiments with nail injury on the ovary during pollination show scabs on the beans. Any minute injury on the early stages of the ovary is stretched longitudinally with the elongation of beans, and thus scabs are formed (Varadarasan et al., 2005).

Fresh vanilla beans are dark green in color and do not impart any aroma because vanillin and other chemical substance are not available in free form at the time of harvest. The beans are ready for harvest when they attain a mild yellow color at the distal end, and it may take 8–10 months to reach the harvesting stage depending on the conditions. Full ripening of the beans leads to splitting and ultimately affects their quality. The beans are to be harvested at the right time, as the immature ones produce inferior commodity. The nonsynchronized flowering and fruiting behavior of the crop leads to selective harvest extending to 1–2 months. Some of the important factors that determine the vanillin content and beans quality are climatic conditions, stage of harvest, and extent of sweating of the pods during curing. During the process of curing, free vanillin is developed in the beans as a result of a series of enzymatic reactions that provide fragrance.

Many processing methods are followed in vanilla-growing countries. However, the Bourbon method of vanilla processing practiced in Madagascar is followed in India, and modified and standardized to suit local conditions. This method is simple and consists of four stages (killing, sweating, slow drying, and conditioning, see Chapter 11) (Krishnakumar et al., 2007, 2008).

Packing

In the final stage of curing, beans are packed in wax paper or any appropriate material and stored in closed boxes for a period of three months or longer to permit the full development of the desired aroma and flavor. Evaluation of different packing methods and methods for keeping up the quality of beans during storage indicated that the beans stored in polyethylene bottle, glass tubes, polypropylene covers (0.011 and 0.035 mm), acrylic boxes, and waxed paper 0.07 mm plus tin were on par with respect to vanillin content and retention of moisture (ICRI, 2006).

According to Purseglove et al. (1981), the primary quality requirement for cured vanilla beans is the aroma/flavor character. The other traits signifying the quality are general appearance, flexibility, size of beans, and vanillin content. Superior quality beans are long, fleshy, supple in nature, very dark brown to black in color, somewhat oily in appearance, strongly aromatic, and free from scars and blemishes.

The quality standards used in India are as per ISO specifications and there are no separate standards specified by Bureau of Indian Standards (BIS). The local traders/ processors have set their own norms based on size of the beans: supergrade beans have a bean size of more than 20 cm, clean without any blemishes; A-grade beans (16–20 cm); B-grade beans (12–16 cm); C-grade (8–12 cm); and low-grade beans comprised of shorter beans, beans with splits, cuts, and wrinkles, and beans with scars/ blemishes. In general, from a healthy plantation, 20–40% of beans produced would be of A grade including supergrade; 30–50% B grade, and the remaining C grade or rejects. In order to attain larger proportion of A-grade beans, farmers restrict the pollination to 10 flowers per bunch and 10–12 bunches per vine. The approximate composition of whole vanilla beans is moisture: 25–30%; protein: 2.56–4.87%; fatty oil: 4.69–6.74%; volatile oil: 0.0–0.64%; nitrogen-free extract: 30.35–32.90%; carbohydrates: 7.1–9.1%; fiber: 15.27–19.6%; ash: 4.5–4.7%; vanillin: 1.48–2.90%; resins: 1.5–2.6%; calcium: 19.7 mg%; potassium: 16.2 mg%; sodium: 6.7 mg%; phosphorus: 9.5 mg%; and iron: 0.3 mg%.

Analysis of Indian vanilla beans carried out inside and outside the country have shown that the vanillin content was invariably above 2.5% with an aroma and flavor comparable to Madagascar vanilla.