Abstract
Pollination is one of the most critical events in sexual reproduction of flowering plants. Pollination is the basis of gene flow and genetic recombination. Studies on pollination ecology were prevalent even before Darwin; they became intense after Darwin formulated the concept of co-evolution between flowers and pollinators. The number of publications on pollination perhaps exceeds those in any other area of reproductive ecology. Pollination is simply the transfer of pollen grains from an anther to the stigma. Pollination ecology is the study of pollen transfer through understanding of interactions between plants and pollinators in relation to the prevailing habitat. Except in some apomictic species which do not depend on fusion of the second male gamete with polar nuclei for endosperm development (pseudogamy), effective pollination is a prerequisite for successful seed development. A comprehensive understanding of pollination ecology of a species needs a thorough familiarity with the phenology, floral morphology and sexuality of the species.
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Pollination is one of the most critical events in sexual reproduction of flowering plants. Pollination is the basis of gene flow and genetic recombination. Studies on pollination ecology were prevalent even before Darwin; they became intense after Darwin formulated the concept of co-evolution between flowers and pollinators. The number of publications on pollination perhaps exceeds those in any other area of reproductive ecology. Pollination is simply the transfer of pollen grains from an anther to the stigma. Pollination ecology is the study of pollen transfer through understanding of interactions between plants and pollinators in relation to the prevailing habitat. Except in some apomictic species which do not depend on fusion of the second male gamete with polar nuclei for endosperm development (pseudogamy), effective pollination is a prerequisite for successful seed development. A comprehensive understanding of pollination ecology of a species needs a thorough familiarity with the phenology, floral morphology and sexuality of the species.
The most important feature of pollination that drives all relevant adaptations is that plants are sedentary and they have to make use of external agencies to achieve pollination. Wind, water and animals are the three pollinating agents. Nearly 90 % of the flowering plants are pollinated by animals (biotic pollination/zoophily), and the remaining 10 % of the species use abiotic agents – largely wind (anemophily ) – and a small proportion of them use water (hydrophily) (Ollerton et al. 2011).
Depending on the origin of pollen, pollination is categorized into the following three categories:
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Autogamy – transfer of pollen grains from the anther to the stigma of the same flower
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Geitonogamy – transfer of pollen from the anther to the stigma of another flower of the same plant or of another plant of the same clone (ramet)
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Xenogamy – transfer of pollen from the anther to the stigma of a different plant (not of clonal origin, genet)
Autogamy does not require pollinating agents, while geitonogamy and xenogamy require pollinating agents for pollen transfer. Another term, allogamy, has often been used in the literature to indicate transfer of pollen from one flower to the other irrespective of whether it is from the same or a different plant.
In several species, autogamous self-pollination (without external agents) takes place to different degrees. Cleistogamy is one such mechanism in which flowers never open and the stamens and the pistil remain enclosed within the flower bud (Lord 1981; Richards 1986; Kaul and Koul 2009). In cleistogamous flowers, the stigma and anthers are in contact with each other; pollen grains germinate inside the anther, or after coming in contact with the stigma, pollen tubes enter the stigma and grow through the pistil. Most of the cleistogamous species such as Commelina benghalensis and species of Viola produce both cleistogamous and chasmogamous (flowers that open) flowers. Often the production of cleistogamous flowers depends on the prevailing environmental conditions particularly the temperature and light. Ruellia, for example, produces chasmogamous flowers during the summer and cleistogamous flowers during the winter under Delhi conditions. Cleistogamous flowers are exclusively autogamous.
Even in several chasmogamous species, autogamous pollination has evolved as a means of reproductive assurance in the absence of pollinators (Kalisz and Vogler 2003; Eckert et al. 2006). Dioecious species and those which are strictly self-incompatible are exclusively xenogamous. Most other species show mixed mating system; both cross- and self-pollinations occur to various degrees depending on the structural features of the flower and the visitation frequency and efficacy of the pollinators.
Some of the major floral adaptations of the three pollination syndromes are summarized below:
7.1 Anemophily
Amongst the abiotic mode of pollinations, anemophily (wind pollination) is more prevalent than hydrophily. Anemophily is common in several families such as Poaceae, Juncaceae and Cyperaceae. Anemophilous species are generally characterized by non-showy flowers. The perianth is much reduced or even absent, and the flowers lack colour, nectar and smell. Wind-pollinated species produce a large amount of pollen grains to compensate for the uncertainties of pollen landing on the stigma. Pollen grains are dry and powdery with a smooth surface. The stamens are large and borne on long filaments and their anthers are well-exposed to the air. The stigma is also large, well-exposed and often feathery to trap airborne pollen. The number of ovules in the ovary is generally reduced, often to just one. Wind pollination is limited in tropical forests (about 5 %), where the movement of wind below the canopy is highly reduced and frequent rains wash the pollen from the air, when compared to temperate forests.
Wind pollination is considered as secondarily derived (Endress 1994; Bronstein et al. 2006; Waser and Ollerton 2006). This is based on the prevalence of predominantly insect pollination in fossil records of primitive angiosperms, lack of wind pollination in extant basal angiosperms and occurrence of wind pollination in specialized families of angiosperms.
7.2 Hydrophily
Hydrophily (water pollination) is rare and is limited to just about 30 genera of 11 families, largely monocotyledons (McConchie 1983). Water acts as a vector in the transportation of pollen in hydrophilous species. A majority of plants growing in water are not hydrophilous; their flowers emerge above the water level and are pollinated by other agents as in terrestrial plants (largely by biotic agents). Marine angiosperms are exclusively hydrophilous.
Basically, there are two types of hydrophily – epihydrophily or ephydrophily and hypohydrophily or hyphydrophily . This distinction is largely based on the movement of pollen; in the former it is two dimensional, and in the latter it is three dimensional. In epihydrophilous species, pollination takes place on the surface of water. In several of them, the reproductive organs are carried just above the water surface (dry epihydrophily), and pollen grains do not come in contact with water during pollination. In others, pollen floats on the surface of water (wet epihydrophily). In hyphydrophilous species, the flowers are submerged in water, and thus pollen grains are dispersed below the water surface and come in contact with submerged stigma. Hypohydrophily is reported in 18 genera of which 17 are monocots and 12 are marine species (Cox 1988, 1993).
Water-pollinated species tend to show unisexual flowers with reduced perianth and absence of colour, nectar and smell. The number of ovules is generally reduced, usually to just one, and the stigma is rigid with a large surface. Adaptations of pollen grains to water medium are found in only those species which come in direct contact with water (wet ephydrophily and hyphydrophily ). In such species, the exine of the pollen grains is highly reduced or even absent. Pollen grains are covered with a coating of mucilage to prevent them from wetting in water. Pollen grains of many such species are filamentous which facilitates their movement in water.
Studies on pollination biology of hydrophilous plants are more difficult when compared to those pollinated by other agents, and the information available is limited (Ducker and Knox 1976; Cox 1988, 1993; Cox et al. 1990). There are not many methods that have been described to study hydrophily. Epihydrophilous species may be studied by direct observation. Exclusion methods used for terrestrial plants need to be modified to suit water medium. Floating rafts or small boats are used to approach flowers in water bodies. For epihydrophilous species, metal or plastic cages suitably modified to prevent the movement of pollen grains on the surface of water can be used in exclusion experiments. Pollination studies on hyphydrophilous plants are more difficult and require diving gears. Growing plants in aquariums is more convenient to study floral biology and pollination biology in hydrophilous species when compared to natural conditions.
7.3 Zoophily
An essential feature of zoophily/biotic pollination is that plants have to develop effective devices to attract suitable animals to visit their flowers in a sustainable way by providing them with some rewards and use them effectively for pollination services . Further, plant species have to use some degree of discretion and restrict the number of animal species visiting the flowers of each species to a reasonable number. If they attract animals indiscriminately, all potential pollinators, present in the habitat, may visit all synchronously flowering plant species; this would bring about extensive heterospecific pollination which seriously reduces the fitness of the plant species. The aim of pollination ecologists is to understand how plants perform these conflicting demands of attraction and restriction of animal species for pollination services. Studies on pollination ecology so far have largely been focused on understanding the details of attraction. Only limited studies have been carried out on the details of restriction of potential pollinators (Shivanna 2014).
As pointed out earlier, the number of species pollinated by animals far exceeds those pollinated by abiotic agents. Zoophilous flowers exhibit an amazing variety in the size, shape, arrangement, colour, scent and sexual system. The evolution of such matchless variety of flowers obviously facilitates the flowers in performing the dual function of attraction and restriction of pollinators. Figure 7.1 presents some of the variations found in flowers and their pollinators. The flowering plants, although last to evolve amongst plant groups (over 100 million years ago in the early and middle cretaceous period), became the most successful group and occupied a pre-eminent position amongst all groups of plants. Evolutionary success of angiosperms has been attributed to the origin of the flower and associated evolution of biotic pollination to bring about cross-pollination (see Pellmyr 2002; Shivanna 2003; Willmer 2011). Amongst animals, insects are the major pollinators. According to one estimate, of the 13,500 genera of angiosperms, 500 contain bird-pollinated species, 250 contain bat-pollinated species, and 874 contain wind- or water-pollinated species; the remainder contain mostly insect-pollinated species (Renner and Ricklefs 1995). Hymenoptera (bees, wasps and ants), Lepidoptera (butterflies and moths), Coleoptera (beetles) and Diptera (flies) are the major orders of insects involved in pollination. Birds and bats are the other important pollinators.
Bees show great variation in body size and length of proboscis. Bees (both medium and large sized) are the most important pollinators. Bees are good in recognizing colours and scents and efficient foragers of nectar and pollen. Unlike humans, many bees can perceive light in ultraviolet range but cannot visualize shades of red which appear black to them (Kevan 2005). Butterflies are active during the day and land on the flower before foraging. Beetles are a very diverse group of insects. Beetles tend to move around the flowers chewing floral parts indiscriminately and get covered with pollen.
Thrips (Thysanoptera) are frequently present in flowers and feed on pollen and nectar. In several species, thrips have been shown to be the major pollinators (Mathur and Mohan Ram 1978; Ananthakrishnan 1993; Garcia-Fayos and Goldarazena 2008). Their limited flights largely promote self-pollination. However, movement of thrips, which are very small insects, is often assisted by wind, and in such situations, their pollen may be carried for longer distances (Ghazoul and Sheil 2010).
Depending on the position of anthers in the flower in relation to the entry of the insects, pollen grains may be deposited on the body of the insect in a diffuse manner, or they may be deposited on the upper (nototrobic) or lower (sternotrobic) surface of the body. The position of the stigma is generally such that it comes in contact with the surface of pollen deposition and the pollination is brought about. Diffuse distribution is generally associated with primitive flowers and unspecialized insects, while the other two types are commonly found in advanced flowers and specialized insects (Faegri and van der Pijl 1979).
Bird-pollinated (ornithophilous) species have been reported in about 65 families (see de Wall et al. 2012). Amongst birds, humming birds (family Trochilidae) are one of the major pollinators and are restricted to the New World; they forage while in flight by hovering near the flower. Sunbirds (Nectariniidae) and sugarbirds (Promeropidae) are common pollinators in Asia and Africa. Sunbirds are capable of hovering, but perch if a perch is available. The other major bird pollinators are honeyeaters (Meliphagidae), restricted to Australia, and honey creepers (Drepanididae) endemic to Hawaii (Kearns and Inouye 1993). In India, as many as 58 bird species belonging to 16 families have been reported to be involved in pollination of 93 species of flowering plants (Subramanya and Radhamani 1993). The information available on bird pollinators is much less than those on insect pollinators. Bird-pollinated flowers are generally bright coloured, most of them being red and scentless. The nectar is the main reward for bird pollinators; bird-pollinated flowers produce copious amount of nectar with low viscosity. The nectar is located in long corolla tubes.
Bats are nocturnal pollinating agents. Bats involved in pollination belong to the order Megachiroptera in the Old World and Microchiroptera in the New World. Bat pollination (Chiropterophily ) has been recorded in a large number of tropical and semitropical plant species of over 40 families (Endress 1994; Gibson 2001). Bats can hover, cling or perch while foraging the flowers. Moths (Lepidoptera) are the other major group of animals involved in nocturnal pollination .
Apart from insects, birds and bats, there are other groups of animals particularly nonflying mammals such as marsupials, rodents and primates (Kress 1993; Endress 1994; Ghazoul and Sheil 2010) which are involved in pollination in a limited number of species. Some of these unusual pollinators are cockroaches (Nagamitsu and Inoue 1997 – Uveria), mice (Wester et al. 2009 – Pagoda lily), squirrels (Tandon et al. 2003 – Butea), snails (Sarma et al. 2007 – Volvulopsis) and lizards (Olesen and Valido 2003; Ortega-Olivencia et al. 2012 – Scrophularia, Hansen et al. 2007 – Trochetia).
Pollination brought about by each group of pollinators has been given a specific terminology (Table 7.1). Variation in floral attractants, rewards and flower morphology determine, to a large extent, the type of animal species that visit the flowers and the specificity of the visiting species. The literature on pollination is full of a large number of terminologies. Some of these terminologies with reference to the flowers and pollinators are presented in Table 7.2.
7.3.1 Floral Attractants and Rewards
Plant species pollinated by animals have to advertise their presence and provide rewards for the visiting animals to sustain their visits. Floral colours, sizes and shapes act as visual attractants, and olfactory attractants are in the form of floral scents. In most of the species, advertisements are provided by floral organs, although in a few, extra floral organs such as bracts may take part in attraction. Most of the species with hidden nectar have contrasting patterns on the corolla termed nectar guides (Fig. 7.2a, b) that guide the visitor to the source of nectar; their size and shape are highly variable. Many investigators have experimentally shown that these nectar guides do play a role in guiding pollinators to the site of nectar (Leonard and Papaj 2011; Hansen et al. 2012).
Several plant species have evolved traits that are beneficial to both the plant species and the pollinator. Postpollination colour change is one such feature that has been recorded in over 214 species of 74 geographically and taxonomically diverse families (Gori 1983; Weiss 1991). In these species the pollinated flowers, instead of senescing, change colour and are retained on the plant. In Lantana camara, for example, the flowers are yellow on the day of anthesis and offer pollen and nectar to the pollinators. They turn orange and then red on subsequent days and do not offer rewards (Mathur and Mohan Ram 1978). Red flowers are retained on the plant for several days depending on the variety and the weather conditions. Experimental studies have shown that the retention of older flowers on the inflorescences increases plant’s attractiveness to pollinators from a distance (Weiss 1991). They discriminate the colour of the flower from a close range and visit yellow flowers significantly more often than orange or red flowers. Thus, the retention of pollinated flowers not only increases the visibility of the flowers to pollinators but also guides them to rewarding flowers.
Olfactory cues are in the form of volatile fragrance compounds emitted by flowers. The chemical composition of floral scents is one of the most extensively investigated areas of floral biology sine long because of their commercial value in perfume industry. The role of floral volatiles in pollinators’ attraction is comparatively recent. The fragrance is in the form of complex mixtures of a large number of volatile compounds. Floral fragrance is largely made up of monoterpenoids, sesquiterpenoids, phenylpropanoids and benzenoid compounds (Williams 1983; Knudsen et al. 2006).
The complex mixture of volatiles is characteristic for each species. No two species, even if they are closely related, have been shown to produce identical mixture of volatiles. The specificity of fragrance of a species is established not by individual fragrance compound but a combination of compounds. Insects are able to distinguish complex mixtures of floral volatiles from different species and respond accordingly (Dudareva and Pichersky 2000 ; Riffell 2011). Maximum emission of fragrance generally coincides with the activities of their pollinators, and the emission often follows endogenous rhythm (Dudareva and Pichersky 2000). Floral scent often changes after pollination and thus enables the pollinators to avoid pollinated flowers (Schiestl et al. 1997).
Pollen grains also emit odours that differ from those of other floral parts and are characteristic of the species (Dobson 1988; Dobson and Bergstrom 1996, 2000). Insects are able to discriminate pollen odours of different species. Flowers of many species emit unpleasant odours, due to the presence of amine-containing compounds which serve as chemical attractants to some insects particularly beetles and flies.
Plant–pollinator interactions are largely mutualistic; they result in reciprocal benefits to both the partners. It is a form of “biological barter” and involves exchange of resources of the plant such as pollen and nectar with the services of the pollinator (Ollerton 2006). Pollen and nectar form the most important rewards for the pollinators. A few species pollinators collect oils, and some others collect nest materials such as resins and waxes (Armbruster 2012). Nectar is largely made up of sugars secreted by the nectary. The major sugars present in the nectar are sucrose, glucose and fructose. Apart from sugars, nectar also contains small amounts of amino acids and traces of lipids, phenolics, alkaloids and proteins (Baker 1977; Nicolson and Thornburg 2007; Heil 2011). The amount of nectar produced per flower (common range 0.1–500 μl) and the concentration of sugars (common range 5–45 %) present in the nectar are highly variable. Nectar of several species has been reported to contain bacteria (Freidman et al. 2012) and a few contain yeasts (Herrera et al. 2009). The role of bacteria and yeasts in nectar is not well understood. In some species, the nectar is coloured (Hansen et al. 2007) and in several, it is scented (Raguso 2004).
Pollen grains are highly nutritious; apart from carbohydrates (ca 25 %), proteins (ca 25 %), amino acids (ca 10%) and lipids (ca 5 %), they are rich in vitamins and minerals (Schmidt and Buchmann 1992; Roulston and Cane 2000). Pollen grains are needed for the larvae and young ones of insects. Bees gather pollen in special parts on their body, the pollen baskets/corbiculum. Pollen baskets in honeybees and bumblebees are on the hind legs, but on leaf-cutting bees, they are under the abdomen.
Many Neotropical orchids produce fragrant compounds largely terpenoids and aromatics to attract as well as reward male euglossine bees. Euglossine bees collect fragrant compounds and store them in their modified hind legs (Dressler 1982). The bees are thought to use these compounds to produce sex pheromones that are released to attract females (Tan 2006).
In some species, the flowers reward the larvae of the pollinators with young seeds (nursery pollination). The pollination of Ficus species by fig wasps and of Yucca by yucca moth (Tegeticula) represents highly specialized obligate nursery pollination systems; each plant species is pollinated by a specific wasp/moth species (Machado et al. 2005). Neither the plant nor the pollinator is able to reproduce without the other. Most of the Ficus species are monoecious and produce male and female flowers in specialized inflorescences termed syconia. The female wasps enter the receptive syconium through the terminal ostiole. They lay their eggs in a proportion of female flowers. The larvae of the wasps feed on the gall formed in oviposited flowers, and the remaining pollinated flowers develop into seeds. The emergence of adult wasps from the larvae coincides with the maturation of male flowers. The wingless male wasps are short lived; they mate with the females, cut an exit tunnel in the wall of the syconium and die. The females loaded with pollen come out through the exit tunnel and enter another receptive inflorescence (which is in the female phase) through the ostiole to reproduce; they bring about pollination during their movement inside the syconium. Several investigators have shown that volatile compounds emitted by the receptive syconia are responsible for the attraction of their specific pollinators (Khadari et al. 1995; Proffit et al. 2008, 2009). The syconia emit volatile compounds only during the period of receptivity of female flowers.
7.3.2 Nocturnal Pollination
In a great majority of species, pollination is diurnal and takes place during the daytime. However, in some species, pollination occurs during night-time. In yet others, pollination occurs during the day as well as night (Young 2002). In species with nocturnal pollination , flowers remain open during the night. In several species with diurnal and nocturnal pollination, the life span of flowers lasts for several days. This strategy of nocturnal and diurnal pollination ensures seed set when pollinators are scarce or unpredictable (Dar et al. 2006 and references therein). Beetles, moths, particularly hawkmoths, bats, and a few species of bees and rodents are the common nocturnal pollinators (Muchhala et al. 2009). Hawkmoths have the longest tongue amongst insects. They are confined to tropical areas.
Although visual cues are the predominant attractants for diurnal pollinators, they are not reliable during night-time; acoustics and olfaction are the principal means to locate flowers by nocturnal pollinators. Flowers of nocturnal pollination species are not brightly coloured but emit strong odour. Bat- and rodent-pollinated flowers produce large quantity of nectar. Bats of Microchiroptera (New World) can produce ultrasonic sound to locate flowers. The sound is reflected by the petals of bat-pollinated flowers, and bats have the ability of recognizing this reflected sound (echolocation) (von Helversen and von Helversen 1999). Echolocation ability is not developed in most of the Megachiroptera (Old World); they depend largely on olfactory and visual cues to locate flowers. When compared to diurnal pollination, the information available on nocturnal pollination is limited largely due to the difficulty of observing flowers and pollinators at night. Logistic problems in conducting observations particularly in tropical forests also deter studies on nocturnal pollination .
7.3.3 Pollination by Ants
Studies on plant–pollinator interactions are largely centred on flying insects as they are the most important pollinators around the world. Ants are active 24 h a day either as individual species or as overlapping guilds of species foraging for particular periods during the day or night. Although ants are amongst the most abundant insects on earth and visit flowers frequently, ant pollination has not evolved as a major pollination syndrome. Ant pollination has been documented in less than 20 species (Beattie et al. 1984; Beattie 1985). Several hypotheses have been put forward to explain the reasons for the absence of frequent evolution of ant pollination syndrome (see Beattie 1985). There are some evidences to indicate that the secretion of antimicrobial substance on their body surface (as a means of protection against bacteria and fungi in the soil) is harmful to the pollen also. So far, reports of ant pollination have been confined to herbs and small shrubs.
7.3.4 Non-mutualistic Pollination
In a number of species, flower–pollinator interactions are non-mutualistic. The flowers do not provide rewards for their pollinators (rewardless flowers), or the floral visitors exploit floral rewards without affecting pollination (nectar and/or pollen robbers) (Wiens 1978; Dafni 1984; Dettner and Liepert 1994; Renner 2006; Bronstein et al. 2006). Rewardless flowers exploit pollinators by signalling the presence of reward without providing the reward (deception). Non-mutualistic plant–pollinator interactions have evolved in all major groups of flowering plants. Orchids form the major group of non-mutualistic interactions; about one third of orchids (ca 3,000 species) are reported to be deceptive (Renner 2006).
Food deception is one of the widely recorded non-mutualistic pollinations. Non-rewarding species (mimic) coexists with rewarding species (model), and the flowers of the mimic resemble the flowers of the model. Floral visitors draw rewards from the model but do not discriminate strongly against non-rewarding flowers. Food deception is largely mediated by visual signals; olfactory signals do not seem to play a major role. The fragrance compounds in several food deceptive orchids have been shown to be quite different and week when compared to its model (Galizia et al. 2005; Salzmann et al. 2007).
A number of orchid species, particularly of Ophrys, achieve pollination through sexual deception – the shape and colour of their flower mimic the female insect of its pollinator, and the floral scent mimics the pheromones of the female insect. The pheromones attract mate-seeking males to the flowers from a distance. The visual cue serves as a close-range attractant and induce sexually stimulated males to land on the flower; the male tries to copulate (pseudocopulation) with the flower and brings about pollination (Ayasse et al. 2003; Schiestl et al. 2003; Huber et al. 2005; Phillips et al. 2014). Although sexual deception is predominant in orchids, it has been reported in a few other species (Asteraceae, Ellis and Johnson 2010; Iridaceae, Vereecken et al. 2012).
Several species belonging to ten families such as Annonaceae, Araceae and Aristolochiaceae mimic brood sites and attract insects whose larvae feed on dung/carcasses. Their flowers (see Wiens 1978; Bernklau 2012) mimic the odours of dung and/or carrion to attract coprophilous beetles and flies that oviposit or feed on carrion or dung. These odours smell like decaying proteins or faeces and are very unpleasant to humans. The odours are composed of sulphide compounds, ammonia, alkyl amines, cadaverine and putrescine. Faecal-like odours are also produced by skatole and indole compounds (Dettner and Liepert 1994).
In some species showing brood site mimicry, the odours are enhanced by the production of heat in the flower/inflorescence (thermogenesis). It has been reported in members of Nym-phaeaceae, Annonaceae, Araceae, Arecaceae, Aristolochiaceae and Magnoliaceae (see Thien et al. 2009). Flowers of some of these species have developed trapping mechanisms to retain insects in the flower for one to several days. In species of Aristolochia (Murugan et al. 2006; Trujillo and Sersic 2006), the flowers are protogynous and emit fragrance during the female phase. The pollinators are attracted to the fragrance and enter the flower through narrow corolla tube. The insects cannot escape because of the presence of downwardly pointed hairs on the inner surface of the corolla tube. When the flower enters the male phase, the insects get coated with the pollen; by this time the downwardly pointed hairs senesce and allow the insects to escape. They are attracted to another flower that is in the female phase; they enter the flower and deposit the pollen on the receptive stigma.
7.3.5 Ambophily
Generally, anemophilous and entomophilous species do not show combination of both the syndromes; however, there are several species in which both the types of pollinations are combined (ambophily). Some examples of ambophily are Plantago lanceolata (Stelleman 1978), Urginia maritime (Dafni and Dukas 1986), species of Salix (Sacchi and Price 1988; Tamura and Kubo 2000; Karrenberg et al. 2002) and Thymelaea velutina (de la Bandera and Traveset 2006). In Ceratonia siliqua, for example, the flowers are showy and entomophilous; they show a combination of diurnal and nocturnal entomophilous pollination as well as wind pollination (Dafni et al. 2012). Wind pollination in species showing entomophilous syndrome compensates low frequency of insect visits. Similarly insect pollination in species showing anemophilous syndrome reduces uncertainty associated with wind pollination.
7.3.6 Floral Visitors and Pollinators
Flowers of most of the species are visited by a number of animal species. All of them need not be pollinators; some of them may rob pollen and/or nectar without affecting pollination. It is, therefore, necessary to distinguish pollinators from non-pollinating floral visitors. Many investigators have identified pollinators on the basis of their visit to the flowers and/or presence of pollen load on their body. However, careful studies have shown that all the floral visitors and those with pollen load may not be pollinators (Sinu and Shivanna 2007; Sinu et al. 2011). Thus, presence of pollen load is not an authentic method for identifying pollinators. To confirm a floral visitor as a pollinator, one has to demonstrate the transfer of pollen grains to the stigma and/or seed set after a visit of a potential pollinator to a virgin flower. When there is more than one pollinator of a given plant species, their frequency of visits and pollination efficacy of each species in terms of pollen removal and/or pollen deposition may vary greatly. The pollination efficiency of some of them may be so low that they may not have any perceptible role in overall pollination success of the plant species.
7.3.7 Restriction to Pollinators
Any natural habitat is made up of a number of plant species and diverse animal species, many of which are potential pollinators. However, each plant species attracts only a proportion of potential pollinators to visit its flowers but prevents the visit of several others present in the habitat. As pointed out earlier, for an efficient pollination system, restriction of visits to a reasonably limited number of pollinator species is important to ensure their visits to largely conspecific flowers. Restriction to potential pollinators to the flower may act at different levels – morphology of the flowers, species-specific fragrance and quantitative and/or qualitative features of the nectar and pollen rewards.
7.3.7.1 Morphological Filters
Flowers of many species are generalized type; their rewards (both pollen and nectar) are open and accessible to any visitor (Fig. 7.2c). Flowers of a large number of species show structural diversification, and their rewards are not accessible to all the visitors (Fig. 7.2d). Such flowers restrict the visits to a limited number of animals. One such floral diversification is the change from radial symmetry to bilateral symmetry. Bilateral symmetry enables the flowers to guide the approaching pattern of the visitors to harvest the rewards efficiently. Flowers of such species are visited only by those animal species that are able to locate and harvest the rewards. Another type of elaboration is the evolution of a long corolla tube or a spur in which nectar is located (Richards 1986; Pellmyr 2002). A number of species in Orchidaceae have spurs of various lengths often reaching up to 40 cm. The nectar in such flowers with spurs or corolla tubes is accessible only to structurally suited animals; only those that have the proboscis/beak of suitable length can harvest the reward. Many studies have shown that pollinator species with different tongue lengths tend to specialize on plant species with matching spur/corolla tube lengths (Pleasants 1983).
Pollination Syndromes: The role of morphological traits of flowers that permit some species of pollinators (and prevent some others) was known since long. Traditional concept, elaborated by Darwin, on pollination systems has been that the evolutionary tendency of a species is towards greater refinement making the pollinator and the flower mutually interdependent. This led to the view that the combination of floral traits reflects pollinator type and each pollinator type is the result of selection pressure exerted by different pollinators. This eventually resulted in the formulation of various “pollination syndromes” (Faegri and van der Pijl 1979; Turner 2001). A pollination syndrome is a combination of floral traits associated with the attraction and utilization of a specific functional group of animals for pollination (Table 7.1). Pollination syndrome concept can explain floral diversity and convergence of floral forms across angiosperms pollinated by similar pollinators (Bronstein et al. 2006). A high level of specialization has the advantage for both the pollinator and the plant species as long as both the partners are adequately available. This also has inherent weakness; if one of the partners becomes rare or absent, the other is also bound to fail unless it evolves compensatory mechanisms such as self-compatibility, autogamy, vegetative propagation or apomixis. High specialization also binds both the partners for spreading to new areas.
There has been a considerable discussion in literature in recent years on specialization and generalization in plant–pollinator interaction (Waser et al. 1996; Waser and Ollerton 2006; Johnson and Steiner 2000). Many recent studies have shown that plant and pollinator assemblages are mostly generalized; a majority of plant species are visited by taxonomically diverse groups of pollinators, and most pollinator species visit several plant species. These studies have questioned the concept that floral traits associated with a pollination syndrome constitute an adaptive response (see Mitchell et al. 2009). An advanced level of specialization is seen in a very limited number of species such as some orchids, Yucca with yucca moth, and Ficus with Ficus wasp, in which pollination of each species is brought about by one specialized insect.
Several investigators have emphasized the utility of pollination syndromes in understanding the mechanisms of floral diversification (Fenster et al. 2004) and have elaborated the evolutionary aspects of mutual interaction between pollinators and the plant species (Mitchell et al. 2009). They argue that costs and benefits of plant–pollinator interactions play an important role in determining whether these interactions are more ‘generalized’ or ‘specialized’. If a plant species has many visitors which provide similar and comparable pollination services , there is little incentive for plants to specialize to attract a particular group of pollinators. On the other hand, if some floral visitors are more effective in the quantity or quality of pollen transfer, selection should favour traits promoting these effective pollinators (Mitchell et al. 2009). Variation amongst floral visitors in pollination efficiency is therefore a requirement for the evolution of specialization (Schemske and Horwitz 1984). These pollination syndromes (permitting some and preventing other visitors) are likely to function in combination with floral fragrance.
7.3.7.2 Fragrance Filters
As pointed out earlier, the composition of fragrance is unique to each plant species and attracts a specific pollinator or a group of pollinators. The fragrance acts as a filter as its attraction is restricted to some species; those animals which are not attracted to the fragrance do not visit the flowers (Williams and Dodson 1972; Omura et al. 2000). In species with obligate specialization, fragrance may be the only filter. A meta-analysis of 18 studies on the response of animals to floral scents by Junker and Bluthgen (2010) has highlighted the dual function of floral scents; obligate floral visitors are attracted to floral scent, while those which are facultative and antagonists are repelled by floral scents.
7.3.7.3 Nectar Filters
Studies on nectar have so far highlighted largely the role of its nutritive components, sugars and amino acids, as rewards for the visitors. The amount of nectar present in the flower and its sugar concentration are, to some extent, correlated with the type of animals visiting the flowers (Baker and Baker 1983). Bee-visited flowers generally have lower amount of nectar with higher sugar concentration, while bat- and bird-visited flowers have higher nectar volume with lower sugar concentration. These features are well recognized and form a component of pollination syndromes.
Several studies have highlighted the role of non-nutritive metabolites such as alkaloids and phenolics in attracting or repelling floral visitors (Stephenson 1981; Adler 2000; Irwin et al. 2004; Adler and Irwin 2005; Raguso 2004). Nectar traits often deter nectar robbers without affecting the visits by pollinators. The floral nectar of Catalpa speciosa, for example, contains iridoid glycosides, catapol and catalposide, which adversely affected potential nectar thieves (ants and a skipper butterfly, Ceratomia catalpae); the legitimate diurnal bee pollinators were not affected by these glycosides (Irwin et al. 2004). Similarly, some South African species of Aloe produce dark brown nectar with a bitter taste because of the presence of phenolic compounds. Bulbuls and white eye, which are effective pollinators, are unaffected by the bitter taste of nectar, while bees and sunbirds which are not the pollinators are deterred by the phenolics in the nectar (Johnson et al. 2006).
Floral nectar of several species is scented. Various components of the fragrance have been shown to have positive or negative effects on different animals. In Nicotiana attenuata (Kessler and Baldwin 2006), benzyl acetone attracted the pollinators (moths and hummingbirds); methyl salicylate repelled ants but attracted moths. Thus, the nectar, at least in several species investigated, has the potential to filter flower visitors, favouring some and deterring others.
7.3.7.4 Pollen Filters
Nutritional quality of pollen is highly variable; some of them lack several essential nutrients and some are poor in proteins (Roulston and Cane 2000; Rasmont et al. 2005), and yet others contain secondary compounds which are repellent or toxic to insects (Pimentel de Carvalho and Message 2004; see Hargreaves et al. 2009; Sedivy et al. 2011). Several studies indicate that pollen can act as a filter to select floral visitors. Analyses of pollen loads of several bee species have shown that some species are specialists at the level of plant families or subfamily or even genera, while others are generalists visiting the flowers of up to 15 plant families (Schmidt 1982; Müller and Kuhlmann 2008; Sedivy et al. 2008). Thus, floral visitors show preferences to pollen of some species and avoid visiting the flowers of other species. Such choices in pollen foragers may be physiologically constrained.
A few studies have been conducted on the effects of host and non-host pollen on the development of larvae of bee species (Sedivy et al. 2011). Pollen of Sinapis arvensis (Brassicaceae) and Echium vulgare (Boraginaceae) failed to support larval development of Colletes bee species specialized on pollen of Campanula (Praz et al. 2008). Similarly, pollen of Asteraceae and Ranunculaceae permitted larval growth of only those bee species that are specialized to harvest pollen from plants belonging to these families; their pollen failed to support larval growth of other bee species. These studies clearly indicate that palatability of pollen can act as an effective filter to restrict the number of floral visitors. Pollen of non-host species may hamper the digestion of the larvae, and the bees seem to have adapted their metabolism to digest pollen of host species (Leonhardt and Bluthgen 2012).
7.4 Pollination Efficiency
Pollination efficiency has been defined in various ways by pollination ecologists. Often different and confusing terminologies have been used (see Dafni et al. 2005). Pollination efficiency can be referred to the efficiency of individual floral visitor to bring about pollination. In this manual pollination efficiency of the visitor is defined as the number of pollen grains deposited on the stigma of a virgin flower after one visit by the pollinator. It can also be assessed on the basis of the number of seeds induced by the pollinator in one visit. Pollination efficiency can also be referred to overall pollination efficiency of all the floral visitors combined. Pollination efficiency under field conditions refers to the per cent of flowers that gets pollinated irrespective of the number of visiting species or number of visits of each species.
7.5 Pollination Limitation
Pollination limitation refers to the reduction in seed production by inadequate deposition of conspecific compatible pollen on the stigma. Pollination limitation has been reported in a large number of species; some of the studies indicate that over 60 % of the species may show pollen limitation under certain conditions (Burd 1994; Wilcock and Neiland 2002; Knight et al. 2005). This is one of the major constraints that often drive the populations to vulnerability.
7.6 Pollen Travel and Gene Flow
The movement of alleles physically through space is referred to as gene flow. Pollen grains and seeds are the agents of gene flow. Often the distance for which pollen travels from its source before it lands on the stigma is used as a measure of gene flow. There are different methods such as staining pollen (before their dispersal) with vital or fluorescent stains and labelling of pollen with radioactive carbon, which can be used to measure the distance of pollen travel. The use of genetic or molecular markers to identify the progeny sired by the pollen of a marked plant is the most authentic method to study gene flow (for details see Kearns and Inouye 1993; Dafni et al. 2005).
7.7 Protocols
7.7.1 Estimation of Nectar Volume and Concentration of Total Sugars in Nectar
Nectar is an important reward for biotic pollinators. It is largely made up of sugars . The amount of nectar and its sugar concentration are highly variable and to some extent reflect the nature of the pollinator. The dynamics of nectar secretion, its location in the flower, the quantity and concentration of sugar are important in understanding pollination ecology.
7.7.1.1 Special Requirements
-
Isolation bags and tags
-
Calibrated capillary tubes or micropipettes (1–50 μl depending on the amount of nectar present in the flower of the focal species)
-
Portable refractometer (with 0–50 % range is sufficient)
7.7.1.2 Procedure
-
1.
Bag flower buds before anthesis (preferably the previous evening of anthesis) to prevent visitors from foraging the nectar before estimation.
-
2.
Soon after anthesis, remove the bags and excise flowers for collecting nectar.
-
3.
Collect nectar by gently inserting calibrated capillary tube of suitable capacity (1, 2, 5, 10, 25 and 50 μl capacity are available), depending on the amount of nectar present, and allow sufficient time for the movement of nectar into the capillary tube through capillary action. When the amount of nectar is abundant as in some bird-/bat-pollinated flowers, micropipettes/microsyringes of various capacities (0.1 ml/1.0 ml) may be used.
-
4.
Estimate the amount of nectar for each flower. As the lumen of the calibrated capillary tubes is uniform, the amount of nectar can be estimated by measuring the length of the tube filled with nectar and calculating its amount on the basis of the total length of the capillary tube up to the calibration point. Generally, 10–20 flowers for each reading are satisfactory.
-
5.
Calculate the average amount of nectar per flower and present with SD or SE.
-
6.
Dispense nectar from capillary tube/micropipette onto the surface of the calibrated portable refractometer. Lower the lid of the refractometer slowly without allowing air bubbles to be trapped. The amount of nectar should be sufficient to cover the entire surface of the refractometer when its lid is lowered. The readings of the refractometer indicate nectar concentration as percentage of sucrose equivalents. Repeat this estimation at least for 6–10 flowers and calculate the average.
7.7.1.3 Modifications
-
1.
Instead of bagging the flower buds the previous evening (step 1), flowers can be collected soon after anthesis before they are visited by any floral visitors.
-
2.
For species in which the longevity of flowers extends for several days and also those that show protandry and protogyny, it would be necessary to study the details of nectar secretion every day until flower senescence, particularly during male and female phases. In such species, more elaborate planning for nectar estimation is needed depending on the objective of the study. Larger number of flower buds is bagged, and specific number of flowers is excised at a given time (with reference to the time of anthesis) for nectar estimation. There should be sufficient number of flowers for each set. The results on the amount of nectar are generally presented in the form of a table/graph/histogram over the period with SD/SE for each value.
-
3.
To check if the nectar is secreted every day, carefully remove the nectar from flowers in the evening and bag them. Next morning open the bags and estimate the amount of nectar secreted and concentration of sugars in the nectar. This would give the amount of nectar secreted during the night.
-
4.
In some studies, information on the amount of nectar consumed by the visitor and resorption of unused nectar by the flower before senescence may also be required. Comparison of the amount of nectar in bagged flowers and those in which the visitor has foraged nectar would give the amount of nectar foraged by the visitor. Comparison of nectar amount in fresh flowers and those bagged until initiation of senescence would indicate if the nectar is resorbed in non-visited flowers. When there is active resorption, there is hardly any nectar left in senescing flowers. Marginal reduction in the amount of nectar may be due to the evaporation of the nectar; in such cases, the amount of sugars generally shows a marginal increase.
-
5.
In several species, the amount of nectar present in the flowers is very small (<1 μl). In such species it is difficult to collect nectar through capillaries and estimate sugar concentration using a refractometer. McKenna and Thomson (1988) suggested a sensitive method to estimate total carbohydrates in the nectar in such species. Instead of capillaries, small filter paper strips are used as wicks to collect available nectar. The paper strips are dried and stored until used for sugar estimation using calorimetric method (for details of protocol, refer to McKenna and Thomson 1988).
7.7.2 Floral Visitors and Frequency of Their Visits
Studies on floral visitors, their visitation frequency, foraging time and behaviour are important aspects of pollination ecology. These studies are needed not only to document floral visitors but also to determine their role in pollination of the species. The frequency of visits (number of visits per flower per unit time) would indicate their possible involvement in pollination and also, to some extent, serve as an indicator of pollinators’ abundance. Foraging time (the visitor spends on the flower) indicates the extent of reward available in the flower. The way the forager harvests the reward from the flower (whether legitimate or illegitimate) and patterns of their movements between flowers of the same and other plants would indicate their efficiency in pollination and also the type of pollen they are likely to deposit on the stigma.
7.7.2.1 Special Requirements
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Suitable tags to label flowers/inflorescences. In herbaceous species, pegs and thread may be required to encircle selected patch for observation.
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Tabulated field notebook – although there is no standard format for preparing the table to record the details, it should contain, on the top, the name of the plant species, date of the study and the number of focal flowers marked for observation. A sample table is given for guidance (Table 7.3). The investigator may modify the same or prepare the table in his/her own format convenient for recording the details.
7.7.2.2 Preliminary Studies
This exercise requires some preliminary studies. It would be desirable to spend a couple of time slots (15/30 min in an hour) to become familiar with all the visitors to the flower and to understand the details of recording various events in the tabulated sheet:
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Selection of the focal flowers: Depending on the number of visiting species, their frequency and the size and density of flowers, the investigator has to decide on the number of flowers/inflorescences to be kept under observation on each day so that all the visitors can be easily recorded during the observation time.
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Observation period of the day: Depending on the active period of the visitors (in some species, the visitors may be active the whole day, and in others the activity may be confined to the morning and/or evening hours), the period of record has to be selected.
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Suitable time slots: When the studies are carried out by one person, it is not feasible to make observations on the visitors continuously throughout the day. Generally, 15/30 min slot for each h is selected depending on the frequency of visits. In some species depending on the visitation details, it may be necessary to observe continuously during the day. In such situations, it is better to have two investigators recording alternately on the same set of flowers.
-
Identity of floral visitors: Identity of each animal species that visits the focal flowers needs to be recorded. If you do not know the name of the species, record each unknown species as a morphospecies for later identification. A few individuals of each of the unknown species have to be collected by using either a sweeping net or an aspirator (see Protocol7.7.7), depending on the size of the visitor, and store them in rectified spirit for identification. Give each morphospecies a letter (A, B, C) so that it can be used for regular recording of its visits.
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Mode of their landing/entry: The approach may be legitimate (when it comes in contact with the anthers and the stigma) or illegitimate when it robs the pollen or nectar without touching the anthers and/or the stigma. In actinomorphic flowers, the visitors generally land on the top of the flower and move amidst anthers and stigma. In zygomorphic flowers the visitors may land on one part of the flower and then move for foraging the pollen and/or nectar.
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Foraging pattern : The visitor may forage only pollen or only nectar or both. Some insects forage exclusively pollen/nectar during some period of the day. These features have to be recorded.
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Foraging time: The time spent by the visitor on each flower indicates the reward available in the flower.
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Contact with the anthers and the stigma: It is important to observe whether the visitor comes in contact with the stigma and the anthers during their foraging. This is essential for effective pollination.
-
Behaviour of the visitor after its exit from the flower: When the visitor exits from a flower, it is desirable to note whether it visits other flowers of the same plant and if so the number of flowers of the same plant visited in each bout before moving to the flowers of a neighbouring plant. Sometimes it may move to the flowers of another species. These details would indicate the type of pollen deposited on the stigma.
Since the above features generally do not change for each visitor, some of the above details can be recorded during the preliminary studies and need not be repeated during all the time slots.
7.7.2.3 Procedure
-
1.
Select a small compact flowering patch containing 10–30 fresh flowers from a single plant or a group of plants which can be observed from the observation spot and mark the patch or the flowers with small tags or by any other means. If the number of species visiting the flowers and their frequency are high, it would be better to select a smaller number of flowers so that the details of all the visitors can be conveniently recorded. If the frequency is low, a larger area with larger number of flowers can be selected. The main consideration for selecting the plot is to ensure that the investigator can clearly see any visitor visiting any of the flowers in the selected patch. Record the number of flowers in the selected area. In annuals, the study plot is generally made up of a group of plants which can be kept under observation from the observation spot without difficulty. If necessary, the selected patch may be enclosed with a square/rectangle/circle made with the help of pegs and thread. If it is a large shrub or a tree, the study unit has to be confined to some branches or inflorescences.
-
2.
Select a comfortable place to sit (for insect visitors, 0.5–2 m away from the selected flowers is satisfactory as they are not very sensitive for human presence), and make sure that all the flowers in the selected plot are clearly visible. Birds are particularly sensitive to human presence. They may even avoid visiting the flowers when the humans are present nearby. For observing birds, it is better to select observation spot several metres away from the focal flowers/inflorescences/branches behind other bushes or branches to camouflage the sitting area.
-
3.
Record the visits of all the visitors to selected flowers in the tabulated sheet (see Table 7.3) during each time slot. In compact inflorescences where visits to individual flowers are difficult to record, the inflorescence may be taken as a unit; the frequency is expressed as the number of visits/inflorescence/h. When the movements of foragers are very swift, it would be more convenient to record observations by using a video camera/handycam for later analysis. One of the advantages of video recordings is that the temporal record of the foraging time of pollinators/robbers can be ascertained with ease, and the recordings can be observed several times when more than one pollinator is visiting the focal flowers.
Apart from recording the frequency of each visitor to the focal flowers, observe and record the following in the pre-tabulated sheet:
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Details of foraging: The visitor may forage only the nectar or only the pollen or both; they can be recorded by the letters P/N/B, respectively. Many of the visitors may forage pollen as well as nectar in the morning hours, but confine only to nectar in the afternoon and evening hours when pollen is no more available. Also record whether the visitor’s body comes in contact with the anthers and/or the stigma during their entry/foraging/exit.
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Duration of visit: Depending on the reward available, the duration of visits on each flower may vary from less than a second to several seconds, often minutes. The duration may change depending on the depletion of nectar/pollen in the afternoon and evening. Duration of visits can be recorded with the help of a digital stopwatch of the mobile phone or digital wristwatch. This method, although most accurate, is cumbersome to record the time spent by each visitor on each flower. Alternatively, start the stopwatch when the visitor lands on the first flower in the observation patch, continue the count as it enters other flowers in the observation patch, and stop the watch when the visitor leaves the last flower of the observation patch. Calculate the duration of visits by dividing the total time spent on the selected patch by the number of flowers visited during each bout. Another method some investigators use to measure the duration of visits is to practise and standardize the counting in seconds so that foraging time starting from landing until departure can be counted and recorded. Combine the results of all the foraging trips for each time slot, and calculate the average time spent on each flower.
It is often difficult to record the number of flowers visited by each visitor and the time spent on each flower. We have found it convenient to record only the number of flowers visited by each visitor (frequency) in the observation patch during each bout in a 15/30 min time slot of each h; the remaining time of the h can be spent in recording other details such as the time the visitor spends on each flower, its foraging behaviour and whether the body of the insect comes in contact with the stigma and anther and so on.
As the pollinators’ activity is affected by environmental conditions particularly temperature, light and precipitation, these parameters have to be recorded on each day of the study period.
Observations are discontinued for the day after the visitors cease to visit the flowers.
7.7.2.4 Modifications
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1.
In tall tree species, it would be better to construct a platform so that the investigator can sit comfortably near the selected flowers for longer periods and record the details. In the absence of a suitable platform, the investigator may be able to find a convenient branch to sit for observation. Alternatively, for small trees, a ground patch may be selected, and a good pair of binoculars may be used to observe visitors. However, by this method many of the smaller insects may be missed particularly at higher canopy. While recording observations with the binocular, it would be convenient to have two investigators, one to vocally announce and the other to record the details. Alternately one can use the recorder.
-
2.
In many species, the flowers remain fresh only for a day; in several others, however, they remain fresh for more than a day, often for several days. In such species the observations have to be extended for the same set of flowers every day until the flowers do not receive any more visitors or until the flowers start senescing. Any new flowers that open in each study patch may be removed to confine observations to older flowers.
The above studies have to be replicated at least for 3–5 days for each population of the focal species using different randomly selected patches each day. It may also be necessary to record details of the visitors and visitation frequency during early, mid- and late periods of flowering as the visitors’ guild/their frequency may change over the flowering period.
After completion of the studies on each patch of flowers, analyse the data. Calculate visitors’ frequency (average number of visits per flower per h or any unit time), duration of foraging (average foraging time per flower in seconds) and foraging pattern (nectar/pollen/both) at different time slots studied during the day. The data on replicates on different days of the same time slot may be combined to determine the mean with ±SD/SE of the means. Represent the frequency of the visits and duration of foraging in the form of suitable graphs/histograms/box plots. Depending on the frequency of visits over time, many investigators combine the data of 2–4 h time slots to plot the graph/histogram (e.g. 06.00–09.00, 09.00–12.00, 12.00–15.00, etc.) instead of plotting for each time slot. Determine the total observation time spent for each species (number of hours spread over the number of days).
7.7.3 Identification of Pollinators Based on Pollen Transfer to the Stigma
As pointed out earlier, all the floral visitors may not be pollinators. Many visitors rob the nectar and/or pollen without affecting pollination. Some animals may visit flowers to feed on smaller insects present on the flowers. Identifying pollinators from non-pollinators is very important in pollination ecology.
7.7.3.1 Special Requirements
Tags and isolation bags
7.7.3.2 Procedure
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1.
Select a set of focal flowers as described in Protocol 7.7.2 and sit at a convenient place to record floral visitors. Based on your earlier/preliminary studies, make sure to start observations before the visits start so that none of the flowers in the selection plot has received a visitor until the start of the observation. If this is not possible, the focal flowers have to be bagged one day before anthesis, and the bags are opened just before the start of the observation. This ensures that you start with a set of virgin flowers.
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2.
Keep track of the visits with reference to each visitor. It is better to focus on one visitor at a time. Chase away other visitor that approach the focal flowers by a stick or a small branch; do not allow them to land on any of the focal flowers.
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3.
Soon after the selected visitor exits a focal flower, excise the flower, label it with the name of the visitor, and keep it in a container with as little disturbance as possible for later studies. Continue observation until you excise enough number of flowers for each visitor (say 10–15 on each observation period).
-
4.
Repeat these studies until you cover all the visitor species. If this cannot be done in one sitting, the studies can be spread to several sittings to cover all the visitors with enough number of replicates.
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5.
Take all the excised flowers to the lab without disturbing their stigmas. In bisexual flowers, it is important to make sure that self-pollen grains do not get deposited on the stigma during handling of the flower.
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6.
In the laboratory, take the stigma/pistil of each flower visited by each visitor individually and observe under a stereomicroscope the presence of pollen grains. If the pollen grains are too small to be seen under a stereomicroscope, mount the stigma in a drop of safranin/acetocarmine and observe under a compound microscope to record the pollen load . If the stigma is thick, tease the stigma, before observation. The presence of conspecific pollen on a substantial proportion of stigmas confirms that the visitor is an effective pollinator. Calculate the percentage of flowers that get pollinated by one visit of the specific visitor. If the conspecific pollen is absent on the stigma of any of the flowers visited by a particular visitor, it is not a pollinator. If such visitors (without pollen load on the stigma) forage pollen and/or nectar during their visit, they can be termed as pollen and/or nectar robbers.
7.7.3.3 Modifications
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1.
Pollination efficiency based on pollen load on the stigma: Pollination efficiency of different pollinators can be compared by calculating the percentage of stigmas pollinated by a single visit of each pollinator to the virgin flower. A further modification is that the number of pollen grains can be counted on each stigma under the microscope. If the number of pollen grains deposited is less than the number of ovules in the ovary, such stigmas are generally considered as insufficiently pollinated. If the pollen grains are more than the number of ovules, they are considered as sufficiently pollinated.
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2.
Pollination efficiency of multiple visits by pollinators: More often, pollinators keep visiting the same flower repeatedly, and in some instances, the same flower may receive as many as 8–10 visits by the same pollinator or different pollinators. In many such species, a single visit to a virgin flower may not be sufficient to deposit sufficient number of pollen grains. The above procedure may be modified to collect flowers visited by once, twice, three times or during the whole day of a specific pollinator.
The following is the brief procedure when the flowers are visited by a single pollinator:
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(a)
Give sequential numbers to all the focal flowers kept under observation in the study patch. Each flower may be labelled by an inconspicuous tag (so that it will not distract the visitor), or a rough diagram can be made on the notebook to indicate the position of all flowers under observation and each flower be given a serial number. As the forager visits each flower, note the serial number of the flower visited in the notebook and continue this record for the entire observation period. At the end of the observation period, count the number of visits for each flower. Excise all the flowers at the end of the observation period and categorize the flowers on the basis of the number of visits it has received.
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(b)
Take the flowers to the laboratory without disturbing their stigmas. Observe all flowers of each category (visited 1 to n number of times) under the microscope and calculate per cent of flowers sufficiently and deficiently pollinated for each category.
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(c)
Represent the results in the form of a table/histogram for each visitor. This is reasonably simple when the flowers are visited by a single pollinator.
In species with multiple pollinators, the same flower is likely to be visited by more than one visitor. Such species require more time and efforts to get data on multiple visits of each pollinator to focal flowers. Under each observation period, only the selected pollinator has to be allowed to visit the flower; others need to be chased away by a stick/small branch. This has to be repeated for each pollinator.
-
(a)
-
3.
Efficiency of pollinator based on fruit set: Pollinators can also be confirmed and their efficiency determined based on the fruit set after one visit or multiple visits by a potential pollinator to a virgin flower.
Follow the same procedure as described in the Protocol 7.7.3. Allow each pollinator species to make one or several visits as explained above. At the end of observation period, label each flower for the specific pollinator visited and the number of visits made. Bag the flowers and allow them to develop into fruits. Determine per cent fruit set and average number of seeds per fruit for each pollinator (for one visit and/or several visits) and compare the same for different pollinators to establish their efficiency.
7.7.4 Pollination Efficiency Under Field Conditions
In the above protocol, pollination efficiency is measured only in flowers visited by a specific pollinator. This does not indicate the pollination efficiency under field conditions. Depending on the density of pollinators, all fresh flowers may or may not be visited by pollinators. Also there may be variation in the number of times the pollinators have visited each flower. Unless there are chances for natural autogamy, the flowers which do not receive visitors will not get pollinated and hence do not set fruits and seeds. This protocol gives the methodology to determine pollination efficiency under field conditions.
7.7.4.1 Special Requirements
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None for the main protocol
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Tags for modification
7.7.4.2 Procedure
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1.
Excise the senescing flowers (which cease to attract pollinators) randomly from different plants in the population. The number of flowers to be selected depends on the number of individuals in the population and the area of the study site. The plants need to be selected randomly to cover the entire population. At least ten plants in a population of 50 individuals are recommended. The total number of flowers in such a population should be around 50–100.
-
2.
Study the stigma of excised flowers under the microscope for the presence of pollen and score them as described under Protocol 7.7.3.
-
3.
Determine the percentage of flowers pollinated. In multiovulate species, the percentage of deficiently pollinated and sufficiently pollinated flowers can also be determined by counting the number of pollen grains on the stigma of each flower.
As pollination efficiency is affected by a number of factors, these studies have to be carried out during the early, peak and late flowering season of the population.
7.7.4.3 Modification
Based on Fruit Set: Tag sufficient number of flowers, selected randomly, in the population, and keep them under observation until they abscise or develop into fruits. Count the number of fruits and the number of seeds in each fruit. Calculate per cent flowers that set fruits and mean number of seeds per fruit. Represent pollination efficiency as per cent fruit set and/or mean number of seeds per fruit.
7.7.5 Estimation of Pollination Efficiency Using Spear’s Pollination Index
Many investigators have used Spear’s pollination index (Spears 1983) to evaluate pollination efficiency of each pollinator. In this method, a pollinator is allowed to make one visit to a virgin flower and is bagged (see Protocol 7.7.3). Pollination efficiency (PE) is assessed on the basis of the number of seeds formed per visit. The PE is calculated as follows:
where Pi is the average number of seeds in the fruit that received only one visit by the pollinator i, Z is the average number of seeds in the fruit that received no visits by the pollinator i and U is the average number of seeds in the fruits that received unrestricted visits. The values of the pollination index range from 0 (when there is no contribution by a given pollinator) to 1 (when the production of seeds or fruits by a given pollinator is equal to that of flowers which received unrestricted visits by pollinators).
7.7.5.1 Special Requirements
Isolation bags and tags
7.7.5.2 Procedure
The procedure is basically the same as described under protocol 7.7.3, but requires the following three sets of virgin flowers:
-
1.
Prepare the following three sets of flowers, each with sufficient number:
-
(i)
Flowers bagged the previous evening of anthesis (no visits).
-
(ii)
Allow one visit of each pollinator to each flower and bag them as in Protocol 7.7.3. This set has to be repeated for each pollinator.
-
(iii)
Tag flowers and allow them for unrestricted visits.
-
(i)
-
2.
Monitor all the three sets of flowers to develop into fruits or until their abscission, and record the response of each flower in each set.
-
3.
Harvest the fruits for each set separately. Open each fruit and collect the seeds and count them.
-
4.
Calculate average number of seeds/fruit for each set of flowers.
-
5.
Calculate pollination efficiency using the above formula. The flowers of set (i) give information on seed set as a result of autogamy (Z). Flowers of set (ii) give information on seed set after one visit by specific pollinator (Pi), and flowers of set (III) provide data on seed set after unrestricted visits by the pollinators (U).
Treatment (i) need not be repeated for each pollinator. Seed set data for Z and U obtained for one pollinator can be used to calculate pollination efficiency of all other pollinators.
7.7.5.3 Modification
This formula can also be used to assess pollination efficiency on the basis of the number of pollen grains deposited on the stigma. Instead of bagging and allowing the three sets of flowers to develop into fruits, excise and label them, and bring them to the laboratory. The flowers of sets (i) and (iii) have to be excised at the end of flower longevity and the flowers of set (ii) have to be excised after one visit of the pollinator.
In the laboratory, count the number of pollen grains on each stigma in each set of flowers instead of scoring them for the presence or absence of pollen. For counting pollen grains on the stigma, place the stigma in a drop of acetocarmine and tease the stigma with a pair of needles to release all the pollen grains into the stain. If the stigma is large, remove the debris, lower a cover glass and count the pollen grains carefully (see Protocol 5.5.1 and 5.5.2 for pollen counts).
7.7.6 Estimation of Pollination Limitation
In most of the plants, particularly in large shrubs and trees, a large proportion of flowers do not produce fruits but abscise resulting in low fruit set. Many causative factors for low fruit-set have been identified (Burd 1994; Larson and Barrett 2000; Wilcock and Neiland 2002; Knight et al. 2005). Pollination limitation is one of the major constraints for low fruit set. One of the standard methods to estimate pollination limitation is through pollen supplementation experiment. In these studies, the fruit set in flowers allowed to open pollinate is compared to that in flowers subjected to pollen supplementation (manual pollination).
7.7.6.1 Special Requirements
Tags to label flowers
7.7.6.2 Procedure
-
1.
Tag a suitable number of freshly opened flowers. The number of flowers to be tagged depends on the availability of flowers. At least 50–100 flowers would be needed to get meaningful results.
-
2.
Make them into two sets. Leave set I flowers as such. This set would represent open pollination. Carry out supplementary pollination with cross-pollen (just to avoid self-incompatibility as a factor for lack of seed set if the species is self-incompatible) on flowers of set II.
-
3.
Monitor open-pollinated and manually pollinated flowers and record their fruit set.
-
4.
When the fruits mature, harvest them. Collect seeds from each fruit and count them.
-
5.
Calculate per cent fruit set and average number of seeds per fruit in both the sets and tabulate the results for open- and manually pollinated flowers.
If the per cent fruit set and average number of seeds per fruit in manually pollinated flowers is significantly more than those in open-pollinated flowers, pollination is a constraint. If there is no significant difference between the two sets of flowers, pollination is not a constraint.
7.7.7 Trapping of Flower Visitors
Trapping of floral visitors is necessary for their identification, analysis of their pollen loads and in some experiments for marking them. There are many devises available to collect various floral visitors. Kearns and Inouye (1993) and Dafni et al. (2005) have given a comprehensive account on collecting and preserving pollinators. A few standard methods are elaborated here.
7.7.7.1 Special Requirements
Suitable devise to trap insects. Any of the following devises can be used:
-
1.
Sweeping net: The use of insect sweeping nets, available in various sizes, is the most commonly used method to trap pollinating insects. The sweep should be swift enough to force the insect to the bottom of the collection net; the net should be turned immediately so that the bottom of the bag folds over the rim of the net to trap the insect inside.
-
2.
Aspirator: Another method of collecting insects particularly the smaller ones is to use an aspirator. A manually operated aspirator is available with suppliers of laboratory apparatus or can be easily constructed (see Appendix A.7, Fig. A.3). It consists of a plastic or glass vial of suitable size closed with a rubber stopper with two holes; two hoses are inserted through these holes in the stopper, one is for sucking through user’s mouth and the other is used as the inlet for the insect. The end of the sucking hose inside the vial is tied with a nylon mesh to prevent insects entering the hose during sucking.
-
3.
Wide-mouthed vial of suitable size: We have been employing routinely a wide-mouthed plastic vial (25–50 ml capacity) to collect smaller insects of many species. After the insect has entered the flower, the vial is held in the mouth of the flower. When the insect exits the flower, it enters the vial. The lid of the vial is closed immediately to prevent its escape. Keeping a piece of tissue or cotton dipped in ethyl acetate inside the bottle before trapping would also help in immobilizing (by keeping for a shorter time) or killing (by keeping for longer time) the insect. Ethyl acetate should not be inhaled during the capture of insects.
7.7.7.2 Procedure
-
1.
Capture the floral visitor by using any of the above devises based on availability, convenience and the size of the insect.
-
2.
Transfer the captured insect to a killing jar.
There are several types of killing jars available in the market (see Kearns and Inouye 1993). The killing jars contain a killing agent, usually ethyl acetate or crystals of sodium/potassium cyanide. Ethyl acetate is a routinely used killing agent for insects. Alternatively, a piece of tissue dipped in ethyl acetate may be kept at the bottom of the vial before trapping. When an aspirator is used to trap insects, the killing agent should not be put inside the aspirator; trapped insects should be transferred to a killing jar, similar to those trapped in the sweeping net. The time of exposure of insects to the killing fumes varies and depends on the size and type of insect. Small insects such as bees and flies are killed in minutes, while larger insects, especially beetles, may take many hours.
Caution: Killing agents have to be handled carefully. As they are highly toxic, one should be careful not to inhale the fumes.
Insects can also be killed by placing the vials in a freezer or by transferring them to ethanol. They can be stored in ethanol for longer periods.
-
3.
Killed insects can be pinned to insect boxes and stored for identification or long-term record.
It is more difficult to trap other pollinating agents such as birds, bats and rodents. For catching birds and bats, mist nets have been commonly used. If studies are being carried out in protected forests, permission has to be taken from the authorities to use mist nets for trapping of these animals. Careful observation with a pair of binoculars, photographing (with a zoom lens) and/or video recordings of the animals while foraging may help in their identification without the need for trapping. Their pollination efficiency may be determined by examining the stigmas of flowers after their visits (see Protocol 7.7.3). For trapping rodents such as rats, suitable traps available in the market may be used by keeping bait inside the trap.
7.7.8 Estimation of Pollen Load on Floral Visitors
Studies on pollen load on flower visitors are important for a number of reasons. The presence of pollen load of conspecific species indicates the possible role of the floral visitor in pollination of the species. These studies also indicate whether the visits are confined to flowers of conspecific species or extend also to synchronously flowering heterospecific species. Species with generalist type of flowers (flowers in which rewards are available to any visitor) are visited by a variety of insects and all of them may not be pollinators. Even if they are pollinators, their efficacy may vary. For effective pollination, the visitor should carry sufficient pollen on its body parts that come frequently in contact with the receptive stigma.
It is important to note that most of the pollen harvested actively and packaged as pollen baskets becomes a part of the larval food in the bee hives and is not available for pollination. Also their viability may be affected. It is only the pollen that remains on the body parts that frequently come in stigmatic contact constitute the pollen used for pollination. Often a distinction is made between the ‘total pollen load’ (pollen grains collected from the total wash of the pollinator) from ‘functional pollen load’ (pollen that has the chance to reach the target stigma due to its location on the animal body). From pollination point of view, it is the functional pollen load that is important. In many instances, the presence of pollen load on the visitors’ body may not be sufficient to confirm them as pollinators as their pollen-loaded parts may not come in contact with the stigma. For determining the pollen load, killing of insects in ethanol should be avoided, as it may dislodge pollen grains from the body unless the entire ethanol used for storage is used for pollen counts.
Qualitative analysis of pollen deposition on the foragers may be done with a powerful magnifying glass or under a dissecting microscope. Pollen identification may also be made by scraping some pollen grains from the body onto a slide and observing them under the microscope. They are compared with the reference pollen mounts prepared from the pollen of the focal and co-flowering species for identification. If all the pollen grains on the insect belong to the same species, it indicates the constancy of the visitor during the study period. If the pollen grains are of more than one species, it shows that the visitor forages flowers of different species.
‘Pollen prints’ taken with an adhesive tape can also be used to study pollen grains. For this, a piece of cellotape/adhesive tape is applied onto the surface of the forager’s body, gently peeled and placed on a slide and observed directly under a microscope for identification. Adhesive tapes peeled from different parts of the body can be used to produce pollen distribution map on the body of the animal. The involvement of floral visitor in removing the pollen in species bearing pollinaria such as orchids and asclepiads can be easily recognized, with naked eye or under a stereomicroscope, by the presence of pollinaria on the visitor’s body or by the absence of pollinaria in the flower visited by the insect.
7.7.8.1 Special Requirements
-
Ethanol (95 %)
-
A suitable stain for staining pollen grains such as acetocarmine
-
A watch glass
7.7.8.2 Procedure
Trap the insect and immobilize it as described in Protocol 7.7.7.
For insects with low pollen load:
-
1.
Hold the insect with a pair of forceps and wash it with droplets of 70 % ethanol to remove the pollen from its body. Carefully collect the drippings falling from the insect into a watch glass. Preserve the insect with all the records for identification and as voucher specimen.
-
2.
Count pollen grains present in ethanol. Follow any of the procedures described in Protocol 5.5.1 or 5.5.2 to count pollen grains in ethanol.
7.7.8.3 Modification
If fluorescence microscope is available in the laboratory, it would be more convenient to mount the pollen in a drop of auramine O (see Appendix A.1 for preparation), a fluorochrome that induces bright fluorescence of pollen exine. Observe the slide under the fluorescence microscope using UV combination filters and count them. Pollen grains show bright yellow fluorescence and permit easy identification of pollen grains from debris.
For insects with heavy pollen loads
Additional requirement: rotary shaker/vortex mix:
-
1.
Take immobilized insect in a clean, pollen-free vial. Add 70 % ethanol enough to cover the insect.
-
2.
Put the vial on a rotary shaker for 2–4 h. We have found vigorous shaking of the vial manually or using a vortex mix for a few minutes is sufficient to release all the pollen from insect’s body.
-
3.
Take out the insect from the vial and preserve it for record.
-
4.
Centrifuge the liquid under low speed (ca 2,000 rpm) and remove the supernatant. Use the pollen pellet for counting pollen grains using any of the procedure described under Protocol 5.5.1 or 5.5.2.
The methods described above are not suitable for birds, bats and other vertebrates. Pollen loads can be removed from vertebrate pollinator by using cubes of glycerin jelly (see Appendix A.3) taken with a pair of forceps and gently swabbing the cubes on the body parts of the animal that come in contact with the anthers during their visit to the flowers. The cubes are then mounted on a slide (see Protocol 5.5.5). Slides can be observed soon or stored for later observation. Pollen grains can also be removed through cellotape peeling and observed under the microscope.
7.7.9 Estimation of Density and Diversity of Insect Pollinators
The density and diversity of floral visitors is important in understanding pollination efficiency and pollination limitation across locations and seasons. This is also important in long-time studies aimed at understanding the decline of pollinator populations as a result of human activity such as habitat degradation, fragmentation and application of non-eco-friendly agrochemicals such as pesticides and fungicides.
The most common method used to estimate pollinators’ density and diversity is by net sweeping along the line transects to collect pollinators. Generally, the investigator walks steadily (about 10 m per min) while sweeping with backward and forward strokes at uniform speed. This needs some practice to familiarize the speed of the sweep and pace of the walk before making systematic collection for record. The length of transects varies depending on the density of insects.
The sweeping is done along transects at fixed time intervals during the day or for a selected period. Several investigators have done sweeping along several 50–100 m transects at specific times of the day (at intervals of 1 or 2 h during the peak activity of pollinators). Depending on the need, the sampling is repeated 1 day per week for 3–4 weeks. For crop plants, it is better to sweep the net between the rows of plants.
7.7.9.1 Special Requirement
-
Standard entomological sweeping net
-
Polyethylene bags
-
Screw cap bottles
7.7.9.2 Procedure
-
1.
Select the area to be studied and fix transects.
-
2.
Sweep the net at selected timings with a steady walk along the transect. After the sweeping is complete, twist the net to retain the collections inside the net.
-
3.
The specimens collected are identified either in the field itself or labelled as morphotypes. Count the number of each species/morphotypes.
-
4.
Fix representative samples of all the identified species/morphotypes for documentation or later identification.
-
5.
Release the remaining specimens to avoid reducing the local population of pollinators. It is desirable to deposit the fixed samples in a recognized entomological collection.
7.7.9.3 Modifications
-
1.
Instead of sweeping, pollinators may be recorded through visual observation. In this method, it may not be feasible to record individual species of insects; instead the identification may be assigned to guilds (honeybees, small solitary bees, etc.). This is particularly useful to get abundance data in the canopy of trees in tropical forests. One may use binoculars/telescope, if needed. Recording of each guild in fixed areas in relation to the known number of flowers/inflorescences and scoring each guild over specific time periods give a crude estimate of insect abundance.
-
2.
Different types of insect traps such as sticky traps, water traps, Malaise trap or light traps and window traps have been used to collect insects (see Kearns and Inouye 1993) to get a rough estimate of insect abundance. However, the use of such traps is less useful in studies on pollination ecology as they also collect non-pollinators. Also, all insects present in the area are not trapped as the sensitivity of different insect types to the traps is variable. However, their use may be necessary under some situations particularly if the collections have to be made during night-times (Kearns and Inouye 1993). The use of insect traps permits simultaneous sampling of several locations. They are particularly suitable for long-term studies.
7.7.10 Demonstration of Nocturnal Pollination Based on Day/Night Exposure of Flowers
It is necessary to check for nocturnal pollination in species in which flowers open during the night and those in which flowers remain open during the day and night. Protocols for studying nocturnal pollination are basically similar to those described for diurnal pollination (Protocol 7.7.2), except that the observations have to be made during night-time.
Major difficulties to study nocturnal pollination are visibility and logistics. Also studies on plants growing in reserve forests and sanctuaries require special permission from the authorities to stay in the forest during the night. The presence of wild animals in the forests is another difficulty. Because of the safety reasons, it is always better to study nocturnal pollination with a minimum of two investigators.
One of the indirect methods to check whether pollination takes place only during the day and/or night is by exposing flowers for pollinators only during the day or only during the night and checking their stigmas for pollen load.
7.7.10.1 Special Requirements
Isolation bags and tags
7.7.10.2 Procedure
-
1.
Select flower buds 1 day before anthesis and bag them. Make them into two sets (1 and 2) and label them. It would be convenient to use tags of different colours for the two sets so that they can be easily distinguished. There should be as many buds as possible for each set, preferably 50 each. If the flowers are bisexual, flower buds have to be emasculated before bagging to prevent possible autogamous self-pollination.
-
2.
When the flowers open, remove the bags of set 1 early morning soon after the daybreak to expose them to the pollinators during the day. Rebag them in the evening before it gets dark.
-
3.
In the dusk before it becomes dark, remove the bags from the second set of flowers to expose them during the night. Rebag them in the early morning.
-
4.
Repeat day and night exposure (of the same set of flowers) as long as the flowers remain fresh.
-
5.
When the flowers start senescing, excise the flowers of set 1 and set 2 separately and observe their stigmas individually for pollen load .
-
6.
Calculate per cent pollinated flowers in the two sets. If possible, one can count the number of pollen grains on the stigma and calculate average number of pollen per stigma in both the sets of flowers.
Repeat the protocol for 2/3 batches of flowers to get dependable results.
If the species is nocturnally pollinated, only the flowers exposed during the night show pollen load on the stigma. If the species is diurnally pollinated, only the flowers exposed during the day show pollen loads on the stigma. If pollination occurs during the day as well as night, pollen loads are found on the stigmas of both the sets.
This protocol will not give any information on the nature of pollinators, frequency of their visits and foraging details.
7.7.10.3 Modification
Nocturnal and diurnal pollinations can be identified on the basis of fruit set in the two sets of flowers. Instead of excising them to study pollen load, allow them to develop into fruits and then calculate per cent fruit set in both the sets.
7.7.11 Demonstration of Nocturnal Pollination Based on Direct Observations
As in the diurnal pollination, this protocol also requires some preliminary studies to identify floral visitors and the timing of their visits. Observations to record the details of pollinators’ visits may have to be carried out continuously or at 15/30 min time slots each hour. Usually nocturnal pollination occurs during the crepuscular period (late evening, 18–20 h, and just before dawn, 04–06 h). Depending on the visitation time, observations may be confined to the late evening or early mornings or extended to the whole night. Lack of sufficient light makes it difficult to observe and record the details of nocturnal visitors to flowers. The use of intense artificial light may repel nocturnal visitors. Many investigators have used torch/lantern covered with red cellophane paper for nocturnal observations (Nagamitsu and Inoue 1997). However, the most convenient method to study nocturnal pollination is to use night vision glasses/binoculars which amplify minimal available light to be able to see night floral visitors with reasonable clarity. For larger animals, camera traps can be set up. Camera traps will only confirm the floral visitors at night but do not provide much detail. Video recording is also useful to record nocturnal visitors.
7.7.11.1 Preliminary Investigations
Nocturnal pollinators are mainly represented by bats and moths and to a lesser extent cockroaches, lizards and rodents. Initially the types of flower visitors can be ascertained by using the combination of battery-operated torch and binoculars. After ascertaining the type of pollinator, a group of observers have to position in a selected spot from where a suitable patch of flowers/inflorescences can be clearly seen. In rodent-pollinated plants, flowers are generally positioned closer to the ground. In case of tree species, we have used the canopy of adjoining tree or a collapsible bamboo ladder of suitable height for recording the observations. The details of observations and recordings elaborated in Protocol 7.7.2, for diurnal pollination, have to be followed.
Observations on bat pollination is relatively easier than the other nocturnal pollinators because in typical bat-pollinated plants, the flowers/inflorescences are generally placed outside the canopies of the plants, either as upright racemes or drooping bunches of inflorescences (e.g. Bignoniaceae, Mucuna spp.).
7.7.11.2 Special Requirements
-
Night vision glasses (goggles/binoculars)
-
Isolation bags
-
Hand-held battery-operated torch
-
At least two investigators and any other items needed for safety
7.7.11.3 Procedure
-
1.
Select a patch of freshly opened flowers which can be clearly seen from the observation place.
-
2.
Keep record of floral visitors and duration of their stay on each flower. Make at least 20 observations of each visitor to flowers (see sample Table 7.3 for details of recording).
-
3.
Record the foraging behaviour of floral visitors and whether they forage nectar or pollen.
-
4.
Excise flowers visited by nocturnal visitors and keep them in a beaker or Petri plate without causing disturbance.
-
5.
Bring excised flowers to the lab in the morning and observe their stigmas for the presence of pollen grains (see Protocol 7.7.3 for details).
-
6.
Plot a graph on visitation frequency (and if data is available, with the temporal details of nectar production).
7.7.11.4 Modifications
-
1.
Nocturnal pollination on the basis of fruit set: Flowers visited by nocturnal agents can be bagged to study fruit set.
-
2.
If pollinators happen to be small insects, they can be trapped by tying sticky traps (3 × 3 cm, available commercially or prepared by smearing nondrying glue, see Protocol 7.7.12 for details) near the flowers/inflorescences. Once the insects come in contact with the tape, they get stuck to the tape. Although it is difficult to remove the insects from the tape, they can be observed under the microscope for pollen load. However, it should be kept in mind that insects caught on the tape are not necessarily the pollinators as many of them may not even visit the flowers.
-
3.
As insects are attracted towards light, they can be conveniently trapped by using a suitable setup near the focal plant when in bloom. One may use a setup comprising a small battery-operated torch covered with blue cellophane paper to impart blue light. The torch then can be placed in bigger size glass jar. Once the insects enter the jar, cover it with a lid. This trapping exercise may be repeated at several locations in the population.
The collected insects can be immobilized and studied for pollen load using the method described in Protocol 7.7.8.
It may not be feasible to trap larger animals. Their role in pollination may be confirmed by allowing them to visit virgin flowers and either excising them to observe pollen load on their stigmas or bagging them to study fruit set.
7.7.12 Demonstration of Ant Pollination
Ants are found on flowers of a number of species. As pointed out earlier, ant pollination is rare and has so far been reported only in a limited number of species. It may be, to some extent, due to the perception of the investigators not to consider them as effective pollinators, even when they are present on the flowers. When ants are present on the flowers of the focal species, it would be necessary to study the details at least to rule out their role in pollination. When ants are present, they are generally present during the day as well as night.
7.7.12.1 Preliminary Studies
This protocol also requires some preliminary studies to understand the behaviour of ants on the flowers and to standardize a suitable method to prevent and permit ants to the flowers used for observation. If these studies clearly establish that the ants do not come in contact with the anthers or the stigma, their role in pollination can be ruled out. If there are possibilities of ants coming in contact with the stigma, the details have to be investigated to study their role in pollination.
Several investigators have used insecticides to prevent ant visits. In herbaceous plants, insecticide can be applied to the soil around the plants. If they are large shrubs or tree species, insecticide has to be sprayed on flowering branches bearing focal flowers. Also, it requires standardization of the concentration and frequency of sprays to prevent ants during the observation period. Some investigators have applied glue (Tanglefoot) around the stem suspending the inflorescences or on the plasticine placed around the whole plant on the ground to prevent ants visiting the flowers (Gomez et al. 1996); they have used nylon mesh bags (0.25 mm mesh) around the flowers to permit visits of only ants to the flowers.
We have used a nondrying strong glue commercially available (Stickem special/Stickem ribbon; Seabright laboratories, Emeryville, CA, USA) to prevent ants’ visit to the flowers. This glue is commonly used to make sticky traps for small insects and it does not dry up for several days. The following is the detailed procedure we have followed to prevent ants from entering focal flowers and to permit only ants to visit the flowers:
Smear Stickem special glue around 2–3 cm of the stem of the flowering branch proximal to the flowers/inflorescences. Crawling ants get stuck to the glue and cannot pass through the glue region, thus preventing ants from reaching the flowers. As an additional precaution, we have bagged (made up of pollen proof cotton cloth) the flowering branch tightly around the stem (distal to the glued part) without leaving any space for the movement of ants into the bag. This dual method prevented ant visits to focal flowers completely.
To study the visits of other insects to focal flowers, we open the bags at their tips and pull them back during the day to expose diurnal pollinators to visit the flowers. The bags are closed in the evening. For allowing only ants to visit the flowers but not any flying insects, we have found that conventional pollination paper bag loosely tied, leaving sufficient gaps around the stem of flowering branches, allows ants to visit the flowers readily.
7.7.12.2 Special Requirements
-
Tags
-
Pollination bags (made up of pollen proof cloth and of paper)
-
Nondrying glue
7.7.12.3 Procedure
-
1.
Prepare the following four sets of flower buds one day before opening:
-
(i)
To prevent ants from entering the flowers by using the method explained above. If individual flowers are used for bagging, smear the nondrying glue around the lower part of the pedicel or on the stem suspending the flower and bag them. If the inflorescence/flowering branches are used for bagging, smear the glue around the stem of the branch/inflorescence and remove opened flowers, very young buds and fruits, if any, before bagging and retain only those buds that would open the next day. Also remove any ants present on the flowers/inflorescences. This set will prevent ants as well as any other insects entering the flowers and indicates the extent of autogamy, if any, in bisexual flowers.
-
(ii)
To allow only ants to enter the flower but not any other flying insects as explained above (by conventional loose bagging without applying nondrying glue).
-
(iii)
To prevent ants but permit other flying insects to enter the flowers (treatment similar to set i), but the bags have to be opened at the tip and pulled back in the morning to allow diurnal insect visits and again closed in the evening. This has to be repeated every day until the flowers in the bag remain fresh.
-
(iv)
To allow all floral visitors by tagging the flowers without bagging them or smearing nonadhesive glue around the stem/pedicel.
-
(i)
-
2.
Maintain the above sets until the flowers start senescing.
-
3.
After initiation of senescence, excise all four sets of flowers and maintain them separately in Petri plates. Handle the flowers carefully to prevent disturbance and possible autogamous pollination (in bisexual flowers) during handling. Take the flowers to the laboratory.
-
4.
Observe the stigma of each flower of each set and record the presence of pollen on the stigma. If possible, count the number of pollen grains on each stigma. Calculate per cent of flowers pollinated and average number of pollen grains present on the stigma of each set.
Ants can be easily lifted with the help of forceps and dipped in 70 % ethanol to study pollen load on their body as described in Protocol 7.7.8.
The presence of pollen on the stigma in significantly greater per cent of flowers in set (ii) than in set (i) demonstrates the role of ants in pollination. Results of set (iii) indicate the role of flying insects in pollination. The results of set (iv) indicate pollination efficiency under field conditions from all the pollinators combined.
The above treatment and observations of the stigma should be replicated using different branches and plants to get a clear information on the role of ants in pollination.
7.7.12.4 Modifications
-
1.
Ant pollination on the basis of fruit set: Retain focal flowers on the plants until fruit development and analyse per cent fruit set and mean number of seeds per fruit in each set of the treatments.
-
2.
Frequency of visits and pollination efficiency of each pollinator can be studied by recording the visits of each visitor (see Protocol 7.7.2 for details). To determine the frequency of ants, observe a set of open flowers and score the frequency of ant visits; chase away any other flying insects from visiting the focal flowers. The frequency of nonflying visitors can be scored in set iii flowers when the bags are opened during the day.
-
3.
It would be useful to make nocturnal observations also if ants turned out to be effective pollinators. Check if ants are present on the flowers during night-time also and if so whether the species present is the same or different. If necessary, their role in pollination can be confirmed by setting up treatment i and ii described above under the ‘Procedure’.
7.7.13 Demonstration of Wind Pollination
Next to biotic pollination, wind is the major pollinating agent. A standard method used to demonstrate wind pollination is to use mesh bags that permit airborne pollen but not any insects.
7.7.13.1 Special Requirements
Insect- and pollen-proof pollination bags
Pollination bags of desired mesh size. Bags of mesh size 1 or 2 mm2 are considered suitable to allow entry of air borne pollen but not the insects. Many workers have used double layered insect netting for preparing such bags. We have found the bags stitched from mosquito net cloth are satisfactory to demonstrate wind pollination. Although mesh bags are effective in preventing insects from bringing about pollination, they do, to some extent, reduce the efficiency of wind pollination as some air borne pollen is likely to get trapped in the mesh and also they may disrupt air currents around the flowers.
7.7.13.2 Procedure
-
1.
Select older flower buds about one day before anthesis. If the flowers are small and cannot be bagged, select inflorescences or flowering branches containing as many buds of suitable stage as possible (see Sect. 2.2). If inflorescences or the flowering branches are selected, remove young buds, opened flowers and fruits, retaining only the flower buds about one day before anthesis.
-
2.
Make them into the following three sets:
-
(i)
Flowers allowed to open pollinate
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(ii)
Flowers enclosed in insect- and pollen-proof bags (to exclude air- and insect-borne pollen)
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(iii)
Flowers enclosed in bags of suitable mesh size that exclude insects but not airborne pollen
Emasculate buds of set (ii) and (iii) before bagging them to prevent autogamy.
Label and record the number of flower buds in each set.
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(i)
-
3.
Remove the bags in sets (ii) and (iii) after the stigmas lose receptivity but keep the tags intact. This period may vary between species.
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4.
Keep track of the three sets of flowers until abscission or fruit development. Maintain the record of abscised flowers.
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5.
Collect mature fruits before their dehiscence from each set, and count the number of seeds from each fruit.
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6.
Calculate per cent fruit set and average number of seeds per fruit in each set of flowers. Compare fruit and seed set in the three sets of flowers.
Flowers isolated with pollen- and insect-proof bags should not set any fruits unless the species is an apomict (which does not require pollination). Fruit set in flowers isolated with mosquito net bags (but not in insect-proof bags), if any, represents fruits developed exclusively as a result of wind pollination. As the mosquito net bags also prevent a proportion of pollen from reaching the stigma, fruit set may be marginally lower in wind-pollinated flowers when compared to open-pollinated flowers.
7.7.13.3 Modification
The protocol can be modified to assess wind pollination on the basis of pollen load on the stigma. Instead of allowing the three sets of flowers for fruit development, excise senescing flowers from each set, and study pollen load on the stigma (see Protocol 7.7.3).
7.7.14 Use of Slide Traps for Airborne Pollen
The release of pollen grains to ambient air and their movement in the air is the basic requirement for wind pollination. This can be easily demonstrated by trapping pollen grains present in the air by using various devices. Although the presence of pollen on slide traps does not demonstrate wind pollination, it provides strong evidence for the involvement of wind in pollination. The use of slide traps is one of the simplest pollen traps.
7.7.14.1 Special Requirements
Prepared microscopic slides: The slides used to trap pollen have to be prepared before they are exposed to collect air borne pollen. As pollen grains trapped on slides need to be quantified generally as number of pollen grains per cm2, marking the 1 cm2 squares on the slide with a marking pen before exposing would make it convenient to count. Mark 3 or 4 squares on one surface of the slide. These squares have to be divided into smaller rectangles by marking about 1 mm parallel lines across each square. This is to facilitate counting of pollen grains between two adjacent lines after trapping pollen grains (see Protocols 5.5.1 and 5.5.2). On the other side of marked slide (which is not used for marking), rub glycerin jelly (see Appendix 13.3 for preparing glycerin jelly) or petroleum jelly (available with chemists) as a thin film. This will help pollen grains adhere to the slides. It is convenient to prepare 20–30 slides at a time and use them as and when needed. Dry the slides in pollen-free and dust-free room and keep them in slide boxes to prevent contamination with pollen.
Thread to hang slides
7.7.14.2 Procedure
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1.
Tie prepared slides with a thread on one end (for vertical exposure) or both the ends (for horizontal exposure). Label the slides with permanent ink on one side of the glycerin jelly-coated surface. Hang one set of slides horizontally (keeping the glycerin-coated surface up) and another set vertically on and around the plant at various distances from the flower. If the flowers are unisexual, the slides have to be exposed with reference to the male as well as the female flowers. If it is a tree species, place the slides at different levels in the canopy and known distances away from the canopy. Record the placements with reference to each numbered slide.
-
2.
Collect exposed slides individually after 24 h, and keep them in a slide box until used for observation.
-
3.
Observe the glycerin-coated surface of the slides under a microscope for the presence of pollen grains of conspecific species. If pollen grains cannot be identified easily, put a drop of acetocarmine, and lower a cover glass before observing. We have found that mounting the slides in a drop of auramine O and observing them under a fluorescence microscope (UV excitation) is very convenient. Auramine O induces bright fluorescence in pollen exine making it easier to distinguish pollen grains from the dust/debris. If pollen grains are present, count the number of pollen grains in each pre-marked cm2 (see Protocol 5.5.1 for pollen counting).
7.7.14.3 Modifications
-
1.
If there is any difficulty in identifying the presence of conspecific pollen on the slide or if there are pollen grains belonging to different species, prepare reference pollen slides of the conspecific and co-flowering species by mounting their pollen grains in glycerin jelly (see Protocol 5.5.5). Identify conspecific pollen grains present on exposed slides with the help of pollen on the reference slides. It would be convenient to prepare reference slides of focal and co-flowering species at the beginning of the study, and keep them in a slide box as permanent records, and use them whenever required.
-
2.
Standard pollen traps can also be used to demonstrate the presence of pollen grains of a given species in the air. Burkard Portable Pollen Sampler, which can be operated by a battery, is the most convenient. The details of these traps can be found in Kearns and Inouye (1993) and Dafni et al. (2005). Instruction manual gives the details of the procedure to be followed.
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Shivanna, K.R., Tandon, R. (2014). Pollination Ecology. In: Reproductive Ecology of Flowering Plants: A Manual. Springer, New Delhi. https://doi.org/10.1007/978-81-322-2003-9_7
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