Article | Open | Published:

Nepenthes pitchers are CO2-enriched cavities, emit CO2 to attract preys

Scientific Reportsvolume 7, Article number: 11281 (2017) | Download Citation

Abstract

Carnivorous plants of the genus Nepenthes supplement their nutrient deficiency by capturing arthropods or by mutualistic interactions, through their leaf-evolved biological traps (pitchers). Though there are numerous studies on these traps, mostly on their prey capture mechanisms, the gas composition inside them remains unknown. Here we show that, Nepenthes unopened pitchers are CO2-enriched ‘cavities’, when open they emit CO2, and the CO2 gradient around open pitchers acts as a cue attracting preys towards them. CO2 contents in near mature, unopened Nepenthes pitchers were in the range 2500–5000 ppm. Gas collected from inside open N. khasiana pitchers showed CO2 at 476.75 ± 59.83 ppm. CO2-enriched air-streaming through N. khasiana pitchers (at 619.83 ± 4.53 ppm) attracted (captured) substantially higher number of aerial preys compared to air-streamed pitchers (CO2 at 412.76 ± 4.51 ppm). High levels of CO2 dissolved in acidic Nepenthes pitcher fluids were also detected. We demonstrate respiration as the source of elevated CO2 within Nepenthes pitchers. Most unique features of Nepenthes pitchers, viz., high growth rate, enhanced carbohydrate levels, declined protein levels, low photosynthetic capacity, high respiration rate and evolved stomata, are influenced by the CO2-enriched environment within them.

Introduction

Nepenthes consists of approx. 160 currently described species distributed in the Madagascar-south east Asia-north Australia-New Guinea region, with hotspots in Borneo, Sumatra and the Philippines. They grow in wet, sunny and nutrient (N, P)-poor habitats. In order to supplement this nutrient deficiency, they evolved strategies to capture insects and other arthropods through their modified leaf tips (pitchers or pitfall traps)1,2,3,4,5,6,7,8. The known factors attracting arthropod preys into the ‘passive’ Nepenthes traps are nectar, olfactory cues, colour and UV/fluorescence patterns1, 3, 6. Toxic metabolites, waxes, physical phenomena, viscoelastic pitcher fluid, chitinases/proteases and antifungal metabolites are also involved in various stages of carnivory displayed by these unique plants2, 4, 5, 7. Other than ‘arthropod trapping strategies’, recent reports show that, pitchers of Bornean Nepenthes species display ‘mutualistic interactions’ with tree shrews, bats and other small mammals, and thereby gain nutrients8.

Nepenthes leaves are highly specialized with two distinct portions, lamina and the pitcher (prey trap). The midribs of Nepenthes leaves protrude from the leaf tip into tendrils, form small buds which inflate into bulb- or tube-shaped pitchers. In other words, Nepenthes pitchers are modified epiascidiate leaves in which their adaxial (upper) surface curls around and fuses to form the inner side of the pitcher1. The tendrils of aerial pitchers are usually coiled in the middle, and once in contact with other objects for long enough they curl around them, forming anchor points for pitchers. In this way, Nepenthes tendrils help to support the growing stem of the plant. As it matures, the pitcher inflates and gets partially filled with an acidic enzymatic fluid. Pitchers also have a flap (operculum), which initially seals (‘hermetically seals’) the growing trap1, and once mature breaks open for prey capture. In N. khasiana, initial development stages to lid opening of pitchers take about 3 weeks. N. khasiana pitchers grow up to an average of 13 cm length, with lid length 3 cm and pitcher fluid 3.25 mL. In most Nepenthes species, the lid covers the pitcher opening and thus protects it from rain, preventing dilution of the pitcher fluid, but in some species the lids are reduced or bent backwards9. Once open, pitcher rims (peristomes) play major initial steps in attracting and capturing preys2. Nepenthes species show considerable variations in size, shape and colour of their pitchers (Fig. S1) and peristomes (Figs S2S4). N. rajah, largest pitcher/carnivorous plant, grows up to 3 m in height, its pitchers grow up to 30 × 14 cm (height or length x width) and secrete up to 2.5 liters of pitcher fluid. The recently described species, N. attenboroughii and N. palawanensis, also produce large pitchers10. Large Nepenthes pitchers are capable of trapping rodents, lizards and birds. Once open, Nepenthes pitchers involve in prey capture from a few weeks up to nine months depending on the species11.

Nepenthes pitchers are even described as ‘hollow leaves’ in the literature12, but they are not entirely ‘hollow’. Unopened N. khasiana pitchers, on pressing with our hands, give a gas-filled sensation, and on further forcing they burst open mostly at the peristome-lid portion. Nepenthes pitchers, their growth, morphology, prey capture, mutualistic interactions, digestion mechanisms and nutrient uptake received lot of attention in recent decades2, 4, 8, 13. But, the gas composition inside Nepenthes pitchers has not been studied so far. In our preliminary tests, we found high levels of CO2 inside growing, unopened N. khasiana pitchers. This led us to look into the role of this CO2 in prey capture, growth and other unique features of Nepenthes pitchers.

Results

CO2 in Nepenthes pitchers, prey capture

We found growing, unopened pitchers of N. khasiana (Fig. S1a) filled with high levels of CO2 (4053.76 ± 1188.84 ppm, n = 9), along with ambient levels of O2, CO, CH4 and N2O. Various Nepenthes hybrids also showed high contents of CO2 in their growing (unopened) pitchers (Fig. S1b–g) (Nepenthes hybrid 01, NH01 3114.38 ± 973.52 ppm, n = 5; NH02 4008.67 ± 1042.38 ppm, n = 3; NH03 3390.03 ppm, n = 1). Gas samples from inside (just below the peristomes) open N. khasiana pitchers showed CO2 levels at 476.75 ± 59.83 ppm (n = 6). Moreover, open N. khasiana pitchers when their lids sealed back (after 24 hours of lid opening) regained the high CO2 levels (3231.33 ± 762.58 ppm, n = 3). Mature, unopened pitchers when cut open and sealed again (after 24 hours) also showed high contents of CO2 inside them (3324.00 ± 959.23 ppm, n = 3). Ambient CO2 levels at the Nepenthes experimental fields were 396.97 ± 6.07 ppm, n = 3, matching global measurements.

Near mature, unopened N. khasiana pitchers, when cut open and quickly re-weighed, showed noticeable reduction in their weights (N. khasiana pitcher length 12.94 ± 3.11 cm, lid length 3.01 ± 0.79 cm, pitcher fluid 3.25 ± 2.29 mL, weight difference 2.50 ± 1.58 mg, n = 45) (Table S1). N. khasiana individual pitchers showed weight differences from 0.80 to 8.50 mg (one exceptionally big pitcher) (Table S1). Nepenthes hybrid pitchers also showed similar weight reduction viz., 0.70 mg (NH 05) to 5.50 mg (NH 01) (details in Table S1).

We passed a stream of CO2-enriched air (1% CO2 in air) through the upper portion (above the liquid zone) of just opened N. khasiana pitchers in the field for 12 days. This CO2-enriched air, mixed with the gas inside the pitcher and discharged through the top of open pitchers (CO2 at 619.83 ± 4.53 ppm, n = 6), attracted substantially higher number of aerial preys (insects) (31.17 ± 11.91, n = 6, Fig. 1) into these traps. In control experiments, when a stream of air at the same flow rate was passed through N. khasiana pitchers (CO2 at 412.76 ± 4.51 ppm, n = 6) for 12 days, we found a relatively lower rate of insect capture, 16.2 ± 5.15 (n = 6). Capture rate in normal (unmodified) pitchers (CO2 at 476.75 ± 59.83 ppm, n = 6) for 12 days was 19.67 ± 5.43 (n = 6) (Fig. 1).

Figure 1
Figure 1

Prey capture rates enhance on streaming CO2 through Nepenthes pitchers. Preys captured in 12 days by [1]: air-streamed (control) N. khasiana pitchers (mean ± s.d., n = 6); [2]: normal (unmodified) N. khasiana pitchers (mean ± s.d., n = 6); [3]: CO2-streamed (test) N. khasiana pitchers (mean ± s.d., n = 6; *significant at p < 0.05, compared to [2]).

CO2 in Nepenthes pitcher fluid

On the average, N. khasiana pitchers produce pitcher fluids at 3.25 ± 2.29 mL (n = 45) (Table S1). Our data show that closed (unopened) N. khasiana pitchers have CO2-enriched gaseous media above their aqueous pitcher fluids, and CO2 remains in equilibrium with these fluids. Partial pressures of oxygen (pO2) and CO2 (pCO2) in mature, unopened N. khasiana pitcher fluids were measured as 140.83 ± 7.60 (n = 6) and 20.47 ± 1.53 mm Hg (n = 6), whereas pO2, pCO2 for opened, prey captured pitcher fluids were 76.78 ± 18.10 (n = 6) and 21.43 ± 2.85 mm Hg (n = 6), respectively (Fig. 2). pO2 and pCO2 in the atmosphere are 159 and 0.30 mm Hg, respectively. We also detected CO2 dissolved in N. khasiana pitcher fluid by headspace GC-MS (VF-5 column, ret. time 1.65 min; EI-MS, m/z: 44 (M+), 32). Mass data of CO2 from the pitcher fluid matched with its authentic standard. We measured the pH of unopened N. khasiana pitcher fluid as 3.54 ± 0.09 (n = 4), and on prey capture the fluid became more acidic (pH 2.47 ± 0.25, n = 4).

Figure 2
Figure 2

Partial pressures of CO2 (pCO2) and oxygen (pO2) in mature, unopened and opened, prey captured N. khasiana pitcher fluids. pCO2 in (1) mature, unopened and (2) opened, prey captured pitcher fluids (mean ± s.d., n = 6); pO2 in (3) mature, unopened and (4) opened, prey captured pitcher fluids, respectively (mean ± s.d., n = 6; *significant at p < 0.05, compared to (3)).

CO2, lid opening, chemical defense

We observed prey captured pitcher fluids in open N. khasiana pitchers turning yellow whereas fluids in netted, open pitchers (with no ants or insects captured) remained colourless. DART-MS of yellow pitcher fluids showed droserone (MW 204.18) and 5-O-methyl droserone (MW 219.00) in them (Figs S5S7). Chitin induction, mimicking prey capture, into N. khasiana pitcher fluid also turned it yellow and demonstrated the release of these antifungal metabolites in DART-MS5 (Figs S5S7).

CO2, stomata in Nepenthes pitchers

In SEM images, we found N. khasiana leaves (laminae) hypostomatic i.e., stomata observed only in their abaxial (lower) sides (Fig. 3a), and not in adaxial (upper) sides. But N. khasiana pitchers (both unopened and open pitchers) showed stomata in their outer sides, and ‘modified stomata’ in their inner sides (Fig. 3b–e). No stomata were seen at the inner sides of N. khasiana pitcher lids (Figs S8S10). Stomata in the abaxial sides of the leaves and at the outer sides of pitchers were normal ones with two guard cells (Fig. 3a–c), whereas stomata inside the pitchers were modified ‘lunate cells’, pointing downwards, with only one guard cell (Figs 3d,e and S3). These modified stomata inside the pitcher were found embedded in crystalline epicuticular wax layers (Fig. 3d,e).

Figure 3
Figure 3

SEM images of root, leaf, tendril and pitcher of N. khasiana. (a), Stoma at the leaf abaxial (lower) surface. (b) Outside surface of pitcher with low stomatal density (stomata circled). (c) Stoma at the outside surface of pitcher. (d) ‘Stomatal’ distribution at the inner surface pitcher. (e) ‘Modified stoma’ at the inner pitcher surface, embedded in wax crystals. (f) Inner liquid zone of the pitcher showing glands. (g) Expansion of a secretory gland. (h) Tendril cross section, showing vascular openings. (i) Tendril cross section, outer layer. (j) Tendril cross section, central portion. (k) Root cross section with vascular openings. (l) Root cross section showing starch granules.

CO2, trichomes, prey capture

Leaf abaxial and adaxial sides of N. khasiana showed only glandular trichomes (data not shown) at a low density. Branched non-glandular and glandular trichomes were observed on N. khasiana tendrils (partially seen in Fig. 3h), at the outer sides of their pitchers and upper sides of their lids (Figs 3b and S11S13), glandular trichomes only were found in the inner sides of lids (Figs S8S10), and no trichomes were observed in other inner sides of pitchers (peristome, slippery and digestive zones) (Figs 3d–g and S2S4).

Respiration as CO2 source within Nepenthes pitchers

SEM micrographs of N. khasiana tendrils and roots (Fig. 3h–l) showed numerous hollow channels (vascular bundles) within them. Starch granules deposited in root cross sections were also observed in the SEM (Fig. 3l). But, no gas flow was detected from tendril (cross-section) into the pitcher cavities (Methods, Field studies). In our comparative measurements, photosynthetic rates (A N) of N. khasiana laminae and pitchers were 3.68 ± 0.53 μmol CO2 m−2 s−1 (n = 6) and −0.60 ± 0.22 μmol CO2 m−2 s−1 (n = 6), respiration rates (R D) 0.82 ± 0.18 μmol CO2 m−2 s−1 (n = 6) and 1.55 ± 0.36 μmol CO2 m−2 s−1 (n = 6) and maximum quantum yield of PSII (F V/F m) 0.80 ± 0.01 (n = 8) and 0.67 ± 0.07 (n = 8), respectively.

CO2, Nepenthes pitcher growth, C/N ratio

We verified the growth rate of N. khasiana pitchers on release of CO2 (within them) against normal CO2-filled pitchers. N. khasiana pitchers in early growth stages, when cut to release the elevated CO2 within them showed diminished growth compared to control pitchers. N. khasiana cut pitchers: initial stage of 6–8 cm to lid opening, average growth of pitchers 6.84 ± 2.03 cm, n = 45; pitcher growth in cm per day 0.61 ± 0.15, n = 45 (Table S2). N. khasiana uncut (control) pitchers: initial stage of 6–8 cm to lid opening, average growth of pitchers 8.00 ± 2.27 cm, n = 45; pitcher growth in cm per day 0.71 ± 0.17, n = 45 (Fig. 4 and Table S2). Growth rate (in cm per day) was diminished by 14.08% in cut pitchers. Growth rate was minimized (to zero) on lid opening of all (cut/uncut) pitchers.

Figure 4
Figure 4

CO2 released-N. khasiana pitchers show diminished growth rates. Blue, cut pitchers, average growth 0.61 ± 0.14 cm per day, mean ± s.d., n = 45; black, uncut pitchers, average growth 0.71 ± 0.17 cm per day, mean ± s.d., n = 45; straight lines display respective averages (more details in Table S2).

We found high carbon and low nitrogen contents in N. khasiana leaves (C 45.72 ± 2.43%, N 2.14 ± 0.30%, n = 4) and pitchers (C 39.07 ± 1.94%, N 1.50 ± 0.25%, n = 4).

Discussion

Our data demonstrate that Nepenthes unopened pitchers are CO2-enriched ‘cavities’, when lids open they release CO2 at a high 3000–5000 ppm to an ambient ~400 ppm atmosphere, and then continue releasing CO2 resulting in its gradient surrounding them. Weight (difference) measurements of Nepenthes pitchers indicate the release of a denser gas (CO2; density CO2/air 1.980/1.225 kg/m3) within them, filled at a slightly higher pressure compared to the atmosphere. Nepenthes pitchers generally stay in ‘upright’ position, and the gas within is emitted through the pitcher-lid opening.

Open pitchers of N. khasiana are constant emitters of CO2 (476.75 ± 59.83 ppm, n = 6), a sensory cue. Most insects pay special attention to ‘subtle variations’ or ‘gradients’ of CO2 in the form of plumes arising from individual point sources14, 15. Insects have well developed CO2 receptors which can detect these variations (even small variations) as a means of locating their food14. Moreover, CO2 emitting devices are widely used as traps against mosquitoes, flies and other insects16. In this study, on CO2-streaming (1% CO2-enriched air) for 12 days through N. khasiana pitcher tops, we found a substantial increase in aerial preys (insects) captured within them (preys captured = 31.17 ± 11.91, n = 6), compared to air-streamed control pitchers (same flow rate; preys captured = 16.20 ± 5.15, n = 6) and unmodified (normal) pitchers (preys captured = 19.67 ± 5.43, n = 6) (Fig. 1). These counts are excluding the ants (dead) crawled into these pitchers from the ground, and the ants count did not show any pattern between the CO2-enriched, air-streamed and unmodified (normal) pitchers. In Fig. 1, the insect capture rates in these three different experimental conditions (CO2-enriched, air-streamed and unmodified pitchers) are proportional to the CO2 emission rates from N. khasiana pitcher tops. These data demonstrate CO2 as an insect attractant emitted by Nepenthes prey traps, and reveals a new prey capture mechanism within them.

Most Nepenthes species secrete pitcher fluids with viscoelastic properties. Fluids in unopened pitchers are sterile17, and once open microbes and inquilines invade them. Our results show that a high level of CO2 is dissolved in N. khasiana pitcher fluids (Fig. 2). Open, prey captured pitcher fluids showed low levels of O2 (Fig. 2), and very low (even anoxia) or decreasing levels of oxygen were reported in Sarracenia purpurea, Utricularia and Genlisea traps18,19,20. Dissolved CO2 in Nepenthes pitcher fluid instantaneously forms equilibrium with its hydrated form H2CO3 which dissociates into H+ and HCO3 − 21. The relative changes to any one of these molecules/ions control the pH and optimum activity of the digestive enzymes secreted into the pitcher fluid by specialized glands21 (Fig. 3f,g). N. khasiana pitcher fluids are acidic, and lid opening and prey capture further reduce the pH. Similar pH trends were observed in pitcher fluids of several Nepenthes species4, 5, 11, 22. pH reduction on prey capture (even after release of the high levels of CO2 on pitcher opening) is critical for optimum enzyme activity (prey digestion) and absorption of nutrients, and this is achieved through a proton (H+) pump22,23,24. This acidic pH could also be controlling the growth of pitcher inhabitants (microbes, mosquito larvae, small aquatic organisms etc.). CO2 dissolved in the pitcher fluid is one of the factors making it acidic and it also acts as a preservative to the pitcher fluid.

Once Nepenthes pitchers become mature, their ‘tight lid sealings’1 open and release the elevated CO2 within, making them ready for prey capture. The sequential events of lid opening, CO2 release and prey capture are sensed by these plants, and they release antifungal naphthoquinones (droserone, 5-O-methyl droserone, plumbagin, 7-methyl juglone) into the pitcher fluid (Figs S5S7), preventing infections from incoming preys5.

Stomata are small pores controlling gas exchange, mainly CO2 and water vapour, found in leaves and other organs in plants25. Stomata inside N. khasiana pitchers were ‘modified’, pointing downwards, with only one guard cell (Figs 3d,e and S3). Similar modified stomata embedded in wax crystals were observed by SEM studies in the inner sides of pitchers of N. rafflesiana 11, 12, 22, N. alata 1, 3, 26, N. mirabilis 3, N. diatas 23 and other Nepenthes species/hybrids27, 28. Most authors described these stomata as ‘transformed’ or as ‘lunate cells’ with a convex structure in the inner surface of Nepenthes pitchers, and explained this modification as an evolutionary adaptation contributing to prey capture by disrupting the adhesion of insect feet and blocking entry of their claws12, 22, 23, 26. Owen and Lennon, 1999 suggested the function of this ‘modified stomatal complex’ as ‘water secretion’ or ‘gas exchange’ or even as a ‘mystery’1. But, absence of ‘pores’1, 22 in these ‘modified stomatal structures’ nullifies the chances of them functioning as vents in ‘gas exchange’.

Similar to our observations, Pavlovič and co-workers reported stomata on the abaxial sides of laminae of N. alata and N. mirabilis, and very low stomatal density in Nepenthes pitchers3. Other studies also reported modified stomata at the interior of pitchers and overall low stomatal density in pitchers of various Nepenthes species1. Stomatal distribution in laminae (abaxial, high) and pitchers (stomata with two guard cells, outer side; low) are matching with their photosynthetic capacities, high (laminae) and very low (pitchers). In most cases, we found high density of the ‘modified stomata’ at the pitcher inner (top) sides (Fig. S3)1, 9, 23. In Nepenthes, pitchers are formed by the folding of leaves with their adaxial (upper) surfaces turning into inner sides of these traps1. It is significant that, the leaf upper surfaces are devoid of stomata, but the pitcher inner surfaces ‘evolved’ these ‘modified stomata’ (Fig. 3d,e). In pitchers of Sarracenia, Darlintonia, Heliamphora and Cephalotus, stomata (normal) are found in their outer surfaces or in their lids/‘hoods’, and ‘stomata-like structures’ present within their pitcher tubes are ‘permanently open’ and not ‘functional’12. It is proven that increase in CO2 even in the range of 100 ppm has a profound effect on the stomata (modifies their morphology) in plants14. The transformed stomatal aperture with a single guard cell (Fig. 3d,e) at the interior (only) of Nepenthes pitchers is most probably a manifestation of the high CO2 (approx. 4000 ppm, nearly 10 times the ambient) atmosphere within them. But, evidences gathered so far are not conclusive on the function of these ‘modified stomata’ or ‘lunate cells’ (Figs 3d,e and S3)1, 12. Crystalline epicuticular wax in thick layers, as observed in the upper part of inner pitcher walls of N. khasiana and several other Nepenthes species, is not distinctly seen in other portions of the pitchers (lid, peristome, liquid zone, outer surface) and in the abaxial and adaxial sides of their leaves (Fig. 3). These inner waxy layers define the hydrophobic slippery zone, which minimizes insect attachment. Recent evidences also demonstrate high level of CO2 as a factor which enhances cuticular wax density in plants29. Nepenthes prey traps display a unique natural model of evolution of stomata in a CO2-enriched atmosphere.

Trichomes, a group of epidermal microstructures, carry out diverse functions in plants, and in carnivorous plants one of their roles is facilitating prey capture30, 31. In fact, relatively high density of branched trichomes was observed at the top outer sides N. khasiana pitchers and their lids1 (Figs S11S12), and no trichomes were observed in deep interior of the pitchers. But, significantly, Sarracenia, Heliamphora, Darlingtonia and Cephalotus pitchers have trichomes in their interior zones, including their innermost digestive zones1, 12, 32. Branched trichomes on the exterior of Nepenthes pitchers (and their lids) provide a foothold to the visitors (termites, ants etc.)30, 31, enhancing the chances of their ultimate ‘luring’ to the interior of the traps. Edible trichomes in N. albomarginata are known to ‘lure’ termites into their pitcher traps23, 33, 34. Elevated CO2 within Nepenthes traps could be one factor reducing the trichome density (particularly branched ones) in the inner sides of Nepenthes pitchers35.

SEM micrographs showed numerous vascular bundles within the roots and tendrils of N. khasiana (Fig. 3h–l), but no gas flow was detected from tendril (cross-section) into the pitcher cavities. Respiration (dark) rates of non-carnivorous herbaceous plants are typically less than 50% of their photosynthetic rates, but, the average respiration/photosynthetic rate in terrestrial carnivorous plants is as high as 63%36. Again, the traps (pitchers, snap trap) of terrestrial carnivorous plants (Nepenthes, Sarracenia, Dionaea muscipula) showed much higher respiratory costs (respiration/photosynthetic rate 158%) than their laminae (lamina, phyllodia, petiole) (respiration/photosynthetic rate 19%)36. More evidences for higher respiration rates (in traps compared to laminae) are available in carnivorous plants with ‘active’ trapping mechanisms (D. muscipula; Utricularia, bladder traps)36,37,38. Our results show that, N. khasiana laminae have significantly higher photosynthetic capacity compared to their pitchers whereas respiration rates are comparatively high in pitchers. Similarly, maximum quantum yield of PSII (Fv/Fm) in N. khasiana laminae is high compared to their pitchers. These parameters are matching with similar previous measurements in other Nepenthes species39. Unlike most plant leaf structures, high growth rate and unique physiological functions (prey attraction, capture, digestion, absorption of nutrients) of Nepenthes pitchers demand more energy, prompting higher respiration rates in the trap tissues, resulting in the release of more of CO2. Carnivorous plants follow the C3 photosynthetic pathway, and high CO2 levels are also known to enhance respiration rates in C3 plants40. Thus, we demonstrate respiration of pitcher tissues as the factor contributing to the high CO2 within the ‘closed cavities’ of Nepenthes traps.

Nepenthes tendrils and pitchers grow at a faster rate from their leaf terminals. ‘Rapid elongation’ of growing Nepenthes pitchers and their limited growth after opening of the lid sealing were previously observed by other authors1. Owen and Lennon, 1999 found a uniform growth rate of 0.0147 ± 0.0001 cm per h (0.35 cm per day) for N. alata pitchers, from initiation to the point of lid opening1. A small incision on defined N. khasiana pitchers (initial length, 6–8 cm) released the high CO2 within them, and these pitchers continued growth at a diminished rate compared to control pitchers (Fig. 4). In control (uncut) pitchers, the balancing of CO2 levels (with atmosphere) occurs only on lid opening. Our data indicate that, as in other CO2-enrichment studies, elevated (entrapped) CO2 within acts as a growth promoter of Nepenthes prey traps. Recent studies revealed key data/facts on comparative anatomy41 and construction costs42 of leaves/pitchers of Nepenthes species, leaf development in S. purpurea 43 and the influence of CO2 on leaf phenology in plants44. More investigations, in the light of the discovery of CO2 within, could possibly unravel similar growth patterns (tissue specific changes in cell division)43 and faster growth rates in Nepenthes pitchers. Carbon contents of N. khasiana leaves are comparable to those of non-carnivorous plants3, 42, but, both C and N contents are comparatively low in the pitchers42. As in other Nepenthes species3, 45, 46, the C/N ratio of N. khasiana pitchers is high, 26.05 (n = 4).

CO2 (high) and CO, CH4 and N2O (ambient) found in Nepenthes pitchers are greenhouse gases. Global CO2 levels are predicted to go up to 800 ppm by 2100 and further onto even higher levels14. Nepenthes prey traps with elevated CO2 contents (3000–5000 ppm) are simulating this futuristic scenario in their ‘closed cavities’ (before trap opening). As in other CO2-enrichment experiments14, high carbohydrate and low protein contents were detected in Nepenthes pitchers3. Carbohydrate accumulation is a major acclimation response to elevated CO2 14. High carbohydrate contents in pitchers, transformed into nectar by nectaries (Figs S3 and S4), act as a major ‘lure’ in prey capture. Chlorophyll content is generally low in pitchers compared to their laminae. In some Nepenthes species, pitchers are red-tinted indicating low chlorophyll contents (Fig. S1). Pitchers in Nepenthes have very low photosynthetic rates compared to their laminae3. Reduction in photosynthetic rates in Nepenthes pitchers is primarily due to factors such as replacement of chlorophyll-containing cells with digestive glands, low nitrogen, chlorophyll contents and low stomatal density3, 14. Photosynthetic Nitrogen Use Efficiency (PNUE) is also significantly low in Nepenthes pitchers compared to their laminae. Recently Pavlovič and Saganová pointed out reduced Rubisco activity in Nepenthes prey traps39, and Rubisco content is known to decrease with elevated CO2. These factors viz., photosynthetic rate, C/N ratio, carbohydrate/protein contents, chlorophyll content and PNUE, of several Nepenthes species were compared between their laminae and pitchers by various groups (N. alata and N. mirabilis 3, N. talangensis 47, 8 Nepenthes species and hybrids42 and 15 carnivorous plants including Nepenthes hybrids46). These parameters of Nepenthes leaves and pitchers were also compared to non-carnivorous plants45, 46.

These trends in Nepenthes pitchers mainly, burst of growth, enhanced carbohydrate levels, declined protein levels, drop in photosynthetic capacity, high respiration rate and evolved stomata, are probable manifestations of the enhanced CO2 atmosphere within them. These evidences also infer Nepenthes pitchers as ideal examples reflecting the effects of an anticipated high CO2 level on Earth’s surface, on the characteristic features of plants. Recently, several groups put forward ‘construction cost or cost/benefit theories’3, 42, 45, 46 on Nepenthes prey traps. Most of these studies estimated the nutritional benefit gained from captured preys above (at least marginally) the cost of constructing traps by leaf modification. Future construction cost estimates need to take into account of the acclimation responses of Nepenthes pitchers due to the ‘so far unknown factor’ of high CO2 content within them.

In conclusion, Nepenthes pitchers are CO2-enriched cavities, and CO2 emission from open pitchers acts as a sensory cue attracting insects towards these traps. Most of the characteristic features of Nepenthes pitchers are influenced by the high content of CO2 entrapped within them. This study also hypothesizes Nepenthes pitchers as natural model systems mimicking an anticipated elevated CO2 scenario on Earth.

Methods

Nepenthes pitchers, gas sampling

N. khasiana mature, unopened pitchers (Fig. S1a) were collected from three established populations (08°45′ 00.05″N77°01′45.35″E, altitude 110 m; 08°45′00.04″N, 77°01′41.09″E, altitude 112 m; 08°44′59.74″N, 77°01′40.31″E, altitude 112 m) in Jawaharlal Nehru Tropical Botanic Garden and Research Institute (JNTBGRI) garden sites and the gas compositions inside them were analyzed by gas chromatography (GC-FID/ECD/TCD). Gas compositions inside mature, unopened pitchers of various Nepenthes hybrids (Fig. S1b–g) grown in a greenhouse (08°45′14.59″N, 77°01′31.37″E, altitude 106 m) at JNTBGRI campus were also tested. Gas samples from inside the opened N. khasiana pitchers (from inside, below the peristome) were collected using syringes (Dispovan, Hindustan Syringes and Medical Devices Ltd., Faridabad, India) with a three way stop cock (IGNA, Ignisol Mediplas-Corp, Mumbai, India) and subjected to gas chromatographic analysis. Air samples from JNTBGRI campus were also analyzed.

Lids of mature (about to open, red colour appears at the peristome portion) N. khasiana pitchers in the field were sealed with super glue (to prevent lid opening). Then a small ‘cut’ (average 5.4 × 5.7 mm) was made on the top half (above liquid zone) of the pitcher (for gas release). After 24 h, the cut portion was sealed with parafilm/super glue. After 2 days of sealing, pitchers were collected and subjected to gas analysis. In another set of experiments, lids of opened N. khasiana pitchers (opened a day before) were sealed back with super glue. After 2 days of sealing, these pitchers were collected and their gas compositions were analyzed.


Gas analysis by GC-FID/ECD/TCD

N. khasiana/Nepenthes hybrid unopened pitchers were opened underwater and the gases inside pitchers were collected by the displacement of water. This is to avoid possible mixing with air and dilution of the contents of the pitchers, when opened in air. The gases from the pitchers were transferred to syringes and analyzed through gas chromatography. A Clarus 580 gas chromatograph (Perkin Elmer, Waltham, USA) equipped with a Flame Ionization Detector (FID) and an Electron Capture Detector (ECD) was used. FID had a Methanator for converting CO and CO2 to methane. ECD measured nitrous oxide in the sample. A gas sampling valve with 100 µl sampling loop was used for injecting the sample to the column. Isothermal separation was achieved at 35 °C in an Elite-PLOT Q column (30 m × 0.53 mm) with nitrogen carrier gas. Another NUCON 5765 gas chromatograph (Aimil, New Delhi, India) with a Thermal Conductivity Detector (TCD) and packed column (PORAPAK Q, 80/100 mesh, 5 m long) with nitrogen as carrier gas was used for the measurement of oxygen in the samples. FID, Methanator and ECD were calibrated with the standard gas mixture containing CH4, CO2, CO and N2O in nitrogen gas.


Head space GC/MS/MS of N. khasiana pitcher fluids

N. khasiana pitcher fluids (3 mL each) and 20 mL standard CO2 (carbon dioxide-N5.0, certified concentration 5.49%, nitrogen-N-5.0 balance, Chemtron Science Laboratories, Mumbai, India) bubbled into 3 mL distilled water were transferred to the head space unit (separately) and analyzed by GC/MS/MS. Injection mode: GC head space (Combi Pal, CTC Analytics, Switzerland), syringe temperature 50 °C, sample agitator temperature 60 °C, incubation time 5 min. GC: CP-3800 (Varian, CA, USA), VF-5 (5% phenyl 95% dimethyl polysiloxane, non-polar, 30 m × 0.25 mm i.d., 0.25 μm film thickness) capillary column, column temperature programme isothermal 60 °C for 20 min, flow rate 0.5 mL min−1, MS: Saturn 2200 GC/MS/MS (Varian, CA, USA), mass range 20–60 m/z.


Partial pressures of CO2, O2 in N. khasiana pitcher fluids

Partial pressures of CO2 and O2 in N. khasiana (mature, unopened and opened, prey captured) pitcher fluids were determined using a calibrated ABL800 Basic Gas Analyzer (Radiometer, Copenhagen, Denmark) (Fig. 2).


SEM of N. khasiana roots, leaves, tendrils and pitchers

SEM analyses of N. khasiana abaxial/adaxial sides of leaves, inner/outer sides of pitchers, lids, tendril and roots were carried out on a S-2400 Scanning Electron Microscope (Hitachi, Tokyo, Japan) (Figs 3, S2S4, S8S13). N. khasiana samples were fixed with 3% gluteraldehyde in phosphate buffer and kept overnight. Samples were then dehydrated sequentially with 30%, 50%, 70% ethanol (15 min each, two changes) and 90%, 100% ethanol (30 min each, two changes). These dehydrated samples were subjected to critical point drying, coated with gold and viewed on the SEM.


DART-MS of N. khasiana pitcher fluids

Pitcher fluids (yellow coloured) from prey captured N. khasiana pitchers (Fig. S5), chitin induced5 (Fig. S6) and uninduced (colourless on opening, before prey capture) pitchers (Fig. S7) were collected, lyophilized and analyzed on an AccuTOF JMS-T100LC Mass Spectrometer having a DART (JEOL, MA, USA). Samples were analyzed directly in front of the DART source. Dry He was used at a flow rate of 4 L min−1 for ionization at 350 °C. Orifice 1 was set at 28 V, spectra were collected, and the data from 6–8 scans were averaged.


Nepenthes pitcher weight measurements

N. khasiana and Nepenthes hybrid (mature, unopened) pitchers were collected and their fresh weights were recorded. Then, pitchers were cut open just above the pitcher fluid level (to release the entrapped gas) and the entire pitcher contents were (very) quickly re-weighed (Table S1).


Field studies

N. khasiana pitchers were covered (netted) with colourless nets to prevent ants and insects entering on lid opening. Netting was done a week before opening on near mature pitchers. Three days after opening pitcher fluids were collected, lyophilized and analyzed.

CO2-enriched air (1% CO2 in air; Bhuruka Gases Ltd., Bangalore, India) was passed into just opened N. khasiana pitchers in the field through a small cut made above the fluid level by inserting a long, colourless tubing (inner diameter 2 mm; average flow 25.72 mL/min), and prey (aerial) capture was monitored for 12 days. Similarly, air at the same flow rate was streamed through control pitchers. On the 6th day, gas samples from inside test/control pitchers (just below the peristomes) were collected in syringes and analyzed by gas chromatography (n = 6, each). On the 12th day after lid opening, the entire contents of test/control N. khasiana pitchers (n = 6, each) were (separately) transferred to petri dishes (Fig. 1), and captured aerial preys (in each dish) were carefully counted. Similarly, prey (aerial) capture rates in normal (unmodified) pitchers (with no CO2/air streaming) in 12 days were also counted. In all three experiments, ants (dead) crawled into these pitchers from the ground were not considered (counted).

Tendrils of live N. khasiana plants were cut just below the pitchers and their cross sections were inserted into inverted syringes partially filled with water (for 6 days) in the field. On repeated experiments, no gas bubbling or any other changes in the water were observed.


N. khasiana pitcher growth measurements

N. khasiana pitchers from the three populations in JNTBGRI garden sites with an initial growth of 6 to 8 cm were marked, their initial pitcher lengths were noted and small cuts (average 5.4 × 5.7 mm, to release the gas inside pitchers) were made above the fluid level. These test pitchers were constantly monitored, pitcher lengths on the day of lid opening and the number of days required till lid opening (from an initial stage of 6 to 8 cm) were noted. Similar measurements were also made on control N. khasiana pitchers (with no cuts) (Fig. 4 and Table S2).


C, N contents in N. khasiana leaves, pitchers

N. khasiana leaves and pitchers were dried at 60 °C for 72 h and their carbon and nitrogen contents were analyzed on a Vario EL III CHN Analyzer (Elementar Analysensysteme GmbH, Hanau, Germany).


Chlorophyll-A fluorescence, photosynthesis (An) and dark respiration (Rd) of N. khasiana laminae and pitchers

Chlorophyll-a fluorescence kinetics, A N and R D of N. khasiana laminae and pitchers were measured using a LI-COR 6400 XT portable infrared analyzer (LI-COR, Lincoln, NE, USA), equipped with a leaf chamber fluorometer. Laminae and pitchers from four N. khasiana plants in the field were subjected to these measurements. Fully grown N. khasiana laminae and healthy, prey captured pitchers (pitcher walls and lids were directly placed into the cuvette, independently) were taken for measurements. A constant PAR (photosynthetically active radiation) of 800 μmol m−2 s−1 of red (90%) and blue (10%) light was chosen as actinic light intensity and the measurement of chlorophyll fluorescence and P N were at ambient CO2 level, temperature (33 ± 1 °C), RH (relative air humidity) ~80% and air flow rate of 300 μmol s−1. R D was measured under similar conditions, except that the plant samples were under dark conditions. The laminae and traps were kept in the chamber for 5–10 min, until steady state of CO2 concentrations were reached. Vapor pressure deficit in the sample cell ranged between 0.7 and 1.3 kPa. Minimal fluorescence (F 0) was measured for overnight dark adapted plant samples whereas maximal fluorescence (F m) was recorded at a PAR of 8000 μmol m−2 s−1 (saturating flash). Maximal quantum yield of PSII was calculated as F v/F m = (F m − F 0)/F m.


Statistical analysis

Prey capture rates (Fig. 1), partial pressure measurements (Fig. 2), pitcher size/weight measurements (Table S1) and growth parameters of cut/uncut pitchers (Fig. 4 and Table S2) are expressed as mean ± s.d. Statistical comparisons were done using student’s t-test (Figs 1 and 2). Values of p < 0.05 were considered as statistically significant.


Data availability

All data generated or analyzed during this study are included in this published article (and its Supplementary Information file).

Additional information

Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  1. 1.

    Owen, T. P. Jr. & Lennon, K. A. Structure and development of the pitchers from the carnivorous plant Nepenthes alata (Nepenthaceae). Am. J. Bot. 86, 1382–1390 (1999).

  2. 2.

    Bohn, H. F. & Federle, W. Insect aquaplaning: Nepenthes pitcher plants capture prey with the peristome, a fully wettable water-lubricated anisotropic surface. Proc. Natl. Acad. Sci. USA 101, 14138–14143 (2004).

  3. 3.

    Pavlovič, A., Masarovičová, E. & Hudák, J. Carnivorous syndrome in Asian pitcher plants of the genus Nepenthes. Ann. Bot. 100, 527–536 (2007).

  4. 4.

    Adlassnig, W., Peroutka, M. & Lendl, T. Traps of carnivorous pitcher plants as a habitat: composition of the fluid, biodiversity and mutualistic activities. Ann. Bot. 107, 181–194 (2011).

  5. 5.

    Raj, G., Kurup, R., Hussain, A. A. & Sabulal, B. Distribution of naphthoquinones, plumbagin, droserone, and 5-O-methyl droserone in chitin-induced and uninduced Nepenthes khasiana: molecular events in prey capture. J. Exp. Bot. 62, 5429–5436 (2011).

  6. 6.

    Kurup, R. et al. Fluorescent prey traps in carnivorous plants. Plant. Biol. (Stuttg.) 15, 611–615 (2013).

  7. 7.

    Chen, H. et al. Continuous directional water transport on the peristome surface of Nepenthes alata. Nature 532, 85–89 (2016).

  8. 8.

    Clarke, C. M. et al. Tree shrew lavatories: a novel nitrogen sequestration strategy in a tropical pitcher plant. Biol. Lett. 5, 632–635 (2009).

  9. 9.

    Bauer, U. & Federle, W. The insect-trapping rim of Nepenthes pitchers: surface structure and function. Plant Signal. Behav. 4, 1019–1023 (2009).

  10. 10.

    Robinson, A. S. et al. A spectacular new species of Nepenthes L. (Nepenthaceae) pitcher plant from central Palawan, Philippines. Bot. J. Linn. Soc. 159, 195–202 (2009).

  11. 11.

    Bauer, U., Willmes, C. & Federle, W. Effect of pitcher age on trapping efficiency and natural prey capture in carnivorous Nepenthes rafflesiana plants. Ann. Bot. 103, 1219–1226 (2009).

  12. 12.

    Adams, R. M. II & Smith, G. W. An S.E.M. survey of the five carnivorous pitcher plant genera. Am. J. Bot. 64, 265–272 (1977).

  13. 13.

    Moran, J. A., Booth, W. E. & Charle, J. K. Aspects of pitcher morphology and spectral characteristics of six Bornean Nepenthes pitcher plant species: Implications for prey capture. Ann. Bot. 83, 521–528 (1999).

  14. 14.

    Sage, R. F. How terrestrial organisms sense, signal, and respond to carbon dioxide. Integr. Comp. Biol. 42, 469–480 (2002).

  15. 15.

    Jones, W. D., Cayirlioglu, P., Kadow, I. G. & Vosshall, L. B. Two chemosensory receptors together mediate carbon dioxide detection in Drosophila. Nature 445, 86–90 (2007).

  16. 16.

    Cooperband, M. F. & Cardé, R. T. Comparison of plume structures of carbon dioxide emitted from different mosquito traps. Med. Vet. Entomol. 20, 1–10 (2006).

  17. 17.

    Buch, F. et al. Secreted pitfall-trap fluid of carnivorous Nepenthes plants is unsuitable for microbial growth. Ann. Bot. 111, 375–383 (2013).

  18. 18.

    Kneitel, J. M. & Miller, T. E. Resource and top-predator regulation in the pitcher plant (Sarracenia purpurea) inquiline community. Ecology 83, 680–688 (2002).

  19. 19.

    Wakefield, A. E., Gotelli, N. J., Wittman, S. E. & Ellison, A. M. Prey addition alters nutrient stoichiometry of the carnivorous plant Sarracenia purpurea. Ecology 86, 1737–1743 (2005).

  20. 20.

    Adamec, L. Oxygen concentrations inside the traps of the carnivorous plants Utricularia and Genlisea (Lentibulariaceae). Ann. Bot. 100, 849–856 (2007).

  21. 21.

    Tresguerres, M., Buck, J. & Levin, L. R. Physiological carbon dioxide, bicarbonate, and pH sensing. Pflugers. Arch. 460, 953–964 (2010).

  22. 22.

    Moran, J. A., Hawkins, B. J., Gowen, B. E. & Robbins, S. L. Ion fluxes across the pitcher walls of three Bornean Nepenthes pitcher plant species: flux rates and gland distribution patterns reflect nitrogen sequestration strategies. J. Exp. Bot. 61, 1365–1374 (2010).

  23. 23.

    Moran, J. A. & Clarke, C. M. The carnivorous syndrome in Nepenthes pitcher plants. Current state of knowledge and potential future directions. Plant Signal. Behav. 5, 644–648 (2010).

  24. 24.

    An, C. I., Fukusaki, E. I. & Kobayashi, A. Plasma-membrane H+-ATPases are expressed in pitchers of the carnivorous plant Nepenthes alata Blanco. Planta 212, 547–555 (2001).

  25. 25.

    Hetherington, A. M. & Woodward, F. I. The role of stomata in sensing and driving environmental change. Nature 424, 901–908 (2003).

  26. 26.

    Scholz, I. et al. Slippery surfaces of pitcher plants: Nepenthes wax crystals minimize insect attachment via microscopic surface roughness. J. Exp. Biol. 213, 1115–1125 (2010).

  27. 27.

    Wang, L., Zhou, Q., Zheng, Y. & Xu, S. Composite structure and properties of the pitcher surface of the carnivorous plant Nepenthes and its influence on the insect attachment system. Prog. Nat. Sci. 19, 1657–1664 (2009).

  28. 28.

    Riedel, M., Eichner, A., Meimberg, H. & Jetter, R. Chemical composition of epicuticular wax crystals on the slippery zone in pitchers of five Nepenthes species and hybrids. Planta 225, 1517–1534 (2007).

  29. 29.

    Holroyd, G. H., Hetherington, A. M. & Gray, J. E. A role for the cuticular waxes in the environmental control of stomatal development. New Phytol. 153, 433–439 (2002).

  30. 30.

    Moran, J. A., Clarke, C. & Gowen, B. E. The use of light in prey capture by the tropical pitcher plant Nepenthes aristolochioides. Plant Signal. Behav. 7, 957–960 (2012).

  31. 31.

    Voigt, D., Gorb, E. & Gorb, S. Plant surface-bug interactions: Dicyphus errans stalking along trichomes. Arthropod Plant Interact. 1, 221–243 (2007).

  32. 32.

    Bauer, U., Sharmann, M., Skepper, J. & Federle, W. ‘Insect aquaplaning’ on a superhydrophilic hairy surface: how Heliamphora nutans Benth. pitcher plants capture prey. Proc. R. Soc. B. 280, 20122569 (2012).

  33. 33.

    Gaume, L., Bazile, V., Huguin, M. & Bonhomme, V. Different pitcher shapes and trapping syndromes explain resource partitioning in Nepenthes species. Ecol. Evol. 6, 1378–1392 (2016).

  34. 34.

    Król, E. et al. Quite a few reasons for calling carnivores ‘the most wonderful plants in the world’. Ann. Bot. 109, 47–64 (2012).

  35. 35.

    Robinson, E. A., Ryan, G. D. & Newman, J. A. A meta-analytical review of the effects of elevated CO2 on plant-arthropod interactions highlights the importance of interacting environmental and biological variables. New Phytol. 194, 321–336 (2012).

  36. 36.

    Ellison, A. M. & Adamec, L. Ecophysiological traits of terrestrial and aquatic carnivorous plants: are the costs and benefits the same? Oikos 120, 1721–1731 (2011).

  37. 37.

    Pavlovič, A., Demko, V. & Hudák, J. Trap closure and prey retention in Venus flytrap (Dionaea muscipula) temporarily reduces photosynthesis and stimulates respiration. Ann. Bot. 105, 37–44 (2010).

  38. 38.

    Adamec, L. Respiration and photosynthesis of bladders and leaves of aquatic Utricularia species. Plant Biol. (Stuttg.) 8, 765–769 (2006).

  39. 39.

    Pavlovič, A. & Saganová, M. A novel insight into the cost-benefit model for the evolution of botanical carnivory. Ann. Bot. 115, 1075–1092 (2015).

  40. 40.

    Leakey, A. D. B. et al. Genomic basis for stimulated respiration by plants growing under elevated carbon dioxide. Proc. Natl. Acad. Sci. USA 106, 3597–3602 (2009).

  41. 41.

    Osunkoya, O. O. & Muntassir, N. A. Comparative anatomy of the assimilatory organs of Nepenthes species. Aust. J. Bot. 65, 67–79 (2017).

  42. 42.

    Osunkoya, O. O., Daud, S. D., Di-Giusto, B., Wimmer, F. L. & Holige, T. M. Construction costs and physico-chemical properties of the assimilatory organs of Nepenthes species in Northern Borneo. Ann. Bot. 99, 895–906 (2007).

  43. 43.

    Fukushima, K. et al. Oriented cell division shapes carnivorous pitcher leaves of Sarracenia purpurea. Nat. Commun. 6, 6450 (2015).

  44. 44.

    Buitenwerf, R., Rose, L. & Higgins, S. I. Three decades of multi-dimensional change in global leaf phenology. Nat. Clim. Change 5, 364–368 (2015).

  45. 45.

    Ellison, A. M. Nutrient limitation and stoichiometry of carnivorous plants. Plant Biol. (Stuttg.) 8, 740–747 (2006).

  46. 46.

    Karagatzides, J. D. & Ellison, A. M. Construction costs, payback times, and the leaf economics of carnivorous plants. Am. J. Bot. 96, 1612–1619 (2009).

  47. 47.

    Pavlovič, A., Singerová, L., Demko, V. & Hudák, J. Feeding enhances photosynthetic efficiency in the carnivorous pitcher plant Nepenthes talangensis. Ann. Bot. 104, 307–314 (2009).

Download references

Acknowledgements

Government of Kerala, India financed this work through Plan Project JNTBGRI/P-132 [SB] and Kerala State Council for Science Technology and Environment, grant no. 010/E&E/2011/CSTE [EJZ]. We acknowledge Dr. Bejoy Mathew, Principal Scientist, Plant Genetic Resources Division, JNTBGRI for providing us Nepenthes hybrid samples. We thank Ms. Deepa Nair Krishnan Nair, NCESS, Thiruvananthapuram, India for helping us in the gas analysis and Dr. Velumani Ravi and Dr. Saravanan Raju, Plant Physiology, Crop Production Division, Central Tuber Crops Research Institute (CTCRI), Thiruvananthapuram, India for their inputs in photosynthesis/respiration measurements. We also acknowledge Mr. G. Balaji, Senior Research Fellow in our group for his support in the field studies.

Author information

Affiliations

  1. Phytochemistry and Phytopharmacology Division, Jawaharlal Nehru Tropical Botanic Garden and Research Institute, Pacha-Palode, Thiruvananthapuram, 695 562, Kerala, India

    • Sabulal Baby
    •  & Anil John Johnson
  2. Atmospheric Sciences Division, National Centre for Earth Science Studies, Post Box No. 7250, Akkulam, Thiruvananthapuram, 695 011, Kerala, India

    • Elavinamannil Jacob Zachariah
  3. Garden Management Division, Jawaharlal Nehru Tropical Botanic Garden and Research Institute, Pacha-Palode, Thiruvananthapuram, 695 562, Kerala, India

    • Abdul Azeez Hussain

Authors

  1. Search for Sabulal Baby in:

  2. Search for Anil John Johnson in:

  3. Search for Elavinamannil Jacob Zachariah in:

  4. Search for Abdul Azeez Hussain in:

Contributions

S.B. developed the concept and wrote the manuscript with inputs from A.J.J. and E.J.Z.; A.J.J., S.B. carried out chemical, S.E.M., field and other studies; A.A.H. provided N. khasiana samples and field support; E.J.Z. carried out gas analysis with A.J.J.

Competing Interests

The authors declare that they have no competing interests.

Corresponding author

Correspondence to Sabulal Baby.

Electronic supplementary material

About this article

Publication history

Received

Accepted

Published

DOI

https://doi.org/10.1038/s41598-017-11414-7

Further reading

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.