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Factors Inducing Successful Anhydrobiosis in the African Chironomid Polypedilum vanderplanki: Significance of the Larval Tubular Nest1
1 National Institute of Agrobiological Sciences, 1-2 Ohwashi, Tsukuba, Ibaraki 305-8634, Japan
2 Saga University, Faculty of Medicine, 5-1-1 Nabeshima, Saga 849-8501, Japan
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SYNOPSIS |
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The African chironomid Polypedilum vanderplanki exhibits anhydrobiosis, i.e., the larvae can survive complete desiccation. Recovery rate and trehalose content were investigated in larvae desiccated slowly or at a rate more than 3 times faster. Upon slow desiccation (evaporation rate 0.22 ml day–1) larvae synthesized 38 µg trehalose/individual before complete desiccation, and all of them recovered after rehydration, whereas larvae that were dehydrated quickly (evaporation rate 0.75 ml day–1) accumulated only 6.8 µg trehalose/individual and none of them revived after rehydration. In the pools that are their natural habitat P. vanderplanki larvae make tubes by incorporating detritus or soil with their sticky saliva. This tubular structure is a physical barrier not only to protect the larva from natural enemies but also induces successful anhydrobiosis by reducing the dehydration rate. When larvae were dehydrated with 100 µl distilled water (DW) in soil tubes, they accumulated 37 µg trehalose/individual and more than half of them could revive after rehydration, whereas larvae without tubes accumulated lower level of trehalose and none recovered after rehydration.
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INTRODUCTION |
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Organisms have a variety of strategies to survive severe dry environments. Anhydrobiosis is the most extreme biological state resistant to drought, in which organisms are completely dehydrated. Metabolic activity is undetectable but can resume after rehydration without ill effect (Keilin, 1959
; Hochachka and Somero, 1984; Crowe et al., 1992). Anhydrobiosis occurs in a wide variety of invertebrates including primitive cysts of crustaceans, insects, nematodes, tardigrades and rotifers (Sømme, 1995; Clegg, 2001). A common characteristic found in anhydrobiotes is that they accumulate large amounts of trehalose upon desiccation (Clegg, 1965; Madin and Crowe, 1975; Crowe, 2002).
When dehydrated too fast, individuals often fail to go into anhydrobiosis (Womersley, 1987; Sømme, 1995; Wharton, 2002). In nature dormant or anhydrobiotic individuals live within or on protective materials such as soil, litter, moss and host plant tissues, and these are assumed to provide them with a physical barrier that slows down evaporation and water loss during desiccation (Sømme, 1995; Danks, 2000). The precise mechanism by which slow dehydration enhances survival is still a matter of conjecture.
The larva of the African chironomid Polypedilum vanderplanki living in a temporary rock pool exhibits anhydrobiosis (Hinton, 1951, 1960a, b). The larvae make tubular shelters in the pools by incorporating detritus or soil with their saliva. The tubes of chironomids are thought to have three functions. First, the tubes allow larvae to obtain oxygen and food efficiently, because the larvae produce water currents in their tubes by undulating their bodies, which serve to trap food in the water at the open mouth of the tubes, feces can also be washed out (Walshe, 1951). Secondly they have a protective role against predacious natural enemies (Macchiusi and Baker, 1992). Because many species of chironomid larvae including P. vanderplanki are red in body color due to hemoglobin in the hemolymph, without their tubes they would be easily visible to natural enemies. Thirdly, the tubes are to be protecting the chironomid larvae also from chemical toxicants (Halpern et al., 2002). In this paper we confirm that slow dehydration enhances survival of P. vanderplanki, and also propose that the tubes play a fourth role by contributing to a decrease in the desiccation rate as required for successful anhydrobiosis in P. vanderplanki.
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MATERIALS AND METHODS |
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Insects rearing
Anhydrobiotic larvae of P. vanderplanki were collected from rock pools in Nigeria in 2000. They were transferred to the laboratory, and put into a plastic container (200 x 300 x 100 mm) containing water (depth, 20 to 30 mm) over autoclaved soil (depth, 20 to 30 mm). The rearing water was always aerated. The container was covered with a nylon-mesh cage (200 x 300 x 250 to 300 mm). The species was reared for successive generations under controlled light (13 hr light : 11 hr dark) and temperature (27°C). Final-instar larvae of a similar body weight (about 1 mg) were used in all the experiments.
Desiccating procedure with a filter paper
Larvae were placed on a filter paper (dia., 50 mm) absorbing various amount of DW in a glass Petri dish (dia., 65 mm; height, 20 mm). The dishes containing larvae were transferred to a desiccation box (250 x 300 x 250 mm) with 1 kg silica gel at less than 5% relative humidity (RH) at room temperature (24– 26°C). When these dishes had a glass top, water evaporated slowly (0.22 ml day–1) over different periods of time depending on the initial amount of distilled water (DW). Water in Petri dishes without the glass top and transferred to a desiccation box evaporates at a quick rate (0.75 ml day–1).
Desiccating procedure with soil
Each of 6 to 24 larvae was placed in a glass Petri dish (dia., 65 mm; height, 20 mm) with different amounts of autoclaved soil and distilled water (soil : DW = 36 g : 4 ml, 0.9 g : 1 ml or 0.45 g : 0.5 ml). A half of the dishes containing larvae were desiccated after pre-incubation for 2 days: the glass top was sealed with vinyl tape to prevent evaporation for 2 days. During the pre-incubation, larvae construct their tubular nest by soil. After the pre-incubation, the glass tops of the dishes were removed and transferred to a desiccation box. Another half of the dishes was directly desiccated in the desiccation box without the pre-incubation treatment.
Desiccating procedure with or without soil tubes
Forty to sixty of larvae were incubated for 2 days with soil and DW in a glass Petri dish, and these larvae in soil tubes were picked up from the dishes. The volume of DW absorbed in a soil tube was ca. 6 µl on average. Each of three larvae with soil tubes was desiccated with 88 or 182 µl DW on a filter paper in a glass Petri dish (total water volume except for larval water were 100 and 200 µl, respectively).
Recovery check
Larvae desiccated in various ways were rehydrated by immersion in DW followed by observation every 0.5 hr or 1 hr during 6 hr after rehydration. Larvae were judged as surviving if they could repeatedly contract their abdominal muscles.
Sugar and polyol measurements
Sample larvae were homogenized individually with 0.1 mg of sorbitol as an internal standard in 0.4 ml of 90% ethanol. The supernatant after membrane filtration (0.45 µm) was dried under vacuum and the dried residue was dissolved in 0.5 ml of MilliQ water (Millipore). The sample was analyzed on a Shimadzu HPLC system (LC-10A system, Shimadzu, Japan) equipped with a guard column (Shim-pack SCR-C, 4.0 x 50 mm, Shimadzu, Japan) connected to an analytical column (Shim-pack SCR-101C, 7.9 x 300 mm, Shimadzu, Japan) and a reflective index detector (RID-6A, Shimadzu, Japan), as described by Watanabe et al. (2002).
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RESULTS |
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Recovery and trehalose content in larvae desiccated with a wet filter paper
Changes of trehalose content in larvae desiccated at different evaporation rates over 48 hr are shown in Figure 1. Upon slow desiccation (DW evaporating at 0.22 ml day–1 in a Petri dish with its glass top) larvae with a filter paper holding 0.44 ml DW started synthesizing trehalose 12 hr after transfer to the desiccation box, and accumulated 38 µg trehalose/individual before complete dehydration. All these slowly desiccated larvae recovered within 1 hr after rehydration. In contrast, during quick desiccation (0.75 ml day–1 in a dish without the glass top) with moisturized filter paper holding 1.5 ml DW, and in spite of the fact that larvae desiccated over a period of 48 hr, larvae accumulated only about 6.8 µg/individual and none of these larvae revived after rehydration.
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Recovery and trehalose content of larvae desiccated in soil
In nature P. vanderplanki larvae make tubular shelters in the pools by incorporating detritus or soil with their saliva (Fig. 2). When P. vanderplanki larvae were pre-incubated with 4 ml DW and enough soil (3.6 mg) in a glass Petri dish in the laboratory, all of them constructed soil tubes within 2 days. Almost all of those larvae desiccated in soil tubes recovered after rehydration because they had accumulated enough trehalose, 61.2 µg/individual (Table 1). When larvae were desiccated in a glass petri dish with 4 ml DW but without the 2 days pre-incubation period, although 63.4% larvae constructed soil tubes and accumulated 58.5 µg/individual of trehalose, only 38.5% attained successful anhydrobiosis. Both recovery rate and trehalose content were significantly greater in larvae desiccated in soil tubes (recovery:
2 test, P < 0.05; trehalose: Mann-Whitney U test, P < 0.05). This tendency was more obvious when the larvae were desiccated with lower amount of soil and DW: 92.5% of larvae that made tubes during the pre-incubation period could revive after rehydration, accumulating trehalose of 37.2 µg/individual, whereas larvae without tubes accumulated only 1.9 µg of trehalose/individual and none recovered after rehydration.
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Trehalose content in larvae desiccated with or without soil tubes
To confirm a significant role of the larval tubular nests in inducing successful anhydrobiosis, trehalose content and recovery in desiccated larvae with and without soil tubes were compared (Table 2). Larvae in soil tubes were desiccated with 182 µl DW (total water volume is 200 µl) accumulated 38 µg trehalose / individual before complete desiccation and 81.3% of them recovered after rehydration. When larvae without the soil tubes were desiccated with 200 µl DW, they accumulated trehalose at significantly lower level of 23 µg (Mann-Whitney U test, P < 0.05) and only 43.3% of them recovered after rehydration. When total water volume in the dish was reduced to 100 µl in the soil tubes accumulated 37 µg of trehalose and 63.3% recovered after rehydration, whereas those without soil tubes synthesized trehalose at lower level (17.3 µg) and none of them recovered after rehydration.
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DISCUSSION |
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Survival increases with slower rates of dehydration in many anhydrobiotes (Womersley, 1987
; Womersley and Ching, 1989; Sømme, 1995; Wharton, 2002). In tardigrades, tun formation is essential for successful induction of anhydrobiosis. Water loss declines rapidly just after tun formation (Wright, 1989; Wright et al., 1992; Sømme, 1995). If tardigrades are dried under anoxia, they do not form a tun but shrink into a flat formless shape and do not survive (Crowe, 1972). Similarly, if the animals are quickly dried at low RHs, they die because water loss is too fast and they are not able to form tuns. The plant parasitic nematode Ditylenchus dipsaci forms aggregations and also decreases cuticular permeability to avoid water loss during desiccation (Wharton, 1996). The worms on the periphery are killed while those in the center survive due to reduced rates of water loss (Ellenby, 1968). Instead of such behavioral adaptations to reduce water loss, some anhydrobiotes live within or on protective materials such as soil, litter, moss and host plant tissues, and these are assumed to provide them with a physical barrier that slows down evaporation and water loss during desiccation. P. vanderplanki larvae could belong to this group; physical barrier of tubes made of saliva and detritus decrease the rate of larval water loss. Here, the laboratory experiments indicated that the tubes play an important role in facilitating successful anhydrobiosis. We emphasize that for successful induction of anhydrobiosis the rate of evaporation is more important than the duration of desiccation treatment, because larvae desiccated at slower evaporation rates accumulated greater amounts of trehalose and had higher recovery rates than those desiccated more quickly, even over the same 48 hr desiccation period (Fig. 1). When P. vanderplanki larvae were desiccated slowly (DW evaporating at 0.22 ml day–1) larvae started synthesizing trehalose approximately 12 hr after transfer to the desiccation box.
We hypothesize that the process of anhydrobiosis proceeds as shown in Figure 3. The starting point for trehalose synthesis by slow desiccation larvae is designated as point "A" from the data in Figure 1, thus the larvae synthesized 38 µg tehalose/individual for 36 hr before complete dehydration. The duration of trehalose synthesis is shown as ‘a’ in the figure. We postulate a threshold of water soaked with a filter paper in a Petri dish (the horizontal line ‘C’ in Fig. 3). We found that this water threshold was the same when larvae were desiccated at the same evaporation rate (0.22 ml day–1) over 7 days (Fig. 1B in Watanabe et al., 2003). Based on this water threshold in a Petri dish, we could estimate roughly when trehalose synthesis takes place under quick evaporation conditions, as "B" in Figure 3, suggesting that quickly desiccating larvae have less than 10 hr (duration of trehalose synthesis = "b" in Fig. 3) to synthesize and accumulate trehalose before complete desiccation. Indeed, quickly desiccated larvae accumulated only 6.8 µg trehalose/ individual (Fig. 1). We conclude that the time available from the triggering of trehalose synthesis to complete desiccation is an important factor, and slow evaporation facilitates anhydrobiosis by prolonging the time available for trehalose synthesis.
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A single larval tubular nest can absorb only 6 µl of DW on average. Six µl DW in a Petri dish evaporate within 40 min even under slow desiccation conditions. According to recovery rate results (Table 2), 6 µl DW absorbed by a larval tube must have maintained its moisture longer than 11 hr, so that 6 µl DW absorbed by the tube kept moisture equivalent to 100 µl DW without the tube. The recovery rate was about 20 % higher in larvae desiccated with 100 µl DW in the soil tubes than in those desiccated with 200 µl DW without soil tubes. Our interpretation is that the structure of the tubes, including soil and larval saliva might retain even a small amount of moisture for a long time, and this micro-climatic moisture is enough to allow the dehydrating larvae to activate the biochemical machinery necessary to achieve successful anhydrobiosis.
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ACKNOWLEDGMENTS |
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This work was supported in part by the Program for Promotion of Basic Research Activities for Innovative Biosciences (PROBRAIN), by the scientific research (young B)(16770058) from Japan Society for the Promotion Science, by the budget for Nuclear Research for the Ministry of Education, Culture, Sports, Science Technology, based on the screening and consulting by the Atomic Energy Commission, and by a grant-in Aid (Bio Design Program) from the Ministry of Agriculture, Forestry and Fisheries of Japan. We thank H. V. Danks for critical reading of the manuscript.
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FOOTNOTES |
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1 From the Symposium Drying Without Dying: the Comparative Mechanisms and Evolution of Desiccation Tolerance in Animals, Microbes, and Plants presented at the Annual Meeting of the Society for Integrative and Comparative Biology, 4–8 January 2005, at San Diego, California.
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