MIAMI UNIVERSITY The Graduate School
Certificate for Approving the Dissertation
We hereby approve the Dissertation
of
Michael A. Elnitsky
Candidate for the Degree:
Doctor of Philosophy
______Director Richard E. Lee, Jr.
______Reader Alan B. Cady
______Reader Jon P. Costanzo
______Reader Kathleen A. Killian
______Graduate School Representative Robert L. Schaefer
ABSTRACT
TOLERANCE AND PHYSIOLOGICAL RESPONSE TO ENVIRONMENTAL STRESS IN ANTARCTIC ARTHROPODS
by Michael A. Elnitsky
The Antarctic Peninsula is characterized by harsh and dynamic environmental conditions. Organisms inhabiting this environment may be challenged by extremes of low temperature, limited water availability, dramatic seasonal fluctuations of light availability and ultraviolet radiation, and high salinity. This dissertation describes three projects examining the tolerance and physiological responses to such environmental stress of two Antarctic arthropods, the midge Belgica antarctica and the collembolan Cryptopygus antarcticus. The first investigation examined the ability of B. antarctica larvae to resist inoculative freezing at subzero temperatures and instead dehydrate as a strategy for winter survival (i.e., cryoprotective dehydration). When cooled to subzero temperatures in the presence of ice, the body fluid melting point was depressed to near equilibrium with the ambient temperature, due to reductions of body water content and the accumulation of several osmolytes, suggesting larvae can undergo cryoprotective dehydration at subzero temperatures. Under more natural conditions, the use of cryoprotective dehydration versus freeze tolerance for winter survival appears to depend upon the moisture content of the surrounding soil. The purpose of the second study was to assess the tolerance and physiological response to desiccation of C. antarcticus under ecologically-relevant conditions. Slow dehydration at high relative humidities characteristic of the austral summer induced the accumulation of several organic osmolytes and increased the tolerance of water loss. A mild drought acclimation further increased the subsequent desiccation tolerance of C. antarcticus. The springtails were also susceptible to water loss at subzero temperatures and likely rely upon such dehydration as a key component for winter survival. As B. antarctica microhabitats may be periodically inundated with seawater, the final investigation examined the osmotic response and tolerance of larvae to hyperosmotic seawater exposure. The larvae displayed an impressive tolerance of the osmotic stress, as ~50% survived a 6-d submergence in pure seawater. Hyperosmotic stress induced the accumulation of organic osmolytes and resulted in a significant positive correlation between the rate of oxygen consumption and larval body water content. Finally, a brief seawater acclimation enhanced the subsequent tolerance of freezing and dehydration, but reduced the tolerance of heat shock.
TOLERANCE AND PHYSIOLOGICAL RESPONSE TO ENVIRONMENTAL STRESS IN ANTARCTIC ARTHROPODS
A DISSERTATION
Submitted to the Faculty of
Miami University in partial
fulfillment of the requirements
for the degree of
Doctor of Philosophy
Department of Zoology
by
Michael A. Elnitsky
Miami University
Oxford, Ohio
2008
TABLE OF CONTENTS
Item Page
Table of contents ii
List of tables iv
List of figures v
Acknowledgements ix
Chapter 1: General introduction 1 References 4
Chapter 2: Cryoprotective dehydration and the resistance to inoculative 6 freezing in the Antarctic midge, Belgica antarctica Summary 7 Introduction 8 Materials and methods 9 Results 12 Discussion 14 References 19 Table 22 Figure legends 23 Figures 24
Chapter 3: Desiccation tolerance and drought acclimation in Antarctic 28 collembolan Cryptopygus antarcticus Summary 29 Introduction 30
ii Materials and methods 31 Results 35 Discussion 37 References 43 Tables 47 Figure legends 49 Figures 50
Chapter 4: Osmoregulation and salinity tolerance in the Antarctic midge, 54 Belgica antarctica: seawater acclimation confers cross tolerance to freezing and dehydration Summary 55 Introduction 56 Materials and methods 57 Results 61 Discussion 63 References 70 Table 74 Figure legends 76 Figures 78
Chapter 5: Concluding remarks 83
iii LIST OF TABLES
Table Page Chapter 2: 2.1 Table 1. Estimated osmotic contribution of initial osmolytes in the 22 hemolymph and osmolytes produced during slow cooling to -3oC in an environment at equilibrium with the vapor pressure of ice. Values are mean ± SEM.
Chapter 3: 3.1 Table 1. The total body water content (N = 25-30) and osmolyte 47 concentrations (N = 6) of Cryptopygus antarcticus during drought acclimation at 4oC and 98.2 or 75.0% RH. Values are mean ± SEM. Within an osmolyte different letters denote significant differences between treatment groups (ANOVA; Bonferroni-Dunn test).
3.2 Table 2. Osmolyte concentrations (N = 6) of Cryptopygus antarcticus 48 during exposure at -3.0oC in an environment at equilibrium with the vapor pressure of ice. Values are mean ± SEM. Within an osmolyte different letters denote significant differences between days of exposure (ANOVA; Bonferroni-Dunn test).
Chapter 4: 4.1 Table 1. Estimated osmotic contribution of initial osmolytes in the 74 hemolymph and osmolytes produced during hyperosmotic seawater exposure. Values are mean ± SEM. Within an osmolyte different letters represent significant differences between days of exposure (ANOVA; Bonferroni-Dunn test).
iv LIST OF FIGURES
Figure Page Chapter 2: 2.1 Fig. 1. Seasonal changes in temperature at a representative larval 24 Belgica antarctica microhabitat site on Torgersen Island, near Palmer Station, Antarctica (64o46’ S, 64o04’ W). Microhabitat temperatures were measured in 2005-2006 using single-channel temperature loggers (HOBO Water Temp Pro, Onset Computer, Pocasset, MA, USA). The dashed line represents the equilibrium freezing point of the body fluids of fully hydrated, control larvae.
2.2 Fig. 2. Changes in (A) body water content (N = 15) and (B) body fluid 25 melting point (N = 6) of larval Belgica antarctica during slow cooling to -3oC in an environment at equilibrium with the vapor pressure of ice. Different letters indicate significant differences between values (ANOVA, Bonferroni-Dunn test, P<0.05). Values are mean ± SEM.
2.3 Fig. 3. Body water content (WC) of individual Belgica antarctica 26 larvae (N = 30) during slow cooling to -3oC in contact with substrates -1 of varying moisture content: 0.80, 1.10, and 1.40 g H2O · g dry soil. Triangles denote WC of individuals at day 0, circles the WC of frost exposed individuals (day 16). Dashes denote the mean WC of individuals at day 0 and 16, separated into ‘high’ (frozen) and ‘low’ (dehydrated) WC groups.
v 2.4 Fig. 4. Percentage of Belgica antarctica larvae frozen, as detected by 27 the maintenance of ‘high’ body water content, during cooling to -3oC in contact with substrates of varying moisture content. Different letters indicate significant differences between values (ANOVA, Bonferroni-Dunn test, P<0.05). Values are mean ± SEM of three groups of 10 individuals.
Chapter 3: 3.1 Fig. 1. (A) Changes in total body water content of Cryptopygus 50 antarcticus during desiccation exposure within various relative humidity (RH) environments at 4oC. Values are the mean ± SEM of 25-30 individuals. (B) Percent survival as a function of total body water content of C. antarcticus during desiccation in various constant relative humidity environments. Values are the mean ± SEM of five groups of 10 individuals.
3.2 Fig. 2. Water loss rate of Cryptopygus antarcticus at 4oC and 0% RH. 51 A linear regression line was fitted to the points (y = -0.208x – 0.00650, R2 = 0.998), where the slope of the regression represents the water loss rate in percent of total body water per hour. Values are mean ± SEM of 25-30 individuals.
3.3 Fig. 3. Survival of Cryptopygus antarcticus desiccated for 5 d at either 52 96.0% RH (A) or 93.0% RH (B) at 4oC. Collembola were previously acclimated at 100% RH (control) or drought acclimated at 98.2 or 75.0% RH prior to assessment of desiccation tolerance. Values are mean ± SEM of five groups of 10 individuals. Asterisks denote a significant difference relative to the control treatment (Student’s t- test).
vi 3.4 Fig. 4. Changes in (A) body water content (N = 15-20 individuals) and 53 (B) osmotic pressure of the body fluids (N = 6) of Cryptopygus antarcticus during slow cooling to -3.0oC in an environment at equilibrium with the vapor pressure of ice. Values are the mean ± SEM.
Chapter 4: 4.1 Fig.1. (A) Survival (N = five groups of 10 larvae), (B) water content 78 (N = 25-30), and (C) hemolymph osmolality (N = 6) of Belgica antarctica larvae exposed to a various concentrations of seawater or freshwater (~0 mOsm kg-1). Values are mean ± 1 SEM.
4.2 Fig. 2. Effect of acclimation to seawater (~1000 mOsm kg-1) and 79 resultant changes of body water content on the rate of oxygen consumption of Belgica antarctica larvae (y = -0.853 + 0.602x; R2 = 0.822; P<0.001).
4.3 Fig. 3. The effect of acclimation to seawater on the freeze tolerance of 80 Belgica antarctica larvae. Larvae were acclimated to either seawater (~1000 mOsm kg-1) or freshwater (~0 mOsm kg-1) for 3 d prior to assessment of freeze tolerance. A third group of larvae (rehydrated) were acclimated to seawater for 3 d followed by rehydration for 24 h in freshwater. Larvae were frozen in groups of 10 in ~100 µL of freshwater for 6 h. Values are mean ± 1 SEM of five groups of 10 larvae. Asterisks denote a significant difference relative to the freshwater (control) treatment (ANOVA, Dunnett’s test, P<0.05).
vii 4.4 Fig. 4. The effect of seawater acclimation on the desiccation tolerance 81 of Belgica antarctica larvae at 98.2 (A) or 75.0% RH (B) and 4oC. Larvae were acclimated to either seawater (~1000 mOsm kg-1) or freshwater (~0 mOsm kg-1) for 3 d prior to assessment of desiccation tolerance. Rehydrated larvae were acclimated to seawater for 3 d and then allowed to rehydrate for 24 h in freshwater prior to desiccation. Values are mean ± 1 SEM of five groups of 10 larvae. Asterisks denote a significant difference relative to the freshwater (control) treatment (ANOVA, Dunnett’s test, P<0.05).
4.5 Fig. 5. The effect of seawater acclimation on the heat shock tolerance 82 (time at 30oC) of Belgica antarctica larvae. Larvae were acclimated to either seawater (~1000 mOsm kg-1) or freshwater (~0 mOsm kg-1) for 3 d prior to assessment of heat shock tolerance. Rehydrated larvae were acclimated to seawater for 3 d and then allowed to rehydrate for 24 h in freshwater prior to heat shock. Values are mean ± 1 SEM of five groups of 10 larvae. Asterisks denote a significant difference relative to the freshwater (control) treatment (ANOVA, Dunnett’s test, P<0.05).
viii ACKNOWLEDGEMENTS
I wish to thank my advisor, Richard E. Lee, Jr., whose mentorship has provided me with the opportunity, guidance, and direction to complete this project. Through his collegiality and professionalism, Rick has been the ideal role model for what I hope to become as a faculty member. He has helped me develop as a learner, teacher and researcher of the biological sciences. I would also like to thank my other committee members, Alan Cady, Jon Costanzo, Kathleen Killian, and Robert Schaefer, for their guidance and assistance with various aspects of this work. I am also grateful to have had the opportunity to work with a number of excellent collaborators. This work would not have been possible if not for the logistical support and assistance from the staff at Palmer Station while working in the Antarctic. Finally, I am especially grateful to my family and friends for their constant support.
ix Chapter 1 General introduction Antarctica is the coldest, driest, and windiest continent on earth. It also is a continent of isolation, with the nearest continental land mass >1000 km away. Most of the continent is permanently covered by snow and ice, with ice-free terrestrial habitats representing <0.5% of the total surface area (Fox and Cooper, 1994). Such terrestrial habitats are largely restricted to the maritime Antarctic, including the west coast of the Antarctic Peninsula. In contrast to the relatively stable marine environment of the Southern Ocean, seasonally ice-free Antarctic terrestrial habitats are characterized by widely varying, often extreme environmental conditions on both daily and seasonal timescales (Convey, 1997). The most obvious environmental challenge facing the terrestrial biota is the low and, at least during summer, highly variable temperatures characteristic of the maritime Antarctic. Winter air temperatures may fall to -25oC near Palmer Station on the Antarctic Peninsula. However, terrestrial microhabitat sites harboring over-wintering arthropods remain between 0 and -3oC for much of winter, thanks to thermal buffering from the oceanic influence and snow and ice (Baust and Lee, 1981). Still, these arthropods must endure continuous, subzero temperatures for as much as 7-9 months (see Chapter 2). At the other extreme, microhabitat temperatures during summer, as result of absorption of solar radiation, can rise rapidly (>20oC in 6 h) and reach temperatures in excess of 30oC (Convey, 1997). Temperature clearly has another important effect in contributing to the availability of water. Water availability, even more so than temperature, is recognized as the most important determinant of the distribution of Antarctic terrestrial organisms (Kennedy, 1993). During winter, terrestrial organisms must endure desert-like conditions, as water is biologically unavailable in the form of ice. Summer presents much more variable moisture availability. Terrestrial microhabitats may dry rapidly as a result of insolation, wind, and rapid drainage of the typically ahumic soils (Kennedy, 1993). Alternatively, microhabitats may be inundated for days or weeks with freshwater from melting snow and ice or frequent rains. Organisms inhabiting these terrestrial microhabitats must be tolerant of such dramatic fluctuations in the availability of water during both the austral summer and winter (see Chapter 3). The extreme environmental challenges of the Antarctic are not restricted solely to temperature and water availability. Soil organisms, as a result of occasional seawater splash and
1 subsequent evaporation, may be subject to increasing salinity and hyperosmotic stress (see Chapter 4). Outwash from penguin colonies and seal wallows may inundate microhabitats with anoxic and acidic (pH 4.0) detrital effluent (Baust and Lee, 1987). Terrestrial organisms also are faced with dramatic seasonal fluctuations of light availability and the risk of exposure to ultraviolet radiation due to the annual formation of the Antarctic ozone hole. As a result of such extreme environmental conditions, along with limited habitat availability and continental isolation, the terrestrial fauna of the maritime Antarctic is rather depauperate (Convey, 1997). Of the terrestrial arthropods, the Acari (mites) and Collembola (springtails) dominate, with two Dipteran (flies) species also represented. Terrestrial organisms inhabiting the Antarctic Peninsula must possess adaptations and elicit responses, both behavioral and physiological, that facilitate survival under such harsh environmental conditions. Therefore, the purpose of this research was to investigate the tolerance and physiological mechanisms used to promote survival of a variety of the aforementioned environmental stressors in two Antarctic arthropods, the midge, Belgica antarctica, and the collembolan, Cryptopygus antarcticus. The terrestrial chironomid B. antarctica Jacobs is the southern-most free-living holometabolous insect, being sporadically dispersed, but locally abundant, on the west coast of the Antarctic Peninsula. Its two-year life cycle includes four larval stages and over-wintering may occur in any instar. Larvae typically winter within the upper few centimeters of the substrate, with pupation and adult emergence occurring in spring and summer. Larvae are freeze tolerant year round to ca. -15oC (Baust and Lee, 1981), and possess an extreme tolerance for desiccation; larvae survive the loss of ~70% of their body water (Baust and Lee, 1987; Benoit et al., 2007). Additionally, we recently documented a rapid cold-hardening (RCH) response in B. antarctica, whereby a brief exposure to -5oC extended the freeze tolerance of the larvae (Lee et al., 2006). Larvae are also tolerant of anoxia (>7 d at 0oC), freshwater immersion (>28 d at 0oC), salinity (>7 d in 0.5 M NaCl) and variations in pH (>14 d at pH 3-12) (Baust and Lee, 1987). This extreme tolerance is likely facilitated by the finding that larvae, but not adults, continually express several members of the heat shock protein (Hsp) family (Rinehart et al., 2006). The springtail C. antarcticus Willem is the most widespread and abundant terrestrial arthropod in the maritime Antarctic (Block, 1984), with large aggregations found on the underside of rocks, beneath mats of terrestrial algae, and in associations with mosses (Worland
2 and Block, 1986). Unlike B. antarctica, the springtail is freeze intolerant and must rely on a well developed ability for supercooling (i.e., remaining unfrozen) to survive subzero temperatures (Block et al., 1978; Sømme, 1978; Lee and Baust, 1981). The ability to supercool is facilitated by a RCH response allowing the springtails to depress their supercooling point in response to rapid changes in temperature (Worland and Convey, 2001). The tolerance and response of C. antarcticus to desiccation stress is less well studied; however, previous investigations have demonstrated that they have a limited resistance to dehydration at 0% relative humidity (Block et al., 1990; Harrison et al., 1991; Block and Harrison, 1995). Three investigations of the physiological tolerance and response to environmental stress in these Antarctic arthropods were conducted. The first project focused on the ability of B. antarctica larvae to resist freezing and undergo dehydration at subzero temperatures (i.e., cryoprotective dehydration) as a strategy for winter survival. The second investigation focused on the tolerance and physiological response of the springtail C. antarcticus to drought stress under ecologically-relevant desiccating conditions characteristic of both the austral summer and winter. Finally, the third study was designed to assess the salinity tolerance of the midge larvae during hyperosmotic seawater stress and the effect of a brief seawater acclimation on the subsequent tolerance of other environmental stressors (i.e., cross tolerance).
3 References Baust, J. G. and Lee, R. E. (1981). Environmental “homeothermy” in an Antarctic insect. Ant. J. US 15, 170-172. Baust, J. G. and Lee, R. E. (1987). Multiple stress tolerance in an Antarctic terrestrial arthropod: Belgica antarctica. Cryobiology 24, 140-147. Benoit, J. B., Lopez-Martinez, G., Michaud, M. R., Elnitsky, M. A., Lee, R. E., Jr., and Denlinger, D. L. (2007). Mechanisms to reduce dehydration stress in larvae of the Antarctic midge, Belgica antarctica. J. Insect Physiol. 53, 656-667. Block, W. (1984). Terrestrial microbiology, invertebrates and ecosystems. In: Laws, R. M. (Eds.). Antarctic ecology, vol. 1. Academic Press, London, pp 163-236. Block, W. and Harrison, P. M. (1995). Collembolan water relations and environmental change in the maritime Antarctic. Global Change Biol. 1, 347-359. Block, W., Harrison, P. M., and Vannier, G. (1990). A comparative study of patterns of water loss from two Antarctic springtails (Insecta, Collembola). J. Insect Physiol. 36, 181-187. Block, W., Young, S. R., Conradi-Larsen, E. M., and Somme, L. (1978). Cold tolerance of two Antarctic arthropods. Experientia 34, 1166-1167. Convey, P. (1996). The influence of environmental characteristics on life history attributes of Antarctic terrestrial biota. Biol. Rev. 71, 191-225. Convey, P. (1997). How are the life history strategies of Antarctic terrestrial invertebrates influenced by extreme environmental conditions? J. Therm. Biol. 22, 429-440. Fox, A. J. and Cooper, A. P. R. (1994). Measured properties of the Antarctic ice sheet derived from the SCAR Antarctic digital database. Polar Res. 30, 206-210. Harrison, P. M., Rothery, P. and Block, W. (1991). Drying processes in the Antarctic collembolan Cryptopygus antarcticus (Willem). J. Insect Physiol. 37, 883-890. Kennedy, A. D. (1993). Water as a limiting factor in the Antarctic terrestrial environment: a biogeographical synthesis. Arctic Alpine Res. 25, 308-315. Lee, R. E., Elnitsky, M. A., Rinehart, J. P., Hayward, S. A. L., Sandro, L. H., and Denlinger, D. L. (2006). Rapid cold-hardening increases the freezing tolerance of the Antarctic midge, Belgica antarctica. J. Exp. Biol. 209, 399-406. Sømme, L. (1978). Cold-hardiness of Cryptopygus antarcticus (Collembola) from Bouvetoya. Oikos 31, 94-97.
4 Worland, M. R. and Block, W. (1986). Survival and water loss in some Antarctic arthropods. J. Insect Physiol. 32, 579-584. Worland, M. R. and Convey, P. (2001). Rapid cold hardening in Antarctic microarthropods. Funct. Ecol. 15, 515-524.
5 Chapter 2
Cryoprotective dehydration and the resistance to inoculative freezing in the Antarctic midge, Belgica antarctica
6 Summary During winter, larvae of the Antarctic midge, Belgica antarctica (Diptera, Chironomidae), must endure 7-8 months of continuous subzero temperatures, encasement in a matrix of soil and ice, and severely desiccating conditions. This environment, along with the fact that larvae possess a high rate of water loss and are extremely tolerant of desiccation, may promote the use of cryoprotective dehydration as a strategy for winter survival. This study investigates the capacity of larvae to resist inoculative freezing and undergo cryoprotective dehydration at subzero temperatures. Slow cooling to -3oC in an environment at equilibrium with the vapor pressure of ice reduced larval water content by ~40% and depressed the body fluid melting point more than 3-fold to -2.6oC. This melting point depression was the result of the concentration of existing solutes (i.e., loss of body water) and the de novo synthesis of osmolytes. By day 14 of the subzero exposure, larval survival was still >95%, suggesting larvae have the capacity to undergo cryoprotective dehydration. However, under natural conditions the use of cryoprotective dehydration may be constrained by inoculative freezing as result of the insect’s intimate contact with environmental ice. During slow cooling within a substrate of frozen soil, the ability of larvae to resist inoculative freezing and undergo cryoprotective dehydration was dependent upon the moisture content of the soil. As detected by a reduction of larval water content, the percentage of larvae that resisted inoculative freezing increased with decreasing soil moisture. These results suggest that larvae of the Antarctic midge have the capacity to resist inoculative freezing at relatively low soil moisture contents and likely undergo cryoprotective dehydration when exposed to subzero temperatures during the polar winter.
Key words: Chironomidae, cryoprotective dehydration, freeze tolerance, supercooling
7 Introduction Cold-hardy invertebrates can be classified most simply as freeze-tolerant or freeze- intolerant. Freeze-tolerant species survive the freezing of their body fluids by promoting ice nucleation at high subzero temperatures and through the seasonal accumulation of cryoprotectants (Zachariassen, 1985; Duman et al., 1991; Lee, 1991). For freeze-intolerant species, by contrast, internal ice formation is ultimately lethal and survival depends upon prolonged, and often extensive, supercooling. Supercooling requires the absence or masking of potential ice nucleators within the body fluids as well as behavioral and/or physiological mechanisms that prevent inoculative freezing by environmental ice (Lee et al., 1995). Water also may be biologically unavailable in the form of ice, and, therefore, supercooled insects may be subjected to extended periods of desiccation as well (Lundheim and Zachariassen, 1993).
A third strategy of over-wintering, termed cryoprotective dehydration, has been described for several freeze-intolerant soil invertebrates (Holmstrup, 1992; Holmstrup and Westh, 1994; Holmstrup and Sømme, 1998; Holmstrup et al., 2002). In this strategy, supercooled invertebrates with high integumental permeability dehydrate when exposed to an environment at equilibrium with the vapor pressure of ice, owing to vapor pressure differences between supercooled water and ice at the same temperature (Holmstrup and Sømme, 1998). Such water loss continues until, at equilibrium, the vapor pressure of the body fluids equals that of the surrounding ice. At this time, the risk of freezing has been eliminated because the melting point (MP) of the animal’s body fluids equals the ambient temperature (Holmstrup et al., 2002). Equilibration of the body fluid MP with that of the environment may also be facilitated by the accumulation of cryoprotectants (Holmstrup, 1995; Worland et al., 1998). More recently, cryoprotective dehydration has been reported for a freeze-tolerant nematode (Wharton et al., 2003) and an enchytraeid worm (Pedersen and Holmstrup, 2003), however, use of this strategy may be constrained by inoculative freezing of the body fluids as result of contact with environmental ice.
The terrestrial chironomid Belgica antarctica is the southern-most free-living holometabolous insect, being sporadically dispersed, but locally abundant, on the west coast of the Antarctic Peninsula. Detailed accounts of the life-history and ecology of B. antarctica are provided by Convey and Block (1996), Sugg et al. (1983), Usher and Edwards (1984), and references cited therein. Briefly, its two-year life cycle includes four larval stages and over-
8 wintering may occur in any instar. Larvae typically over-winter within the upper few centimeters of the substrate, with pupation and adult emergence occurring in spring and summer. The adults are wingless, like many insects in wind-swept alpine and oceanic habitats, and live for fewer than 14 days.
While ambient air temperatures on the Antarctic Peninsula may reach winter lows of - 30oC, larvae of B. antarctica survive freezing to only ca. -15 to -20oC (Baust and Lee, 1981; Lee et al., 2006). However, thermal buffering of the over-wintering hibernaculum, provided by the oceanic influence and up to a meter of ice and snow, apparently explains this anomaly; at one cm depth, substrate temperatures remain between 0 and -2oC for more than 300 days of the year, and rarely decrease below -5oC (Baust and Lee, 1981). As larvae maintain relatively constant supercooling points between -6 and -8oC throughout the year (Baust and Lee, 1987), freezing of body fluids during over-wintering likely occurs via inoculation from the external environment. Alternatively, upon freezing of the surrounding substrate larvae may dehydrate, equilibrating their body fluid MP with the ambient temperature, thereby remaining unfrozen during over- wintering. Such a strategy of cryoprotective dehydration necessitates a high rate of water loss and/or a resistance to avoid inoculation of the supercooled body fluids.
Anecdotal reports suggest that several Arctic chironomids dehydrate during the winter (Scholander et al., 1953; Danks, 1971), thus the capacity for dehydration may be present within this taxonomic group. In addition, B. antarctica is highly desiccation tolerant, as larvae tolerate dehydration to ~30% of their initial body weight (Baust and Lee, 1987; Hayward et al., 2007; Benoit et al., 2007), and possess a high rate of water loss even at high relative humidities. Therefore, the purpose of the present study was to assess the capacity of larval B. antarctica to resist inoculative freezing and undergo cryoprotective dehydration when exposed to subzero temperatures.
Materials and methods Source of insects Substrate containing larval B. antarctica Jacobs was collected from sites near penguin rookeries on Torgersen Island, near Palmer Station on the Antarctic Peninsula (64o46’ S, 64o04’ W) in January 2005. Samples were shipped frozen (ca. -5oC for 7 days) to Miami University and subsequently stored at 4oC (0L: 24D) prior to use. Larvae were handpicked from the substrate
9 and held in water at 4oC for 12-24 h to ensure clearance of the gut (mean gut clearance ~6 h; Baust and Edwards, 1979) and to standardize body water content prior to use. Only 4th instar larvae were used for experiments.
Microhabitat temperature Ten miniature temperature loggers (HOBO Water Temp Pro, Onset Computer, Pocasset, MA, USA) were deployed in microhabitat sites containing larval and adult B. antarctica on Torgersen Island in January 2005. Loggers recorded temperature at 30-min intervals for the duration of the study period. The loggers were recovered in January 2006 and the resulting data analyzed using Boxcar Pro 4.3 software (Onset Computer, Pocasset, MA, USA).
Cryoprotective dehydration The capacity of B. antarctica to undergo cryoprotective dehydration was assessed by exposing larvae to an environment at equilibrium with the vapor pressure of ice as described in Pedersen and Holmstrup (2003). Groups of five individuals were blotted dry and placed within 0.6-ml polyethylene microcentrifuge tubes. Larvae were confined by means of fine (~20 μm) nylon mesh that allowed free movement of water vapor. Microcentrifuge tubes were in turn placed within 15-ml glass vials containing ~5 g of crushed ice and closed with tightly-fitting lids. Vials containing larvae were allowed to equilibrate in refrigerated baths at -0.6 ± 0.1oC for 24 h. The temperature of the bath was then lowered incrementally (~0.5oC · day-1) to approximately - 3.0 ± 0.1oC and held there for an additional 10 days. A control group of larvae was held at -0.6 ± 0.1oC until termination of the experiment (day 14). Groups of larvae for body water content (WC) and body fluid melting point (MP) measurements were removed at 1-4 day intervals. The WC of individual larvae was assessed gravimetrically from measurements of fresh weight (to the nearest 0.01 mg) at the time of sampling and dry mass (DM) after drying to constant mass at 65oC. Melting point determinations were made using a vapor pressure depression technique (Holmstrup and Sømme, 1998). Groups of 5 larvae were placed in a sample holder, crushed with a Teflon rod to expose the body fluids, and rapidly sealed within a C-52 sample chamber (Wescor Inc., Logan, UT, USA). Samples were then allowed to equilibrate for 30 min prior to measurement of the body fluid MP using a Wescor HR-33T Dew Point Microvoltmeter (Wescor Inc., Logan, UT, USA)
10 operated in the dew point mode. Sample melting point was determined from standard curves produced from known salt solutions (Opti-Mole, Wescor Inc., Logan UT, USA). Survival was assessed on remaining larvae upon termination of the subzero exposure. Water (~80-100 µl) was added to the microcentrifuge tubes and larvae allowed to rehydrate/recover for 24 h at 4oC prior to survival assessment. Individuals displaying spontaneous movements were considered to have survived.
Cryoprotectant analysis Cryoprotectant analysis was performed on larvae following slow cooling to -3oC in an environment at equilibrium with the vapor pressure of ice as described above. Control larvae were maintained at -0.6 ± 0.1oC until termination of the experiment (day 14). Six groups of ~25 larvae were weighed and immediately frozen at -80°C until whole body concentrations of cryoprotectants were determined. Prior to cryoprotectant analysis, larvae were homogenized in 7% perchloric acid and neutralized with equal volumes of 0.789 M potassium hydroxide. Glycerol content was determined enzymatically as described by Holmstrup et al. (1999). Sorbitol concentrations were measured on the same individuals using an enzymatic assay described in Bergmeyer et al. (1974). Trehalose content was determined following digestion with trehalase as described by Chen et al. (2002). Glucose concentration was determined using the glucose oxidase procedure (no. 510; Sigma, St. Louis, MO, USA).
Exposure in frozen substrate The ability to use the strategy of cryoprotective dehydration may be limited in B. antarctica, as over-wintering larvae are likely in direct contact with ice and may be susceptible to inoculative freezing. Therefore, to assess their ability to resist inoculative freezing and undergo protective dehydration larvae were slowly cooled in contact with frozen substrate. Groups of 10 larvae were placed in 35-mm diameter Petri dishes containing ~ 4 g of loosely packed substrate, predominantly sand and organic matter, collected from Torgersen Island. Larger stones were removed and the soil mixed to achieve a relatively homogeneous substrate. Substrate samples were dried at 65oC prior to rehydration to the desired soil moisture contents.
The soil moisture content of field samples collected near Palmer Station was 1.10 ± 0.08 g H2O · g-1 dry soil (R. E. Lee and L. Sandro, unpublished). However, due to the sandiness of the soil,
11 larvae in the field likely experience large fluctuations of soil moisture. Therefore, three soil -1 moistures (0.80, 1.10 and 1.40 g H2O · g dry soil) were tested. Petri dishes were subsequently sealed with Parafilm to prevent evaporation. Larvae were allowed to equilibrate at -0.2 ± 0.1oC for 24 h and an additional 24 h at -1.0 ± 0.1oC within refrigerated baths. Freezing of the soil was then induced by lightly applying freezing spray to the exterior of the Petri dish; soils were allowed to freeze overnight at -1.0 ± 0.1oC and visually confirmed to have frozen by the presence of ice crystals within the soil matrix. The temperature of the bath was then lowered incrementally (~0.5oC · day-1) to approximately -3.0 ± 0.1oC and held there for an additional 7 days. Control groups of larvae were held in unfrozen substrate at -0.2 ± 0.1oC until the termination of the experiment. The WC of individual larvae was assessed at approximately 7-day intervals. Frozen soils were rapidly thawed and larvae collected and gently blotted to remove surface water prior to determination of WC as described above. As it was not possible to monitor individual larvae for freezing exotherms, WC was used as an indication of whether larvae remained supercooled, and therefore lost body water, or were frozen inoculatively (i.e., maintained high WC) during the subzero exposure. Following termination of the exposure (day 16), remaining larvae were allowed to recover at 4oC for 24 h prior to survival assessment as described above.
Statistical analysis Means were compared using Student’s t-tests or analysis of variance (ANOVA) and Bonferroni-Dunn tests (Statview from SAS Institute, Cary, NC, USA). Survival data were arcsin-square root transformed prior to analysis. Data are presented as mean ± 1 SEM. Statistical significance was set at P<0.05.
Results Microhabitat temperatures Summer temperatures (i.e., January and February) in microhabitat sites of larval B. antarctica often exceeded 15oC (Fig. 1), and occasionally 20oC. Conversely, subzero temperatures also occurred during summer months. Microhabitat temperatures declined throughout the summer, finally stabilizing at subzero temperatures during mid to late-April, and remained below zero until mid-November (Fig. 1). Thanks to oceanic thermal buffering (Baust
12 and Lee, 1981), as well as thermal buffering from snow and ice, winter temperatures in microhabitat sites generally remained between -1 and -3oC, and only rarely fell below -5oC.
Cryoprotective dehydration o -1 Larvae equilibrated to -0.6 C had a mean (± SEM) WC of 2.67 ± 0.05 g H2O · g DM (N = 15), with no significant change in the WC or DM of control animals during the experiment. In contrast, slow cooling in an environment at equilibrium with the vapor pressure of ice resulted in a significant (P<0.001) reduction of larval WC (Fig. 2A). Larval WC decreased rapidly through day 10, prior to leveling off for the remainder of the experiment. Water content was reduced by -1 ~40%, to 1.63 ± 0.03 g H2O · g DM (N = 15), by day 14; larval DM did not change significantly over the course of the experiment. These larvae appeared mildly dehydrated, however, survival at termination of the experiment was >96% (N = 30). Similarly, all control larvae (N = 30), maintained at -0.6oC, survived the 14-day exposure. The MP of larvae equilibrated to -0.6oC was -0.74 ± 0.02oC (N = 6) and did not change in control animals during the course of the experiment. However, cooling while at equilibrium with the vapor pressure of ice resulted in a significant (P<0.0001) reduction of larval MP (Fig. 2B). In contrast to WC, the MP decreased throughout the 14-day experiment. Relative to the controls, the MP was reduced more than 3-fold, to -2.61 ± 0.03oC (N = 6), by day 14 in larvae exposed to the vapor pressure of ice. At the termination of the experiment, the body fluid MP had been depressed to within 0.4oC of the final ambient temperature (Fig. 2B). However, there remained a vapor pressure deficit of approximately -5.0 bar between the body fluids of the larvae and the surrounding environment. Therefore, larvae would be expected to continue to lose water to their environment, further depressing the MP of the body fluids.
Cryoprotectant analysis Of the cryoprotectants measured in larvae, only glucose and trehalose were found in significant concentrations (Table 1). Control larvae maintained relatively low concentrations of both glucose and trehalose throughout the experiment. Conversely, both glucose and trehalose were increased significantly (P<0.0001) in larvae that were cooled at equilibrium with the vapor pressure of ice. By day 14 of the subzero exposure, glucose and trehalose concentrations in larvae were increased by ~9 and 11-fold, respectively.
13
Exposure in frozen soil The WC of larvae equilibrated at -0.2oC did not differ significantly among the soil -1 moisture contents tested (combined mean 2.48 ± 0.04 g H2O · g DM; N = 30). Similarly, neither WC nor DM changed significantly in control animals during the 16-day exposure for any soil moisture tested. However, cooling larvae in contact with frozen substrate resulted in a clear -1 -1 separation, based upon WC, into high (>2.20 g H2O · g DM) and low (<1.80 g H2O · g DM) groups (Fig. 3). Larvae in the ‘high’ WC groups, designated as frozen in Fig. 3, presumably froze inoculatively soon after the soil was frozen. These larvae remained in vapor pressure equilibrium with the surrounding environment, and, therefore, did not lose substantial amounts of water. For all levels of soil moisture tested, the mean WC of frozen larvae did not differ significantly from control larvae maintained at -0.2oC (Fig. 3). In contrast, larvae in the ‘low’ WC group, designated as dehydrated in Fig. 3, presumably remained unfrozen and dehydrated; larvae lost body water as a result of the vapor pressure gradient between the unfrozen body fluids and the surrounding environmental ice. By day 16, the WC of larvae that dehydrated was significantly (P<0.001) lower than that of control larvae for all three soil moistures tested (Fig. 3). There was no significant difference in DM between ‘high’ and ‘low’ WC groups in any soil moisture tested at the termination of the experiment. Based upon reductions of larval WC during cooling in contact with frozen soil, the ability of larvae to resist inoculative freezing and, therefore, use a strategy of cryoprotective dehydration was significantly (P<0.001) affected by soil moisture content (Fig. 4); the percentage of larvae frozen inoculatively increased with increasing soil moisture. Less than 50% of larvae were frozen when cooled in contact with substrate at a soil moisture content of 0.80 g -1 -1 H2O · g dry soil. However, at soil moisture contents of 1.10 and 1.40 g H2O · g dry soil <30% of larvae were able to resist inoculative freezing and use a strategy of cryoprotective dehydration (Fig. 4). Regardless, larval survival of the 16-day exposure was >96% (N = 30) for all soil moistures tested and did not differ significantly from control larvae maintained at -0.2oC.
Discussion Cryoprotective dehydration in a polar insect
14 During winter B. antarctica larvae are likely to be encased in a matrix of soil and ice for 7 – 8 months. However, microhabitat temperatures remained between 0 and -3oC throughout much of winter, and only rarely fell below -5oC (Fig. 1; Baust and Lee, 1981). Such conditions may result in the freezing of the body fluids through inoculation from environmental ice. Alternatively, if larvae can resist inoculative freezing, these environmental conditions would create a gradient for water loss from supercooled larvae due to the lower vapor pressure of the surrounding ice. Our results demonstrate that B. antarctica larvae do indeed dehydrate when exposed to an environment at equilibrium with the vapor pressure of ice (Fig. 2A). This water loss, in addition to the de novo synthesis of osmolytes, depressed the MP of the larvae’s body fluids to nearly -3oC (Fig. 2B), suggesting B. antarctica larvae have the capacity to undergo cryoprotective dehydration at ecologically relevant subzero temperatures. While protective dehydration has previously been documented in earthworm cocoons (Holmstrup, 1992; Holmstrup and Westh, 1994), Collembola (Worland et al., 1998; Holmstrup et al., 2002), a nematode (Wharton et al., 2003), and an enchytraeid worm (Pedersen and Holmstrup, 2003), our study is the first report of cryoprotective dehydration in a true insect. The melting point depression observed in B. antarctica was a consequence of both water loss, and resultant increase in the concentration of the original solutes, as well as a concomitant synthesis of osmolytes (Table 1). As a result of the vapor pressure gradient between body fluids and the surrounding ice, larvae lost water throughout the subzero temperature exposure (Fig. 2A). At the termination of the experiment (day 14), larvae had lost ~40% of their total body water. However, estimates of the osmotically active water content, based upon measurements in Worland et al. (1998), suggest that water loss, and the associated concentration effect, explained only ~57 and 51% of the observed change in osmotic pressure at days 6 and 14 during cryoprotective dehydration, respectively (Table 1). This is in contrast to the enchytraeid Fridericia ratzeli, in which MP depression to nearly -6oC during cryoprotective dehydration is accomplished largely (~83%) through water loss and the concentration of the original solutes (Pedersen and Holmstrup, 2003). At the termination of the experiment, there still remained a vapor pressure gradient of approximately -5 bar between the body fluids and the surrounding ice. Therefore, larvae would be expected to continue to lose water to their environment, further depressing the MP of their body fluids. Additionally, as the WC continues to decrease, even small reductions in WC would
15 have a large effect on the resulting MP, as the effects of further water loss will increase the MP in a hyperbolic manner (Ring, 1982; Holmstrup and Westh, 1994). Such continued dehydration is unlikely to affect survival, at least in the short-term, as B. antarctica larvae tolerate the loss of ~70% of their total body water (Benoit et al., 2007). During cryoprotective dehydration, larvae accumulated significant amounts of osmolytes that further depressed the MP of the body fluids (Table 1). Of the sugars and polyols assayed, only glucose and trehalose were detected in B. antarctica larvae; relative to control larvae, glucose and trehalose concentrations were ~9 and 11-fold higher, respectively, following cryoprotective dehydration. The Arctic collembolan, Onychiurus arcticus, likewise accumulates significant concentrations of both glucose and trehalose during cryoprotective dehydration (Worland et al., 1998). Further, F. ratzeli accumulates similar concentrations of glucose during cryoprotective dehydration as we observed in B. antarctica larvae (Pedersen and Holmstrup, 2003). In addition to facilitating depression of the melting point, such sugars and polyols are well known to protect membranes and proteins against the deleterious effects of low temperature and desiccation (Crowe et al., 1992; Storey, 1997). While the observed increase in glucose and trehalose levels was substantial, these osmolytes accounted for a rather small fraction of the overall increase (<12%) in osmotic pressure during dehydration (Table 1). Together with the concentration of the original solutes, as the result of dehydration, the synthesis of the measured sugars accounted for <65% of the observed change in osmotic pressure. The identity of the solutes making up the remainder of the observed osmotic pressure following dehydration is unknown. However, B. antarctica larvae are known to accumulate a variety of other sugars and polyols, including erythritol, sucrose, and fructose (Baust and Edwards, 1979; Baust, 1980; Baust and Lee, 1983). Additionally, other cold-hardy insects accumulate significant concentrations of free amino acids during desiccation and acclimation to low temperature (Storey and Storey, 1988). Such increases in other cryoprotectants may account for a portion of the change in osmotic pressure during cryoprotective dehydration. It should also be noted that estimates of the contribution of dehydration and the synthesis of osmolytes to account for the observed increase in osmotic pressure during cryoprotective dehydration are highly dependent upon the fraction of the total water content of larvae that is osmotically active (Table 1). This is especially true when total body water is low, as even a
16 slight change in the osmotically active water (OAW) content would significantly affect estimates of the contribution to the measured osmotic pressure. Our estimates in accounting for the observed osmotic pressure necessarily rely on measures of osmotically active and inactive water from the collebolan O. arcticus (Worland et al., 1998), as, unfortunately, similar data do not exist for the midge larvae. These values may or may not be representative for B. antarctica larvae, and, therefore, may also account for a portion of the unexplained increase in osmotic pressure during dehydration.
Resistance to inoculative freezing The ability to use the strategy of cryoprotective dehydration for subzero temperature survival may be constrained by inoculative freezing, as over-wintering larvae are likely to be in intimate contact with environmental ice. During the period in which the larvae are in a supercooled state (i.e., during cooling), they are at risk of inoculation from the surrounding ice. If the insect remains unfrozen during cooling, body water is lost, as a result of the vapor pressure gradient, the melting point of the body fluids equilibrates with the ambient temperature, and the risk of freezing is eliminated. Therefore, a high rate of water loss, which B. antarctica larvae certainly possess (this study; Benoit et al., 2007), allows the organism to ‘track’ environmental changes and rapidly equilibrate the MP of the body fluids to that of the ambient temperature, thereby eliminating the risk of freezing. Based upon reductions of larval WC, the resistance to inoculative freezing of B. antarctica depended on the moisture content of the soil in which they were cooled. As the water content of the soil increased, and therefore the amount of ice surrounding the larvae, the -1 percentage of larvae that froze likewise increased (Fig. 4). At soil moistures of >1.10 g H2O · g dry soil, fewer than 30% of larvae resisted inoculative freezing. These larvae almost certainly froze due to inoculation during cooling to -3oC, since the supercooling point of larvae remains between approximately -6 and -8oC year-round (Baust and Lee, 1987). Additionally, larvae were likely frozen soon after freezing of the substrate, as evidenced from the maintenance of the high -1 body water content in frozen larvae (Fig. 3). In contrast, at 0.80 g H2O · g dry soil, nearly 60% of larvae remained unfrozen and dehydrated. This suggests that under relatively dry conditions larvae can resist inoculative freezing and use a strategy of cryoprotective dehydration to survive subzero temperatures.
17 The rate of cooling is also likely to determine the resistance to inoculative freezing and the use of cryoprotective dehydration. Wharton et al. (2003) demonstrated that in the Antarctic nematode Panagrolaimus davidi, slower cooling rates significantly increase the percentage of nematodes that resist freezing and undergo cryoprotective dehydration. The cooling rate of 0.5oC · day-1 used in our study corresponded to a realistic, but relatively rapid rate of cooling compared to natural conditions (Fig. 1). As winter in the Antarctic begins and soils freeze, the cooling of larvae would likely be moderated by the thermal inertia of the soil and buffering from snow and ice. At slower cooling rates, water loss from the larvae, and the corresponding MP depression, could likely keep pace and ‘track’ changes of ambient temperature, therefore, increasing the likelihood that larvae undergo cryoprotective dehydration.
Cryoprotective dehydration vs. freezing for winter survival Our results suggest that B. antarctica larvae can survive ecologically relevant subzero temperatures using either freeze tolerance or cryoprotective dehydration. The strategy used for winter survival likely depends upon the ambient environmental conditions upon entrance into winter. At relatively low soil moistures, the high subzero temperatures and slow rates of cooling within the larval microhabitat, in addition to the extreme tolerance of desiccation and high rate of water loss of B. antarctica, increase the likelihood that larvae can resist inoculative freezing and undergo cryoprotective dehydration during the polar winter. However, even if larvae are frozen after a period of dehydration this likely only enhances survival, as mild dehydration increases the freeze tolerance of B. antarctica larvae (Hayward et al., 2007). Finally, while there did not appear to be differences in survival of low temperature between strategies, future studies should address the long-term fitness consequences (e.g., Irwin and Lee, 2002) of the use of freeze tolerance or cryoprotective dehydration for winter survival.
Acknowledgements This research was supported by NSF grants OPP-0337656 and OPP-0413786. We thank the support staff at Palmer Station for their excellent assistance with this project. Ben Philip provided useful comments and suggestions on a draft of the manuscript.
18 References Baust, J. G. (1980). Low temperature tolerance in an Antarctic insect: A relict adaptation?. CryoLetters 1, 360-371. Baust, J. G. and Edwards, J. S. (1979). Mechanisms of freezing tolerance in an Antarctic midge, Belgica antarctica. Physiol. Entomol. 4, 1-5. Baust, J. G. and Lee, R. E. (1981). Environmental “homeothermy” in an Antarctic insect. Antarct. J. US 15, 170-172. Baust, J. G. and Lee, R. E. (1983). Population differences in antifreeze/cryoprotectant accumulation patterns in an Antarctic insect. Oikos 40, 120-124. Baust, J. G. and Lee, R. E. (1987). Multiple stress tolerance in an Antarctic terrestrial arthropod: Belgica antarctica. Cryobiology 24, 140-147. Benoit, J. B., Lopez-Martinez, G., Michaud, M. R., Elnitsky, M. A., Lee, R. E., and Denlinger, D. L. (2007). Mechanisms to reduce dehydration stress in the Antarctic midge, Belgica antarctica. J. Insect Physiol., in press. Bergmeyer, H. U., Gruber, W. and Gutmann, I. (1974). D-Sorbitol. In Methods of Enzymatic Analysis, vol. 3 (ed. H. U. Bergmeyer), pp. 1323-1326. New York: Academic Press. Chen, Q., Ma, E., Behar, K. L., Xu, T., and Haddad, G. G. (2002). Role of trehalose phosphate synthase in anoxia tolerance and development in Drosophila melanogaster. J. Biol. Chem. 277, 3274-3279. Convey, P. and Block, W. (1996). Antarctic Diptera: Ecology, physiology and distribution. Eur. J. Entomol. 93, 1-13. Crowe, J. H., Hoekstra, F. A., and Crowe, L. M. (1992). Anhydrobiosis. Ann. Rev. Physiol. 54, 579-599. Danks, H. V. (1971). Overwintering of some north temperate and Arctic Chironomidae. II. Chironomid biology. Can. Entomol. 103, 1875-1910. Duman, J. G., Wu, D. W., Xu, L., Tursman, D., and Olsen, T. M. (1991). Adaptations of insects to subzero temperatures. Quart. Rev. Biol. 66, 387-410. Hayward, S. A. L., Rinehart, J. P., Sandro, L. H., Lee, R. E., and Denlinger, D. L. (2007). Slow dehydration promotes desiccation and freeze tolerance in the Antarctic midge Belgica antarctica. J. Exp. Biol. 210, 836-844.
19 Holmstrup, M. (1992). Cold hardiness strategy in cocoons of the lumbricid earthworm Dendrobaena octaedra (Savigny). Comp. Biochem. Physiol. 102, 49-54. Holmstrup, M. (1995). Polyol accumulation in earthworm cocoons induced by dehydration. Comp. Biochem. Physiol. 111, 251-255. Holmstrup, M. and Sømme, L. (1998). Dehydration and cold hardiness in the Arctic collembolan Onychiurus arcticus Tullberg 1876. J. Comp. Physiol. B 168, 197-203. Holmstrup, M. and Westh, P. (1994). Dehydration of earthworm cocoons exposed to cold: a novel cold hardiness mechanism. J. Comp. Physiol. B 164, 312-315. Holmstrup, M., Bayley, M., and Ramløv, H. (2002). Supercool or dehydrate? An experimental analysis of overwintering strategies in small permeable Arctic invertebrates. Proc. Natl. Acad. Sci. USA 99, 5716-5720. Holmstrup, M., Costanzo, J. P. and Lee, R. E. (1999). Cryoprotective and osmotic responses to cold acclimation and freezing in freeze-tolerant and freeze-intolerant earthworms. J. Comp. Physiol. B 169, 207-214. Irwin, J. T. and Lee, R. E. (2002). Energy and water conservation in frozen vs. supercooled larvae of the goldenrod gall fly, Eurosta solidaginis (Fitch) (Diptera: Tephritidae). J. Exp. Zool. 292, 345-350. Lee, R. E. (1991). Principles of insect low temperature tolerance. In: Insects at Low Temperature, (ed. R.E. Lee and D.L. Denlinger), pp. 17-46, New York: Chapman and Hall. Lee, R. E., Elnitsky, M. A., Rinehart, J. P., Hayward, S. A. L., Sandro, L. H., and Denlinger, D. L. (2006). Rapid cold-hardening increases the freezing tolerance of the Antarctic midge, Belgica antarctica. J. Exp. Biol. 209, 399-406. Lee, R. E., Warren, G. J., and Gusta, L. V. (1995). Biological Ice Nucleation and its Applications. St. Paul: APS Press. Lundheim, R. and Zachariassen, K. E. (1993). Water balance of over-wintering beetles in relation to strategies of cold tolerance. J. Comp. Physiol. B 163, 1-4. Pedersen, P. G. and Holmstrup, M. (2003). Freeze or dehydrate: only two options for the survival of subzero temperatures in the arctic enchytraeid Fridericia ratzeli. J. Comp. Physiol. B 173, 601-609.
20 Ring, R. A. (1982). Freezing-tolerant insects with low supercooling points. Comp. Biochem. Physiol. 73A, 605-612. Scholander, P. F., Flagg, W., Hock, R. J., and Irving, L. (1953). Studies on the physiology of frozen plants and animals in the arctic. J. Cell. Comp. Physiol. 42, 1-56. Storey, K. B. (1997). Organic solutes in freezing tolerance. Comp. Biochem. Physiol. A 117, 319-326. Storey, K. B. and Storey, J. M. (1988). Freeze tolerance in animals. Physiol. Rev. 68, 27-84. Sugg, P., Edwards, J. S., and Baust, J. (1983). Phenology and life history of Belgica antarctica, an Antarctic midge (Diptera: Chironomidae). Ecol. Entomol. 8, 105-113. Usher, M. B. and Edwards, M. (1984). A dipteran from south of the Antarctic Circle: Belgica antarctica (Chironomidae), with a description of its larva. Biol. J. Linn. Soc. 23, 19-31. Wharton, D. A., Goodall, G., and Marshall, C. J. (2003). Freezing survival and cryoprotective dehydration as cold tolerance mechanisms in the Antarctic nematode Panagrolaimus davidi. J. Exp. Biol. 206, 215-221. Worland, M., Grubor-Lajsic, G., and Montiel, P. (1998). Partial desiccation induced by sub- zero temperatures as a component of the survival strategy of the Arctic collembolan Onychiurus arcticus (Tullberg). J. Insect Physiol. 44, 211-219. Zachariassen, K. E. (1985). Physiology of cold tolerance in insects. Physiol. Rev. 65, 799-832.
21 Table 1. Estimated osmotic contribution of initial osmolytes in the hemolymph and osmolytes produced during slow cooling to -3oC in an environment at equilibrium with the vapor pressure of ice. Values are mean ± SEM.
Control Cryoprotective Dehydration
Day 6 Day 14
Observed osmotic pressure of body fluids 398 ± 10 991 ± 6 1392 ± 16 (mOsm)
-1 Total body water content (g H2O · g DM) 2.67 ± 0.05 1.99 ± 0.06 1.63 ± 0.03
Osmotically active water (OAW) 2.19 1.55 1.22 -1 content (g H2O · g DM)*
Loss of OAW (%) --- 29.2 44.3
Osmotic contribution of original solutes --- 562 715 due to loss of OAW (mOsm)
Concentration of osmolytes (µg · mg-1 DM) Glycerol ~ 0 ~ 0 ~ 0 Sorbitol ~ 0 ~ 0 ~ 0 Glucose 2.15 ± 0.24 12.41 ± 0.12 18.46 ± 0.24 Trehalose 3.32 ± 0.52 19.48 ± 0.67 37.42 ± 0.31
Osmotic contribution of synthesized --- 67.2 155.9 osmolytes† (mOsm) Total explainable osmotic pressure (%) --- 63.5 62.6
* OAW was calculated from Worland et al. (1998). [(OIW) = 0.069(TBW) + 0.3, where OIW is osmotically inactive water content, TBW is total body water content and is the sum of OIW and OAW] † Assuming that osmolytes are dissolved in OAW.
22 Figure legends Fig. 1. Seasonal changes in temperature at a representative larval Belgica antarctica microhabitat site on Torgersen Island, near Palmer Station, Antarctica (64o46’ S, 64o04’ W). Microhabitat temperatures were measured in 2005-2006 using single-channel temperature loggers (HOBO Water Temp Pro, Onset Computer, Pocasset, MA, USA). The dashed line represents the equilibrium freezing point of the body fluids of fully hydrated, control larvae.
Fig. 2. Changes in (A) body water content (N = 15) and (B) body fluid melting point (N = 6) of larval Belgica antarctica during slow cooling to -3oC in an environment at equilibrium with the vapor pressure of ice. Different letters indicate significant differences between values (ANOVA, Bonferroni-Dunn test, P<0.05). Values are mean ± SEM.
Fig. 3. Body water content (WC) of individual Belgica antarctica larvae (N = 30) during slow cooling to -3oC in contact with substrates of varying moisture content: 0.80, 1.10, and 1.40 -1 g H2O · g dry soil. Triangles denote WC of individuals at day 0, circles the WC of frost exposed individuals (day 16). Dashes denote the mean WC of individuals at day 0 and 16, separated into ‘high’ (frozen) and ‘low’ (dehydrated) WC groups.
Fig. 4. Percentage of Belgica antarctica larvae frozen, as detected by the maintenance of ‘high’ body water content, during cooling to -3oC in contact with substrates of varying moisture content. Different letters indicate significant differences between values (ANOVA, Bonferroni-Dunn test, P<0.05). Values are mean ± SEM of three groups of 10 individuals.
23 Fig. 1
20
15 C) o 10
5 Temperature ( Temperature
0
-5 Jan May Sep Jan '05 '06
24 Fig. 2
a 2.8
-0.5 A H (g content Water a 2.6 Water content -1.0 2.4 C)
o b
-1.5 2.2 c
2.0 2
O g -2.0 d Temperature ( 1.8 -1 DM) d -2.5 Ambient temperature 1.6 -3.0 1.4 02468101214
-0.5 a a Melting B point -1.0
C) b o
-1.5 c
-2.0 d Temperature ( Temperature e -2.5 Ambient temperature -3.0 02468101214 Time (days)
25 Fig. 3
0.0 2.8 -1 0.80 g H2O g dry soil
2.6 content (g H Water -0.5 Frozen 2.4 -1.0 C) o 2.2 -1.5 2.0 2
-2.0 1.8 0 g -1 Temperature (
Dehydrated DM) 1.6 -2.5 Ambient temperature 1.4 -3.0 1.2 0 5 10 15 0.0 2.8 -1 1.10 g H2O g dry soil
2.6 (g H content Water -0.5 Frozen 2.4 -1.0 C) o 2.2 -1.5 2.0 2
-2.0 1.8 O g Temperature ( Dehydrated -1 1.6 DM) -2.5 1.4 -3.0 1.2 0 5 10 15 0.0 2.8 -1 1.40 g H2O g dry soil
2.6 H (g content Water -0.5 Frozen 2.4 -1.0 C) o 2.2 -1.5 2.0 2
-2.0 1.8 O g Temperature ( Dehydrated -1 1.6 DM) -2.5 1.4 -3.0 1.2 0 5 10 15 Time (days)
26 Fig. 4
100 b b 80
60 a
40 Frozen (%)
20
0 0.80 1.10 1.40 -1 Soil moisture (g H2O g DM)
27 Chapter 3
Desiccation tolerance and drought acclimation in the Antarctic collembolan Cryptopygus antarcticus
28 Summary The availability of water is recognized as the most important determinant of the distribution and activity of terrestrial organisms within the maritime Antarctic. Within this environment, arthropods may be challenged by drought stress during both the austral summer, due to increased temperature, wind, insolation, and extended periods of reduced precipitation, and the winter, as a result of vapor pressure gradients between the surrounding icy environment and the body fluids. The purpose of the present study was to assess the desiccation tolerance of the Antarctic springtail, Cryptopygus antarcticus, under ecologically-relevant conditions characteristic of both summer and winter along the Antarctic Peninsula. In addition, this study examined the physiological changes and effects of mild drought acclimation on the subsequent desiccation tolerance of C. antarcticus. The collembolans possessed little resistance to water loss under dry air, as the rate of water loss was >20% h-1 at 0% relative humidity (RH) and 4oC. Even under ecologically-relevant desiccating conditions, the collembolans lost water at all relative humidities below saturation (100% RH). However, slow dehydration at high RH dramatically increased the desiccation tolerance of C. antarcticus, as the springtails tolerated a greater loss of body water. Relative to animals maintained at 100% RH, a mild drought acclimation at 98.2% RH significantly increased subsequent desiccation tolerance. Drought acclimation was accompanied by the synthesis and accumulation of several sugars and polyols that could function to stabilize membranes and proteins during dehydration. Drought acclimation may permit C. antarcticus to maintain activity and thereby allow sufficient time to utilize behavioral strategies to reduce water loss during periods of reduced moisture availability. The springtails were also susceptible to desiccation at subzero temperatures in equilibrium with the vapor pressure of ice; they lost ~40% of their total body water over 28 d when cooled to -3.0oC. The concentration of solutes in the remaining body fluids as a result of dehydration, together with the synthesis of several osmolytes, dramatically increased the body fluid osmotic pressure. This increase corresponded to a depression of the melting point to approx. -2.2oC, and may therefore allow C. antarcticus to survive much of the Antarctic winter in a cryoprotectively dehydrated state.
Keywords: desiccation, drought acclimation, Collembola, cold-hardiness, Antarctica, cryoprotective dehydration
29 Introduction The abundance and activity of many soil-dwelling organisms depends upon the moisture characteristics of their environment. This is especially true in the Antarctic where water availability, even more so than temperature, is recognized as the most important determinant of the distribution of Antarctic terrestrial organisms (Kennedy, 1993). During winter, habitat moisture is likely to be limited, as water is biologically unavailable in the form of ice. Similarly, during summer terrestrial microhabitats may dry depending on the vagaries of precipitation, wind, temperature, and insolation in relation to soil and vegetation type (Kennedy, 1993). Therefore, desiccation resistance and/or tolerance of varying relative humidity conditions are likely to be as important as cold tolerance for the survival of terrestrial organisms in polar environments (Ring and Danks, 1994; Block, 1996). Collembolans are among the most abundant and widespread terrestrial arthropods and have been classified into three groups based upon their response to desiccating conditions (Vannier, 1983). Type II (mesophilic) and III (xerophilic) species show increasing control, through morphological, behavioral, and/or physiological means, over water loss when exposed to a desiccating environment. Because of their small size, and correspondingly high surface-to- volume ratio, and highly permeable cuticle, Type I species (hygrophilic) show little to no control over water loss. However, most previous studies assessing rates of water loss and the survival of springtails under desiccating conditions were conducted at extremely low relative humidities, which often were not ecologically relevant. Such experimental treatments may mask mechanisms involved in desiccation resistance/tolerance under a more ecologically-relevant setting. Traditionally, studies on the desiccation physiology of springtails have focused on the role of the cuticle as a barrier for water loss (Verhoef and Witteveen, 1980; Block et al., 1990; Harrison et al., 1991), while desiccation-induced osmolyte production has also received considerable attention (Verhoef, 1981; Verhoef and Prast, 1989; Bayley and Holmstrup, 1999; Holmstrup et al., 2001; Kaersgaard et al., 2004). Recently, a drought acclimation response that increases the tolerance of springtails to subsequent desiccation and cold shock was documented (Sjursen et al., 2001; Bayley et al., 2001; Holmstrup et al., 2002b). Drought acclimation is facilitated by the accumulation of sugars and polyols, which reduce the gradient for water loss and likely stabilize membranes and proteins during subsequent desiccation.
30 In polar and temperate regions, over-wintering arthropods within the soil column possessing a limited resistance to water loss, such as many species of Collembola, may also be challenged by dehydration. As temperatures decline and the soil water freezes, a vapor pressure gradient is established between the unfrozen body fluids of the arthropods and the surrounding frozen environment. Dehydration occurs because the vapor pressure of the environment is lower than that of the unfrozen body fluids at the same temperature. During such dehydration, the melting point of the body fluids may be depressed, as solutes become concentrated in the remaining body fluids and equilibrate with the surrounding environmental temperature, thereby eliminating any risk of freezing (Holmstrup et al., 2002a). Several arthropods are now known to have the capacity to use such cryoprotective dehydration as a viable strategy for winter survival (Holmstrup et al., 2002a; Holmstrup and Sømme, 1998; Worland and Block, 2003; Elnitsky et al., 2008). The springtail Cryptopygus antarcticus is the most widespread and abundant terrestrial arthropod in the Maritime Antarctic (Block, 1984). Large aggregations are found on the underside of rocks, beneath mats of terrestrial algae (i.e., Prasiola crispa), and in association with mosses (Worland and Block, 1986). Previous studies have focused on the desiccation resistance/tolerance of C. antarcticus, suggesting that this species has a limited resistance to desiccation and no physiological control over water loss (Block et al., 1990; Harrison et al., 1991; Block and Harrison, 1995). These studies exposed C. antarcticus to extremely low, constant relative humidity (RH) environments (i.e., 0-35% RH). However, Worland and Block (1986) reported diurnal ranges of 37-100% RH under stones in springtail microhabitats. As yet, no study has assessed the response of C. antarcticus under these more ecologically-relevant conditions of dehydration characteristic of the austral summer. Similarly, Worland and Block (2003) demonstrated that C. antarcticus is susceptible to desiccation at subzero temperatures corresponding to conditions during winter on the Antarctic Peninsula, but did not assess the physiological response to such dehydration. Therefore, the purpose of the present study was to assess the desiccation tolerance and physiological response to dehydration of C. antarcticus under ecologically-relevant conditions characteristic of both the austral summer and winter. In addition, this study examined the physiological changes and effects of drought acclimation on the subsequent desiccation tolerance of C. antarcticus.
31
Materials and methods Source of animals Cryptopygus antarcticus (Willem) (Collembola, Isotomidae) were collected from sites on Humble and Torgersen Islands, near Palmer Station on the Antarctic Peninsula (64o46’ S, 64o04’ W) during January and February 2007. Collembola were subsequently stored at 100% RH and 4oC (0L: 24D) for a minimum of 2 d to ensure animals were fully hydrated prior to experimental use. Only adult C. antarcticus (>1.0 mm long and typically >50 µg; Worland and Block, 2003) were used in all experiments.
Water balance and desiccation tolerance The desiccation tolerance and changes in water content of C. antarcticus were assessed at 4oC at several constant relative humidities. Specific relative humidities were produced in glass desiccators containing 500 ml NaCl solutions. The air inside the closed system quickly equilibrated with the salt solution (following Raoult’s law) to create a 98.2 (31.60 g NaCl L-1), 96.0 (71.20 g NaCl L-1), 93.0 (126.57 g NaCl L-1), or 75.0% (supersaturated NaCl solution) RH environment. Control animals were maintained at 100% RH (double-distilled water). For tests of desiccation tolerance, groups of 10 C. antarcticus were placed within mesh-covered cages (20 µm mesh size), which allowed the free movement of water vapor, and transferred to the experimental relative humidity. Thirty individuals for measurement of total body water content (WC) and five groups of 10 C. antarcticus for assessment of survival were removed daily from each RH treatment. The WC of individual springtails was assessed gravimetrically from measurements (to the nearest 1 µg; Cahn C-31 electrobalance, Ventron Co., Cerritos, CA, USA) of fresh weight at the time of sampling and dry mass (DM) after drying to constant mass at 65oC. Prior to assessing survival, C. antarcticus were rehydrated for 24 h at 100% RH. Animals were considered to have survived if they displayed coordinated walking behavior following rehydration. For comparison to previous studies of water balance in arthropods, the rate of water loss at 0% RH (dry calcium sulfate) and 4oC was also determined by fitting measurements taken at 0.5 h intervals to Wharton’s (1985) model for exponential water loss -kt mt = m0e or ln(mt/m0) = -kt
32 where m0 is the initial water mass, mt is the water mass at time t, and kt is the amount of water
lost between the measurements m0 and mt. The slope of ln(mt/m0) plotted against time is the water loss rate expressed as percent of the total water content per hour. The water mass of individual springtails was assessed gravimetrically as the difference between the mass at time t and DM values.
Drought acclimation To determine the effect of a mild drought stress on the subsequent tolerance of desiccation, C. antarcticus were drought acclimated at 98.2 or 75.0% RH prior to tests of desiccation tolerance. The springtails were drought acclimated at 4oC and 98.2% RH for 24, 48, or 96 h, or at 75.0% RH for 24 h, in glass desiccators as described above. A control group of C. antarcticus was maintained at 100% RH prior to tests of desiccation tolerance. Springtails were subsequently transferred to 96.0 or 93.0% RH for 5 d prior to assessment of survival as described above. During drought acclimation, samples were also removed to assess the role of sugar and polyol accumulation. Glycerol, trehalose, and glucose were measured in C. antarcticus drought acclimated at 98.2% RH for 24, 48, and 96 h, or 75.0% RH for 24 h, as described above. A control group was maintained at 100% RH prior to assessment of sugar and polyol content. Six groups of ~500 animals were weighed and immediately frozen at -80°C until whole body concentrations of sugars and polyols were determined. Animals were subsequently homogenized in 1 N perchloric acid and neutralized with an equal volume of 1 N potassium hydroxide prior to determining sugar and polyol content. Glycerol concentration was determined enzymatically as described by Holmstrup et al. (1999). Trehalose content was determined following digestion with trehalase as described by Chen et al. (2002). Glucose concentration was determined using a glucose oxidase procedure (no. 510; Sigma, St. Louis, MO, USA).
Subzero temperature-induced desiccation Specimens were also desiccated by exposure to an environment at equilibrium with the vapor pressure of ice while monitoring changes in WC and the osmotic pressure of the body fluids. Groups of ~30 springtails were placed within 0.6 ml microcentrifuge tubes and confined by means of fine plastic mesh (20 µm mesh size) which allowed the free movement of water
33 vapor. Microcentrifuge tubes were in turn placed within 10-ml glass vials containing ~4 g of crushed ice and closed with tightly fitting lids. Vials containing C. antarcticus were allowed to equilibrate in refrigerated baths at -0.6oC for 24 h. The temperature of the bath was subsequently lowered incrementally (~0.5oC d-1) to -3.0oC and remained at this temperature until termination of the experiment (day 28). A reference group of C. antarcticus was held at -0.6oC throughout the experiment. Samples were removed at 2 to 7-d intervals for assessment of WC, as described above, and osmotic pressure of the body fluids. Determinations of the body fluid osmotic pressure were made using a vapor pressure depression technique (Holmstrup and Sømme, 1998). Groups of ~30 C. antarcticus were placed in a sample holder and quickly crushed with a Teflon rod to expose the body fluids. Samples were then allowed to equilibrate for 30 min following placement within a C-52 sample chamber (Wescor Inc., Logan, UT, USA). The osmotic pressure of the body fluids was measured using a Wescor HR-33T Dew Point Microvoltmeter (Wescor Inc., Logan, UT, USA) operated in the dew point mode. Additionally, sugar and polyol accumulation during subzero temperature-induced desiccation was examined. On days 0, 6, 15, and 28 during exposure to -3.0oC in an environment at equilibrium with the vapor pressure of ice, groups of C. antarcticus were removed and stored for subsequent analysis of glycerol, trehalose, and glucose as described above. Control springtails were maintained at -0.6oC until termination of the experiment.
Statistical analysis Variations in survival, WC, and DM during dehydration under constant RH environments were analyzed with two-way (treatment x time with their interaction) analysis of variance (ANOVA) following tests of parametric assumptions. When there were significant treatment effects, Student-Newman-Keuls (SNK) multiple comparison procedure was used to test for significant differences over time. Differences in survival of desiccation between control and drought-acclimated Collembola were compared with Student’s t-tests. Mean sugar and polyol concentrations and changes in WC and osmotic pressure during subzero temperature induced desiccation were analyzed with one-way ANOVA and Bonferroni-Dunn tests. Percentage data were arcsin-square root transformed prior to analysis. Data not meeting parametric assumptions were log transformed to correct for non-normality or heteroscedasticity. All data are presented as mean ± SEM with statistical significance set at P<0.05.
34
Results Water content and desiccation tolerance When tested at 4oC, C.antarcticus lost water at all relative humidities except at water saturation (100% RH; Fig. 1A). Within a relative humidity treatment, the DM of C. antarcticus did not differ significantly during the course of the experiment, suggesting that differences in fresh mass were due solely to water loss. The WC of live Collembola declined significantly (P<0.0001) at all relative humidities below water saturation through day 5, prior to leveling throughout the remainder of the experiment. Rates of water loss from C. antarcticus were inversely related to the desiccation treatment (i.e., higher rates of water loss occurred at lower relative humidities). By day 10 of desiccation, only the 98.2% RH treatment had a sufficient number of surviving collembola to allow accurate measurement of WC. The WC of C. antarcticus maintained at 100% RH did not change during the course of the experiment. The survival of C. antarcticus during desiccation declined as a function of WC (Fig. 1B). -1 At 96.0, 93.0, and 75.0% RH, survival dropped rapidly as WC dropped below 1.1 g H2O g DM , -1 and only at 96.0% RH did >50% of the adults survive at a WC below 0.95 g H2O g DM . This contrasts with C. antarcticus exposed to 98.2% RH, of which >75% of individuals survived at a -1 WC below 0.90 g H2O g DM . These results suggest that the desiccation tolerance of C. antarcticus is highly dependent upon the severity of the desiccation stress and rate of dehydration; desiccation tolerance was increased at lower rates of water loss. At the termination of the experiment (day 10), the osmotic pressure of body fluids in springtails exposed to 98.2% RH had equilibrated to that of the environment (approx. -25 bar; N = 5) and, therefore, they would not have been expected to lose more water. Springtails displayed little resistance to water loss in dry air (0% RH). Applying Wharton’s (1985) model, the hourly rate of water loss at 0% RH and 4oC of C. antarcticus was ~21% h-1 (Fig. 2). Under such extreme conditions of desiccation, survival dropped rapidly, as -1 fewer than 50% of the individuals survived body water contents below ~1.1 g H2O g DM (Fig. 1B), corresponding to only 1.5 h of desiccation at 0% RH.
Drought acclimation
35 Mild drought stress at 98.2% RH significantly increased the subsequent tolerance of desiccation of C. antarcticus to both 96.0 and 93.0% RH (Fig. 3A, B). The survival of springtails acclimated at 100% RH (controls) was ~55 and <20%, respectively, when exposed to 96.0 or 93.0% RH for 5 d. Drought acclimation at 98.2% RH reduced the body water content (Table 1); WC was reduced by nearly 45% in the 96 h acclimation. Relative to the controls, acclimation at 98.2% RH for 48 or 96 h significantly (P<0.001) increased subsequent survival of exposure to either 96.0 or 93.0% RH for 5 d. Acclimation at 98.2% RH for 24 h had no significant effect on subsequent desiccation tolerance. Drought acclimation at 75.0% RH for 24 h similarly reduced WC by ~35% (Table 1); however, it had no significant effect on the subsequent survival of C. antarcticus at 96.0 or 93.0% RH (Fig. 3A, B). Insufficient numbers of collembola survived longer drought acclimations at 75.0% RH; therefore, their subsequent tolerance of dehydration was not tested. Drought acclimation at 98.2% RH was accompanied by the de novo synthesis and accumulation of several osmolytes within the body fluids (Table 1). Glycerol concentrations were increased more than 5-fold to >45 µg mg DM-1 after a 96 h acclimation at 98.2% RH. Glucose and trehalose concentrations were more modestly elevated by ~2 and 3-fold, respectively, following acclimation for 48 or 96 h. Acclimation at 75.0% RH resulted in a small, but significant (P<0.001), increase in glycerol concentration, but failed to induce the synthesis of glucose or trehalose.
Subzero temperature-induced desiccation Slow cooling of C. antarcticus to -3.0oC in an environment at equilibrium with the vapor pressure of ice resulted in a significant (P<0.0001) reduction of WC (Fig. 4A). The WC decreased through day 14, prior to leveling off over the remainder of the experiment. By day 28, -1 the WC had been reduced by ~40% to 1.41 ± 0.03 g H2O g DM . The DM of C. antarcticus did not change significantly over the course of the experiment, suggesting all mass changes were due solely to water loss. At the termination of the experiment collembolan survival was nearly 75% (N = 150). Control C. antarcticus equilibrated to -0.6oC had a body water content of 2.36 ± 0.05 -1 g H2O g DM , with no significant change in WC or DM during the experiment. Similarly, at day 28 survival of control animals was >81% (N = 150).
36 The osmotic pressure of the body fluids of C. antarcticus equilibrated to -0.6oC was -9.4 ± 0.4 bar (N = 6) and did not change significantly in control animals during the course of the experiment. However, cooling at equilibrium with the vapor pressure of ice resulted in a significant (P<0.0001) increase of the hemolymph osmotic pressure (Fig. 4B). Relative to controls, the osmotic pressure of C. antarcticus exposed to the vapor pressure of ice increased nearly 3-fold, to -26.4 ± 0.3 bar (N = 6) by day 28. However, at termination of the experiment there remained a vapor pressure deficit of approximately -10 bar between the collembolans and the surrounding environment. Therefore, the springtails would have likely continued to lose water to their environment. Water loss alone could not account for the observed increase in the osmotic pressure of the body fluids during cooling at equilibrium with the vapor pressure of ice. Desiccation at subzero temperatures also induced the de novo synthesis of several osmolytes (Table 2). During desiccation, the concentrations of trehalose and glucose increased significantly (P<0.0001) by ~4 and 5-fold, respectively, by day 28 of the subzero exposure. At termination of the experiment, glycerol concentration had increased ~8-fold to nearly 70 µg mg DM-1. Such osmolyte synthesis likely contributed substantially toward the observed increase in the osmotic pressure of the body fluids, especially at low body water contents. At termination of the experiment, glucose and trehalose concentrations were unchanged in control individuals maintained at -0.6oC, whereas glycerol concentration increased significantly (P<0.001) by ~2-fold.
Discussion Long-term field monitoring of C. antarcticus populations on maritime Antarctic Signy Island suggests that springtails are faced with drought stress during both the austral summer and winter (Block and Convey, 2001; Convey et al., 2003). Monthly sampling over an 11-year period from 1984-1995 revealed a distinct seasonal pattern of body water content in C. antarcticus; WC was lowest during mid-summer (December-January) and mid-winter (June- August). During summer, increased solar radiation, temperature, wind, and periods of reduced precipitation contribute to the drying of microhabitat sites. In winter, when ice is present within the soil matrix, springtails may lose water to the surrounding environment due to the lower vapor pressure of ice compared to the unfrozen body fluids (Holmstrup et al., 2002a). Such conditions necessitate an inherent tolerance of desiccation. The present study sought to characterize the
37 desiccation tolerance of C. antarcticus under ecologically-relevant conditions characteristic of both summer and winter along the Antarctic Peninsula, and also to identify physiological mechanisms that may reduce water loss during drought stress and/or increase the inherent tolerance for dehydration. These findings are discussed in the context of our current knowledge regarding the physiology and ecology of this species.
Desiccation tolerance Cryptopygus antarcticus showed little or no resistance to desiccation under dry-air conditions, thus confirming the results from previous investigations (Worland and Block, 1986; Block et al., 1990; Harrisson et al., 1991). At 4oC and 0% RH, the collembolans lost ~20% of their total body water per hour and succumbed to dehydration stress within two hours. This high rate of water loss is comparable to that of other hygrophilic collembolans desiccated under conditions of dry air (Vannier, 1983). However, while useful for comparative purposes in assessing the absolute rates of water loss, such extreme conditions of desiccation are likely of little ecological relevance. Further, these tests of desiccation resistance may mask processes that operate to increase the tolerance and/or resistance to dehydration under more ecologically- relevant conditions. Within their microhabitat, C. antarcticus rarely, if ever, experiences conditions below ~35% RH (Worland and Block, 1986) and when presented with drought stress within their natural environment, they may employ behavioral [e.g., the tendency to aggregate in large numbers or actively seek moist refuges (Hayward et al., 2001, 2004)], and/or physiological mechanisms [e.g., through the accumulation of osmolytes (present study)] to limit water loss. Due to their high surface-to-volume ratio and limited cuticular resistance (Harrisson et al., 1991) to water loss, C. antarcticus are especially susceptible to dehydration. This was evident even at high relative humidities, as the springtails lost water under all conditions below saturation, with the rate of water loss inversely proportional to the desiccation treatment. No evidence of water vapor absorption from subsaturated air was observed, as has been reported for some other species of Collembola (Bayley and Holmstrup, 1999). However, springtails maintained at 98.2% RH for 10 d had equilibrated the osmotic pressure of their body fluids to that of the environment (approx. -25 bar). These individuals would, therefore, not be expected to undergo further reductions of body water. At this time survival was >75% (compared to ~90%
38 for control springtails maintained at 100% RH), suggesting C. antarcticus can tolerate extended periods in the absence of free water at high relative humidities. Under mild desiccation stress at high relative humidities, corresponding to lower rates of water loss, C. antarcticus tolerated a greater loss of body water. When desiccated at 0% RH, -1 survival declined to below 50% at ~1.15 g H2O g DM , corresponding to a loss of only ~50% of the total body water. However, at 93.0% RH, springtails survived the loss of ~58% of their body water and >60% at the highest relative humidity tested (98.2%). Such increases in desiccation tolerance at lower rates of water loss are well known amongst arthropods (Hadley, 1994). Similarly in the maritime Antarctic, Benoit and colleagues (2007) recently reported that the Antarctic midge, Belgica antarctica, tolerates a significantly greater loss of body water when dehydrated at higher relative humidities. However, the mechanisms accounting for this increased tolerance of dehydration are unknown. It has been suggested that desiccation at lower rates of water loss permits adjustments of the water stores, in an attempt to maintain sufficient tissue hydration and preserve cellular metabolic activity, by shifting water from the extracellular compartment to the cells (Hadley, 1994). Slow dehydration of both C. antarcticus (present study) and B. antarctica (Benoit et al., 2007) also stimulated the synthesis and accumulation of higher concentrations of several sugars and polyols. In addition to further reducing the gradient for water loss, even modest concentrations of these osmolytes are well known to stabilize and protect membranes and proteins thereby preventing/reducing dehydration-induced cellular damage (Crowe et al., 1992; Yancey, 2005).
Drought acclimation increases desiccation tolerance Mild drought acclimation increased the desiccation tolerance of C. antarcticus. Acclimation for 48 or 96 h at 98.2% RH increased the subsequent survival of springtails at either 96.0 or 93.0% RH for 5 d. Whereas analogous thermal acclimations are well known to confer increased tolerance to high or low temperature, drought acclimation is less well studied. However, increases of dehydration tolerance following acclimation are known from nematodes (Crowe and Madin, 1975), reptiles (Lillywhite, 2004), insects (Hoffmann, 1990; Benoit et al., 2007), and other collembolan species (Sjursen et al., 2001). For example, when the soil-dwelling springtail Folsomia candida was similarly acclimated at 98.2% RH for 6 d, survival of subsequent drought stress, as low as 94% RH, and the tolerance of water loss was dramatically
39 increased (Sjursen et al., 2001). Among other Antarctic organisms, workers recently reported a similar drought acclimation response in the midge B. antarctica (Benoit et al., 2007). The reduction of the total body water content during drought acclimation of C. antarcticus was accompanied by the synthesis and accumulation of several osmolytes. This contrasts with previous investigations that suggested this species does not accumulate osmolytes during dehydration (Worland and Block, 1986; Block and Harrisson, 1995). The reason for these differing results is unknown. However, in previous studies dehydration likely occurred rapidly, as springtails were exposed to extremely low relative humidity environments. Our results suggest that C. antarcticus can accumulate moderate concentrations of osmolytes if desiccation occurs at low rates of water loss and high RH. This point is exemplified by the finding that springtails accumulated significantly higher concentrations of osmolytes when desiccated to a similar extent at 98.2% RH (96 h) compared to those desiccated at 75.0% RH (24 h). A similar pattern is observed in B. antarctica, in which midge larvae accumulate greater osmolyte concentrations when desiccated at high relative humidities (Benoit et al., 2007). The accumulation of osmolytes appears to be a common component to the drought acclimation response and likely contributes mechanistically, by protecting membranes and proteins, to the increased tolerance of dehydration (Sjursen et al., 2001; Benoit et al., 2007; present study). In F. candida, drought acclimation also results in a higher degree of unsaturation of membrane phospholipid fatty acids (Bayley et al., 2001; Holmstrup et al., 2002b), a change that resembles membrane alterations seen in ectothermic animals acclimated to low temperature. Holmstrup and colleagues (2002b) suggest such membrane desaturation may counter the increased packing of membrane lipids that occurs as water is removed from the cell during dehydration, thereby maintaining membrane fluidity and metabolic function. Whether similar changes in the membrane unsaturation occur during and contribute to the drought acclimation observed in C. antarcticus deserves investigation. In the maritime Antarctic, the availability of water is widely recognized as the most important determinant of the distribution and activity of terrestrial organisms (Kennedy, 1993). The ecological significance of the enhanced desiccation tolerance of C. antarcticus may allow springtails to make use of microhabitats prone to dehydration. Further, Sjursen et al. (2001) suggested that drought acclimation likely permits springtails to maintain activity in the face of increasing environmental drought stress. This may be especially important as microhabitats
40 continue to dry, allowing C. antarcticus to use behavioral mechanisms, such as aggregation or dispersal to moist refuges, to reduce further water loss.
Subzero temperature induced desiccation Cryptopygus antarcticus is believed to over-winter within air spaces in the upper soil layers under vegetation, rocks and stones (Convey et al., 2003). The springtails are, therefore, vulnerable to desiccation at subzero temperatures during winter as the vapor pressure of the surrounding environment is lower than that of the unfrozen body fluids at the same temperature. Our results confirm previous reports (Worland and Block, 2003) and demonstrate that C. antarcticus may lose a considerable portion of their total body water when exposed to the vapor pressure of ice at subzero temperatures. In the present study, springtails lost ~40% of their total body water over the course of 28 d at -3.0oC. Similar to their response to drought acclimation, desiccation at subzero temperatures was accompanied by the synthesis and accumulation of several osmolytes. Long-term monitoring of field populations suggests C. antarcticus is regularly challenged by drought stress during winter as WC may be reduced by 25% or more to -1 as low as 1.2 g H2O g DM (Block and Harrisson, 1995; Convey et al., 2003). Additionally, over-wintering field populations are known to accumulate significant concentrations of various osmolytes, including glycerol, glucose, and trehalose (Montiel, 1998). Taken together, it appears that partial dehydration and the accumulation of several sugars and polyols form an important component to the over-wintering strategy of C. antarcticus. Desiccation at subzero temperatures and the resulting concentration of solutes in the remaining body fluids, together with the de novo synthesis of osmolytes, dramatically increased the osmotic pressure of C. antarcticus to approx. -25 bar by day 28 at -3.0oC. Such an increase in the osmotic pressure corresponds to a ~2.2oC depression of the melting point of the body fluids. At least near Palmer Station, Antarctica where springtail microhabitat temperatures remain between -1 and -3oC throughout much of the austral winter (Elnitsky et al., 2008), due to oceanic buffering, C. antarcticus may be able to remain in a cryoprotectively-dehydrated state. Even if temperatures subsequently decline forcing reliance upon supercooling for survival, such dehydration would likely only increase the supercooling capacity of the springtails and reduce the risk of lethal freezing (Sømme and Block, 1982; Worland and Block, 2003).
41 Conclusions On the Antarctic Peninsula, C. antarcticus may be challenged by drought stress throughout the year. In addition to behavioral strategies, our results suggest that the springtails also rely upon a number of physiological mechanisms to limit water loss and/or increase desiccation tolerance. When presented with water stress, the selection of moist microhabitat sites would not only reduce the rate of water loss, but may enhance desiccation tolerance through the synthesis and accumulation of various organic osmolytes. Additionally, the drought acclimation response may permit C. antarcticus to maintain metabolic and cellular activity, thereby extending survival during subsequent desiccation stress. During winter, the collembolans likely rely upon the partial dehydration and accumulation of sugars and polyols, similar to the Arctic collembolan Onychiurus arcticus (Worland et al., 1998), as important components of their over-wintering strategy. These physiological responses and their effects on activity and survival likely contribute to the success of C. antarcticus in this harsh environment.
Acknowledgements This research was supported by NSF grants OPP-0337656 and OPP-0413786. We are especially thankful to the staff at Palmer Station for their support and assistance in Antarctica.
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43 Hadley, N. F. (1994). Water Relations of Terrestrial Arthropods. Academic Press, New York, 356 pp. Harrisson, P. M., Rothery, P., and Block, W. (1991). Drying processes in the Antarctic collembolan Cryptopygus antarcticus (Willem). J. Insect Physiol. 37, 883-890. Hayward, S. A. L., Bale, J. S., Worland, M. R., and Convey, P. (2001). Influence of temperature on the hygropreference of the Collembolan, Cryptopygus antarcticus, and the mite, Alaskozetes antarcticus from the maritime Antarctic. J. Insect Physiol. 47, 11- 18. Hayward, S. A. L., Worland, M. R., Convey, P., and Bale, J. S. (2004). Habitat moisture availability and the local distribution of the Antarctic Collembola Cryptopygus antarcticus and Frisea grisea. Soil Biol. Biochem. 36, 927-934. Hoffmann, A. (1990). Acclimation for desiccation resistance in Drosophila melanogaster and the association between acclimation responses and genetic variation. J. Insect Physiol. 36, 885-891. Holmstrup, M. and Sømme, L. (1998). Dehydration and cold hardiness in the Arctic collembolan Onychiurus arcticus Tullberg 1876. J. Comp. Physiol. B 168, 197-203. Holmstrup, M., Bayley, M., and Ramløv, H. (2002a). Supercool or dehydrate? An experimental analysis of overwintering strategies in small permeable Arctic invertebrates. Proc. Nat. Acad. Science USA 99, 5716-5720. Holmstrup, M., Costanzo, J. P., and Lee, R. E. (1999). Cryoprotective and osmotic responses to cold acclimation and freezing in freeze-tolerant and freeze-intolerant earthworms. J. Comp. Physiol. B 169, 207-214. Holmstrup, M., Hedlund, K., and Boriss, H. (2002b). Drought acclimation and lipid composition in Folsomia candida: implication for cold shock, heat shock, and acute desiccation stress. J. Insect Physiol. 48, 961-970. Holmstrup, M., Sjursen, H., Ravn, H., and Bayley, M. (2001). Dehydration tolerance and water vapor absorption in two species of soil-dwelling Collembola by accumulation of sugars and polyols. Funct. Ecol. 15, 647-653. Kaersgaard, C. W., Holmstrup, M., Malte, H., and Bayley, M. (2004). The importance of cuticular permeability, osmolyte production and body size for the desiccation resistance of nine species of Collembola. J. Insect Physiol. 50, 5-15.
44 Kennedy, A. D. (1993). Water as a limiting factor in the Antarctic terrestrial environment: a biogeographical synthesis. Arctic Alpine Res. 25, 308-315. Lillywhite, H. B. (2004). Plasticity of the water barrier in vertebrate integument. Inter.Congress Series 1275, 283-290. Montiel, P. O. (1998). Profiles of soluble carbohydrates and their adaptive role in maritime Antarctic terrestrial arthropods. Polar Biol. 19, 250-256. Ring, R. and Danks, H. (1994). Desiccation and cryoprotection: overlapping adaptations. Cryo- Letters 15, 181-190. Sjursen, H., Bayley, M., and Holmstrup, M. (2001). Enhanced drought tolerance of a soil- dwelling springtail by pre-acclimation to a mild drought stress. J. Insect Physiol. 47, 1021-1027. Sømme, L. and Block, W. (1982). Cold hardiness of Collembola at Signy Island, maritime Antarctic. Oikos 38, 168-176. Vannier, G. (1983). The importance of ecophysiology for both biotic and abiotic studies of soil. In: Leburn, P., Andre, H. M., deMets, A., Gregoire-Wibo, C., Wauthy, G. (Eds.), New Trends in Soil Biology. Dieu-Brichart, Ottignies-Louvain-La-Neuve, Belgium, pp. 289- 314. Verhoef, H. A. (1981). Water balance in Collembola and its relation to habitat selection: water content, haemolymph osmotic pressure and transpiration during an instar. J. Insect Physiol. 27, 755-760. Verhoef, H. A. and Prast, J. E. (1989). Effects of dehydration on osmotic and ionic regulation in Orchesella cincta (L.) and Tomocerus minor (Lubbock) (Collembola) and the role of the coelomoduct kidneys. Comp. Biochem. Physiol. 93A, 691-694. Verhoef, H. A. and Witteveen, J. (1980). Water balance in Collembola and relation to habitat selection: cuticular water loss and water uptake. J. Insect Physiol. 26, 201-208. Wharton, G. W. (1985). Water balance of insects. In: Kerkut, G.A., Gilbert, L.I. (Eds.), Comprehensive Insect Physiology, Biochemistry and Pharmacology, vol. 4. Pergamon Press, Oxford, pp. 565-603. Worland, M. R. and Block, W. (1986). Survival and water loss in some Antarctic arthropods. J. Insect Physiol. 32, 579-584.
45 Worland, M. R. and Block, W. (2003). Desiccation at sub-zero temperatures in polar terrestrial arthropods. J. Insect Physiol. 49, 193-203. Worland, M. R., Grubor-Lajsic, G., and Montiel, P. O. (1998). Partial desiccation induced by sub-zero temperatures as a component of the survival strategy of the Arctic collembolan Onychiurus arcticus (Tullberg). J. Insect Physiol. 44, 211-219. Yancey, P. H. (2005). Organic osmolytes as compatible, metabolic and counteracting cytoprotectants in high osmolarity and other stresses. J.Exp. Biol. 208, 2819-2830.
46 Table 1. The total body water content (N = 25-30) and osmolyte concentrations (N = 6) of Cryptopygus antarcticus during drought acclimation at 4oC and 98.2 or 75.0% RH. Values are mean ± SEM. Different letters denote significant differences between treatment groups (ANOVA; Bonferroni-Dunn test).
Hours of exposure
98.2% RH 75.0% RH
0 24 48 96 24
-1 a b c d e Water content (g H20 g DM ) 2.27 ± 0.03 1.96 ± 0.06 1.67 ± 0.04 1.26 ± 0.05 1.42 ± 0.06
Osmolyte concentration (µg mg DM-1)
Glycerol 9.2 ± 1.2a 19.7 ± 2.6b 31.3 ± 3.1c 47.4 ± 5.2d 13.8 ± 0.9b
Glucose 5.6 ± 0.9a 6.2 ± 1.2a 12.3 ± 1.1b 19.5 ± 2.2c 7.8 ± 1.4a
Trehalose 4.1 ± 0.7a 6.4 ± 1.1a 10.6 ± 1.4b 18.6 ± 1.9c 6.8 ± 1.5a
47 Table 2. Osmolyte concentrations (N = 6) of Cryptopygus antarcticus during exposure at -3.0oC in an environment at equilibrium with the vapor pressure of ice. Values are mean ± SEM. Different letters denote significant differences between days of exposure (ANOVA; Bonferroni-Dunn test).
Days of exposure
Desiccation at -3.0oC Control (-0.6oC)
0 6 15 28 28
Osmolyte concentration (µg mg DM-1)
Glycerol 8.7 ± 1.3a 15.3 ± 2.2b 27.6 ± 2.7c 68.4 ± 5.6d 16.5 ± 2.0b
Glucose 5.4 ± 0.8a 8.3 ± 1.1a 16.7 ± 2.1b 26.8 ± 3.3c 7.8 ± 1.2a
Trehalose 3.8 ± 0.7a 7.1 ± 0.9a 10.1 ± 1.6b 16.2 ± 1.4c 6.1 ± 1.1a
48 Figure legends Fig. 1. (A) Changes in total body water content of Cryptopygus antarcticus during desiccation exposure within various relative humidity (RH) environments at 4oC. Values are mean ± SEM of 25-30 individuals. (B) Percent survival as a function of total body water content of C. antarcticus during desiccation in various constant relative humidity environments. Values are mean ± SEM of five groups of 10 individuals.
Fig. 2. Water loss rate of Cryptopygus antarcticus at 4oC and 0% RH. A linear regression line- of-best fit described the equation y = -0.208x – 0.00650 (R2 = 0.998), where the slope of the regression represents the water loss rate in percent of total body water per hour. Values are mean ± SEM of 25-30 individuals.
Fig. 3. Survival of Cryptopygus antarcticus desiccated for 5 d at either 96.0% RH (A) or 93.0% RH (B) at 4oC. Collembola were previously acclimated at 100% RH (control) or drought acclimated at 98.2 or 75.0% RH prior to assessment of desiccation tolerance. Values are mean ± SEM of five groups of 10 individuals. Asterisks denote a significant difference relative to the control treatment (Student’s t-test).
Fig. 4. Changes in (A) body water content (N = 15-20 individuals) and (B) osmotic pressure of the body fluids (N = 6) of Cryptopygus antarcticus during slow cooling to -3.0oC in an environment at equilibrium with the vapor pressure of ice. Values are mean ± SEM.
49 Fig. 1.
A ) -1
2.0 100% RH O g DM
2 98.2% RH 96.0% RH 1.5 93.0% RH 75.0% RH
1.0 Water content (g H 0.5 0 48 96 144 192 240
Hours of exposure
B 100
75
98.2% RH 50 96.0% RH 93.0% RH 75.0% RH Survival (%) 0% RH 25
0 0.4 0.6 0.8 1.0 1.2 1.4 1.6 1.8 2.0 2.2 -1 Water content (g H2O g DM)
50 Fig. 2.
0.1
0.0
-0.1 ) 0 -0.2 /m t -0.3 ln(m
-0.4
-0.5
-0.6 0.0 0.5 1.0 1.5 2.0 2.5
Time (h)
51 Fig. 3.
A. 96.0% RH
100 * 80 *
60
40 Survival (%)
Control - 100% RH 20 Drought acclimation - 98.2% RH Drought acclimation - 75.0% RH
0 024487296
B. 93.0% RH 100
80
60 * * 40 Survival (%) Survival
20
0 0 24487296 Acclimation time (h)
52 Fig. 4.
A
2.4 -0.5 Water content (g H
2.2 -1.0 C) o 2.0 -1.5
1.8 -2.0 2 O g Water 1.6
Temperature ( Temperature -2.5 Acclimation content -1
DM) temperature 1.4 -3.0
1.2 0 5 10 15 20 25 30
B
-30 -0.5 Osmotic pressure (bar) pressure Osmotic
-1.0 Osmotic -25
C) pressure o -1.5 -20 -2.0
-15 Temperature ( -2.5 Acclimation temperature -3.0 -10
0 5 10 15 20 25 30 Time (d)
53 Chapter 4
Osmoregulation and salinity tolerance in the Antarctic midge, Belgica antarctica: seawater acclimation confers cross tolerance to freezing and dehydration
54 Summary Summer storms along the Antarctic Peninsula can cause microhabitat sites of the terrestrial midge Belgica antarctica to become periodically inundated with seawater from tidal spray. As microhabitats dry, larvae may be exposed to increasing concentrations of seawater. Alternatively, as a result of melting snow or following rain, larvae may be immersed in freshwater for extended periods. The present study assessed the tolerance and physiological response of B. antarctica larvae to salinity exposure, and examined the effect of seawater acclimation on their subsequent tolerance of freezing, dehydration, and heat shock. Midge larvae tolerated extended exposure to hyperosmotic seawater; nearly 50% of larvae survived a 10-d exposure in 1000 mOsm kg-1 seawater and ~25% of larvae survived 6 d in 2000 mOsm kg-1 seawater. Exposure to seawater drastically reduced body water content and increased hemolymph osmolality. In contrast, larvae were effective osmoregulators at seawater concentrations less than the osmotic concentration of the body fluids (~400 mOsm kg-1). Hyperosmotic seawater exposure resulted in a significant correlation between the rate of oxygen consumption and larval water content and induced the de novo synthesis and accumulation of several organic osmolytes. A 3-d acclimation to hyperosmotic seawater increased freezing tolerance relative to freshwater-acclimated larvae. Even after rehydration, the freezing survival of larvae acclimated in seawater was greater than freshwater-acclimated larvae. Additionally, seawater acclimation increased larvae’s subsequent tolerance of dehydration. Our results further illustrate the similarities between these related, yet distinct forms of osmotic stress, and add to the suite of physiological responses used by larvae to enhance survival in the harsh and unpredictable Antarctic environment. Keywords: Osmotic stress, dehydration, cold-hardiness, Antarctica, Chironomidae
55 Introduction Insects tolerant of seawater submergence can be classified as either osmoregulators or osmoconformers (Bradley, 1987). During exposure to high salinity, osmoregulators maintain the hemolymph at a concentration hyposmotic to the external medium. This strategy is common among dipteran larvae, including saline-water mosquitoes of the genus Aedes (for review see Bradley, 1987) and chironomids (Neumann, 1976; Kokkin, 1986). Saline-water larvae constantly lose water to their environment, and must counter such water loss by actively ‘drinking’ the external medium. To prevent a substantial accumulation of ions that may be detrimental to protein function (Somero and Yancey, 1997), larvae must also excrete a concentrated ‘urine.’ Alternatively, osmoconformers maintain water balance by equilibrating the osmotic pressure of the body fluids with that of the saline environment. Indeed, other genera of mosquitoes (Culex, Garrett and Bradley, 1987; Culiseta, Deinocerites, Bradley, 1994) and insect orders that contain euryhaline species (e.g., Chironomus sp., a dragonfly nymph Enallagma clausam, Stobbart and Shaw, 1974) osmoconform while inhabiting saline environments. Garrett and Bradley (1987) demonstrated that larvae of the mosquito Culex tarsalis osmoconform by accumulating high concentrations of proline and trehalose. It is thought that accumulation of these organic osmolytes, as opposed to inorganic ions (predominantly Na+ and Cl-), preserves the function of enzymes despite an increase in osmolality (Somero and Yancey, 1997). Most insects do not tolerate prolonged seawater submergence, and at least for most larval dipterans, the upper limit of salinity tolerance is equal to the initial osmotic concentration of the hemolymph (Bayly, 1972). Exposure to hyperosmotic saline results in an inability to osmo- and ionoregulate. However, physiological responses to salinity exposure that ameliorate this hyperosmotic and ionic stress (e.g., accumulation of organic osmolytes, reduced rate of water loss, etc.) may extend survival time. Belgica antarctica Jacobs (Diptera: Chironomidae) is the southern-most free-living holometabolous insect, being sporadically dispersed, but locally abundant, on the west coast of the Antarctic Peninsula. Detailed accounts of the life-history and ecology of this terrestrial, Antarctic midge are provided by Convey and Block (1996), Sugg et al. (1983), Usher and Edwards (1984), and references cited therein. Briefly, its two-year life cycle includes four larval stages and over-wintering may occur in any instar. Larvae are freeze tolerant to ca. -15oC (Baust and Lee, 1981; Lee et al., 2006), and are extremely tolerant of desiccation; larvae survive the loss
56 of ~70% of their total body water (Benoit et al., 2007). During freezing and desiccation, larvae are necessarily challenged with extensive osmotic stress, as solutes become concentrated in the remaining extracellular body water. Such tolerance suggests the larvae have a well developed ability to maintain a critical minimum volume necessary for cell survival and the maintenance of metabolic function when faced with osmotic stress. During summer, Antarctic storms can result in terrestrial microhabitats of B. antarctica becoming periodically inundated with seawater from tidal spray (Baust and Lee, 1987; M. Elnitsky, personal observation). Subsequently, as microhabitats dry due to evaporation, larvae may be exposed to increasing concentrations of seawater. On the other hand, as a result of melting snow or following rain, larvae may be immersed in freshwater for extended periods. Therefore, the physiological tolerance and response of larvae to such osmotic perturbations are likely critical for survival. A previous study of B. antarctica found >95% survival of larvae following a 7-d submergence in 0.5 M NaCl (Baust and Lee, 1987). However, the osmotic response to such salinity and the tolerance of more severe hyperosmotic stress is unknown. Therefore, the purpose of the present study was to assess the salinity tolerance and acute osmotic response of larval B. antarctica, including changes in water content, body fluid osmotic pressure, and osmolyte accumulation during salinity stress. Additionally, we characterized the physiological effects of seawater exposure on the rate of oxygen consumption and investigated the effect of a brief seawater acclimation on the subsequent tolerance of freezing, desiccation, and heat shock.
Materials and methods Source of insects Substrate containing larval B. antarctica was collected from sites near penguin rookeries on Cormorant and Humble Islands, near Palmer Station on the Antarctic Peninsula (64o46’ S, 64o04’ W) in January and February 2006 and 2007. Samples were stored at 4oC (0L: 24D) in moist native substrate prior to use. Larvae were then handpicked from the substrate on ice-cold water and held in freshwater at 4oC for 12-24 h to ensure clearance of the gut (mean gut clearance ~6 h; Baust and Edwards, 1979) and to standardize body water content prior to use. Only 4th instar larvae were used for all experiments.
57 Osmoregulation and salinity tolerance To assess the osmotic response and salinity tolerance of B. antarctica, larvae were exposed to various concentrations of seawater. The salinity tolerance of larvae was tested during exposure to hypoosmotic (0 mOsm kg-1; freshwater), isoosmotic (~400 mOsm kg-1), and hyperosmotic (~1000 [pure seawater], ~1500, and ~2000 mOsm kg-1) solutions of seawater. The desired seawater solutions were produced by either dilution or concentration, via evaporation, of pure seawater collected adjacent to Palmer Station. Groups of 10 larvae were placed in open- top, 1.6-ml microcentrifuge tubes and ~1 ml of the test seawater solution added. Larvae were subsequently placed at 4oC and samples removed on days 0, 1, 3, 6 and 10 for assessment of survival and measurement of total body water content (WC) and hemolymph osmolality. Five groups of 10 larvae were removed for survival assessment and individuals displaying spontaneous movement were deemed to have survived. The WC of individual larvae was assessed gravimetrically from measurements (to the nearest 1 µg; Cahn C-31 electrobalance, Ventron Co., Cerritos, CA, USA) of fresh weight at the time of sampling and dry mass (DM) after drying to constant mass at 65oC. Osmolality determinations were made using a vapor pressure depression technique (Holmstrup and Sømme, 1998). Groups of 5 larvae were placed in a sample holder and quickly crushed with a Teflon rod to expose the body fluids. Samples were then allowed to equilibrate for 30 min following placement within a C-52 sample chamber (Wescor Inc., Logan, UT, USA). The osmolality of the sample was measured using a Wescor HR-33T Dew Point Microvoltmeter (Wescor Inc., Logan, UT, USA) operated in the dew point mode. Only surviving larvae were used for measurement of WC and osmolality.
Osmolyte analysis The role of osmolyte accumulation during osmotic stress was also examined. Glycerol, trehalose, and glucose analyses were performed on larvae following exposure to seawater (~1000 mOsm kg-1) at 4oC. A group of control larvae was maintained in freshwater (0 mOsm kg-1) at 4oC until termination of the experiment (day 6). On days 1, 3, and 6, groups of ~25 larvae were weighed and immediately frozen at -80°C until whole body concentrations of osmolytes were determined. Larvae were homogenized in 1 N perchloric acid and neutralized with equal volumes of 1 N potassium hydroxide prior to determining osmolyte content. Glycerol concentrations were determined enzymatically as described by Holmstrup et al. (1999).
58 Trehalose content was determined following digestion with trehalase as described by Chen et al. (2002). Glucose concentration was determined using the glucose oxidase procedure (no. 510; Sigma, St. Louis, MO, USA).
Oxygen consumption Recently, Benoit et al. (2007) reported a reduced rate of oxygen consumption of B. antarctica larvae following desiccation. Therefore, the effect of osmotic stress, and the accompanying changes in WC, on the rate of oxygen consumption of the midge larvae was assessed following acclimation to seawater (~1000 mOsm kg-1) at 4oC. Groups of 10 larvae were acclimated in open-top, 1.6-ml microcentrifuge tubes containing ~1.0 ml of either seawater or freshwater. Daily, five groups of 10 larvae were removed for measurement of oxygen consumption via closed-system respirometry using an Instech Fiber Optic Oxygen Monitor (Model 110; Instech Laboratories, Inc., Plymouth Meeting, PA, USA). Larvae were placed in the FOXY chamber (Instech Laboratories, Inc., Plymouth Meeting, PA, USA) along with 1.0 ml of the acclimation solution. Larvae were allowed to equilibrate within the chamber for ~10 min, prior to recording changes in dissolved oxygen concentration for the subsequent 12 min using OOISensors Software (Instech Laboratories, Inc., Plymouth Meeting, PA, USA). The rate of oxygen consumption per unit time was calculated using the slope calculator on the OOISensors Software. At the end of each trial, larvae were removed from the chamber, blotted dry, and weighed (to nearest 0.01 mg). The WC of larvae was determined following drying to constant o -1 -1 weight at 65 C. Oxygen consumption was expressed as µl O2 g DM h . Prior to all measurements, the oxygen monitoring system was calibrated using solutions of 0% oxygen concentration (200 µM sodium hydrosulfite) and a saturated oxygen solution at 4oC.
Cross tolerance to freezing, heat shock, and desiccation Hayward and colleagues (2007) recently demonstrated that a mild desiccation stress conferred cross tolerance to freezing in B. antarctica larvae. Therefore, we assessed the effect of a mild osmotic stress on the subsequent tolerance of freezing, heat shock, and desiccation by acclimating larvae for 3 d to seawater (~1000 mOsm kg-1) at 4oC prior to tests of environmental stress tolerance. In addition, to test whether seawater acclimation conferred increased cross tolerance independent of reductions in body water content, a group of larvae was acclimated in
59 seawater for 3 d followed by a 24-h rehydration in freshwater prior to assessment of stress tolerance. A control group of larvae was maintained in freshwater (0 mOsm kg-1) at 4oC for 3 d prior to measurement of stress tolerance. All larvae were acclimated to either seawater or freshwater as described above. Immediately prior to tests of freeze tolerance, larvae were removed from their respective treatment and placed in microcentrifuge tubes containing 100 µl of fresh water (to standardize the amount of ice formation within the freezing media). Larvae were subsequently placed directly at the target temperature (-10, -12, -15, or -20oC) and allowed to freeze for 6 h. As larvae have a limited resistance to inoculative freezing (Elnitsky et al., 2008), the freezing of the body fluids likely occurred at a high subzero temperature, but was not controlled or monitored. Prior to survival assessment, larvae were allowed to thaw/recover for 24 h at 4oC. For tests of heat shock tolerance, larvae were placed directly at 30oC and five groups of 10 larvae removed at 0.5 h intervals to assess survival. Prior to survival assessment, larvae were permitted to recover for 24 h at 4oC. Finally, for tests of desiccation tolerance, groups of 5 larvae were removed from their respective treatment, blotted dry and placed within mesh-covered cages (20 µm mesh size), which allowed the free movement of water vapor. Cages containing larvae were in turn placed on a dry platform within glass desiccators containing 500 ml NaCl solutions. The air inside the closed system quickly equilibrated with the salt solution (following Raoult’s Law) to create a relative humidity (RH) at either 98.2 (31.6 g NaCl l-1) or 75.0% (supersaturated NaCl solution). At 1-2 d intervals, 20 individuals were removed for assessment of WC and five groups of 10 individuals removed for assessment of survival. Water content was measured gravimetrically as described above. Larvae were allowed to rehydrate for 24 h at 100% RH at 4oC before survival was assessed. For all tests of cross-tolerance to other environmental stressors, individuals displaying spontaneous movement were deemed to have survived.
Statistical analysis Changes in survival, WC, and osmolality over the course of the saline or freshwater exposure were analyzed with two-way (treatment x time along with their interaction) analysis of variance (ANOVA) following a test of parametric assumptions. Where there were significant treatment effects, the Student-Newman-Keuls comparison was used to test for significant differences over time. Linear regression analysis was used to evaluate the relationship between
60 the rate of oxygen consumption during acclimation to seawater and WC. Mean sugar and polyol concentrations of larvae were compared with one-way ANOVA and Bonferroni-Dunn tests. Survival following tests of cross tolerance to other environmental stressors was analyzed relative to the freshwater (control) acclimation with two-way ANOVA and Dunnett’s test. Survival data were arcsin-square root transformed prior to analysis. Data not meeting parametric assumptions were log transformed to correct for non-normality or heteroscedasticity. All data are presented as mean ± 1 SEM with statistical significance set at P<0.05.
Results Osmoregulation and salinity tolerance The survival of larval B. antarctica exposed to solutions of hyperosmotic seawater was significantly (P<0.0001) affected by both the time of exposure and the strength of the seawater solution (Fig. 1A); survival declined with submergence time and the concentration of seawater. Still, greater than 75% of larvae survived exposure in pure seawater (~1000 mOsm kg-1) for 6 d whereas ~45% survived to the termination of the experiment (day 10). Even after 6 d, >50 and 25% of larvae survived exposure in 1500 and 2000 mOsm kg-1 seawater, respectively. Nearly all larvae exposed to freshwater or isoosmotic seawater (400 mOsm kg-1) survived to day 10 of exposure. Exposure of larvae to hyperosmotic seawater resulted in significant (P<0.05) reductions of WC (Fig. 1B). The WC of larvae exposed to 1000 mOsm kg-1 seawater declined significantly through day 6 prior to stabilizing over the remainder of the experiment. By day 10, the WC of -1 -1 1000 mOsm kg exposed larvae was reduced by ~30% to 1.78 ± 0.05 g H2O g DM . Similarly, by day 6, the WC of larvae exposed to 1500 and 2000 mOsm kg-1 seawater was significantly reduced by ~32 and 43%, respectively. The WC of larvae exposed to 400 mOsm kg-1 seawater varied slightly, but significantly (P<0.05), throughout the experiment, but by day 10 did not differ from control larvae maintained in freshwater. Freshwater submergence did not affect larval WC over the course of the experiment. The reduction of WC during exposure of larvae to hyperosmotic seawater solutions necessarily resulted in significant (P<0.0001) increases in hemolymph osmolality over the course of the experiment (Fig. 1C). The osmolality of larvae exposed to 1500 and 2000 mOsm kg-1 seawater increased by ~2.2 and 2.3-fold, respectively, by day 6. Similarly, the hemolymph
61 osmolality of larvae exposed to 1000 mOsm kg-1 seawater nearly doubled by day 6 prior to leveling off throughout the remainder of the experiment. The hemolymph osmolality of larvae exposed to isoosmotic seawater also increased slightly, but significantly, over the course of the exposure. Submergence in freshwater did not affect hemolymph osmolality over the 10 d exposure.
Osmolyte accumulation The reductions of WC as a result of exposure to hyperosmotic seawater were insufficient to account for the observed increase in hemolymph osmolality (Table 1). As osmolyte synthesis in B. antarctica larvae has been documented in response to dehydration (Benoit et al., 2007), we measured the concentrations of several osmolytes during osmotic stress. Exposure to hyperosmotic seawater induced the de novo synthesis of osmolytes in larval B. antarctica. Glucose and trehalose concentrations increased approximately three-fold over the course of a 6-d exposure in 1000 mOsm kg-1 seawater. Similarly, by day 6 the concentration of glycerol increased more than four-fold to nearly 10 µg mg DM-1. The osmolyte concentration of control larvae maintained in freshwater was not affected over the course of the experiment.
Oxygen consumption Acclimation to seawater resulted in a significant correlation (P<0.001) between the rate of oxygen consumption and larval WC (Fig. 2). In general, lower rates of oxygen consumption were found in larvae having a lower WC. The rate of oxygen consumption ranged from 0.32 – 0.68 µl g DM-1 h-1 and the linear regression model suggested that larvae having the lowest WC had rates of oxygen consumption that were ~50% below fully hydrated larvae.
Cross tolerance to other environmental stressors Relative to the freshwater controls, a 3-d acclimation in 1000 mOsm kg-1 seawater significantly (P<0.0001) increased the freeze tolerance of B. antarctica larvae (Fig. 3). Nearly all larvae survived a 6-h freeze at -10oC. However, of larvae acclimated 3 d in freshwater less than 65% survived freezing at -12oC and <15% survived a 6-h freeze at -15oC. In contrast, nearly 95 and 55% of 1000 mOsm kg-1 seawater acclimated larvae survived freezing at -12 and - 15oC, respectively. Even when frozen at -20oC, nearly 15% of seawater-acclimated larvae
62 survived. This contrasts with freshwater acclimation, in which no larvae survived a 6-h freeze at -20oC. Additionally, the freeze tolerance of larvae rehydrated for 24 h, during which the WC of the larvae was restored to pre-acclimation levels, following seawater acclimation was significantly (P<0.05) greater than freshwater-acclimated larvae (Fig. 3). Relative to freshwater- acclimated larvae, seawater acclimation followed by rehydration increased survival by 18 and 22% following freezing at -12 and -15oC, respectively. Similarly, acclimation to 1000 mOsm kg-1 seawater significantly (P<0.001) affected the subsequent desiccation tolerance of B. antarctica, however, survival as a function of time was increased only in larvae rehydrated prior to desiccation (Fig. 4A, B). At both 98.2 and 75.0% RH, the survival of rehydrated larvae was significantly higher than the control, freshwater- acclimated larvae. The desiccation tolerance of larvae acclimated 3 d in seawater did not differ relative to freshwater-acclimated larvae. However, this result may have been simply due to the -1 reduced WC of the seawater-acclimated larvae (~2.0 vs. ~2.5 g H2O g DM for freshwater- acclimated larvae). This is supported by the observation that at any given WC during desiccation at 75.0% RH the survival of both the seawater-acclimated and rehydrated groups was greater than that of the freshwater-acclimated larvae (i.e., seawater-acclimated larvae tolerated a greater loss of body water during dehydration). The increased survival of freezing and desiccation following seawater acclimation, however, was contrary to the response of the larvae to heat shock (Fig. 5). Survival of heat shock at 30oC declined rapidly with exposure time, such that less than 75 and 25% of freshwater- acclimated larvae survived a 1.5 and 2.5 h heat shock, respectively. However, relative to freshwater acclimation, the survival of seawater-acclimated larvae was significantly (P<0.001) lower at all exposure times. The survival of larvae rehydrated following the 3-d seawater acclimation did not differ significantly (P>0.05) from that of larvae acclimated in freshwater.
Discussion Larval B. antarctica must endure potentially severe osmotic perturbation, as they are regularly challenged by a variety of environmental conditions that generate hydric and osmotic stress. On the Antarctic Peninsula, the freeze-tolerant larvae may be faced with subzero temperatures throughout the year. Freezing of the body fluids results in a cellular dehydration, whereby water is osmotically drawn from intracellular stores to the now concentrated
63 extracellular fluids (for a review, see Lee, 1991). Similarly, larvae may be exposed to severely desiccating conditions as microhabitats dry due to the vagaries of summer precipitation, elevated temperature, wind, and insolation. Desiccation may also occur during winter, as freezing of the surrounding soil solution establishes an osmotic gradient for water loss from the yet unfrozen body fluids of the larvae (Elnitsky et al., 2008). Since microhabitat sites may be occasionally immersed in increasing concentrations of seawater from Antarctic storms, we investigated the tolerance and physiological response of the larvae to hyperosmotic seawater exposure. Together with previous investigations (Baust and Edwards, 1979; Baust and Lee, 1987; Hayward et al., 2007; Benoit et al., 2007; Michaud et al., 2008), our results demonstrate the impressive tolerance of the midge larvae to variation in environmental salinity. This inherent tolerance is facilitated by a variety of physiological responses (e.g., osmolyte accumulation, etc.) induced by the osmotic challenge, that likely also contribute to the observed cross-tolerance to freezing and dehydration. These results are discussed below in the context of our current knowledge of this and other species’ tolerance and response to osmotic stress.
Tolerance and physiological response to salinity Terrestrial larvae of B. antarctica tolerated extensive osmotic dehydration when challenged by hyperosmotic seawater. Nearly 50% of the larvae survived a 10-d exposure in 1000 mOsm kg-1 seawater during which time the total body water content of the larvae was -1 reduced by ~30% to <1.80 g H2O g DM . Survival declined rapidly during exposure to higher seawater concentrations; however, even in 2000 mOsm kg-1 seawater (i.e., twice the concentration of pure seawater), ~25% of larvae survived a 6-d exposure. As the larvae are known to tolerate an extensive loss of body water, the survival of hyperosmotic seawater is likely not limited solely by the larvae’s tolerance to dehydration. During desiccation in air, larvae -1 survive the loss of nearly 70% of their body water to <1.0 g H2O g DM (Benoit et al., 2007). Instead, it seems likely that an incurred salt load from the external medium contributed to the observed mortality during seawater exposure. Inorganic ions, and especially Na+ and Cl-, are well known to disrupt cellular activity by binding to and destabilizing proteins and nucleic acids (Somero and Yancey, 1997; Hochachka and Somero, 2002; Yancey, 2005). Together with the reduced body water content, such salt load likely results in a breakdown of cellular homeostasis that may limit salinity tolerance in B. antarctica.
64 The larvae were unable to effectively osmoregulate during exposure to hyperosmotic seawater. The reduced water content of the larvae, and associated concentration of solutes in the remaining body fluids, necessarily contributed to the increased osmotic concentration (Table 1). However, water loss alone was insufficient to account for the observed increase in hemolymph osmolality. At day 6 of exposure to 1000 mOsm kg-1 seawater, the reduction of body water could explain only ~70% of the observed osmotic concentration of the body fluids. Salinity exposure also induced the de novo synthesis and accumulation of several organic osmolytes, as glycerol, glucose and trehalose concentrations all increased 3 to >4-fold over the 6-d exposure. Assuming that the organic osmolytes were dissolved in the osmotically active fraction of the body water (Worland et al., 1998), such osmolyte accumulation likely contributed ~100 mOsm kg-1 (~13%) toward the observed osmolality at day 6 during exposure to seawater (Table 1). The larvae likely also incurred a significant accumulation of inorganic ions from the seawater medium that contributed to the osmolality; however, we can not rule out the additional accumulation of other organic osmolytes. The accumulation of organic osmolytes is a well known response used by organisms to reduce osmotic stress (Hochachka and Somero, 2002). These compatible organic solutes are generally favored over the accumulation of inorganic ions because they limit the perturbation of macromolecules even at high concentrations (Yancey, 2001). In addition to their colligative effect in reducing the gradient for water loss, many organic solutes also have other physiological properties, such as stabilizing membranes and proteins during cell shrinkage (Crowe et al., 1984; Crowe et al., 1992; Sano et al., 1999), which preserve cellular function. Among arthropods, larvae of the euryhaline mosquito C. tarsalis accumulate high concentrations of proline and trehalose in response to increased environmental salinity (Garrett and Bradley, 1987; Patrick and Bradley, 2000). We previously demonstrated that larval B. antarctica accumulate significant concentrations of glycerol and trehalose in response to desiccation stress at high relative humidities in air (Benoit et al., 2007), and glucose and trehalose at subzero temperatures in the presence of environmental ice (Elnitsky et al., 2008). In the present study, we observed a similar response to hyperosmotic seawater exposure. In all cases, the accumulation of organic osmolytes likely serves to slow and/or limit cellular dehydration while preserving metabolic function. This may be especially important during exposure to hyperosmotic seawater to extend
65 survival time and allow the larvae to use other behavioral mechanisms to reduce further osmotic stress. At seawater concentrations less than the osmotic concentration of the body fluids (~400 mOsm kg-1) the larvae were effective at osmoregulation. During submergence in freshwater (0 mOsm kg-1), the larvae maintained the total body water content and osmotic concentration of the -1 -1 body fluids at ~2.5 g H2O g DM and ~400 mOsm kg , respectively. In isoosmotic seawater (400 mOsm kg-1), hemolymph osmolality increased slightly (~10%) over the 10-d exposure. This increase could not be accounted for by a simple concentration effect due to changes in WC, as by day 10 of the exposure the WC did not differ from that of larvae maintained in freshwater. Therefore, it suggests these larvae too incurred an increased salt load from the external medium and/or the salinity exposure resulted in the accumulation of organic osmolytes.
Reduced O2 consumption during osmotic stress Exposure to hyperosmotic seawater and the accompanying osmotic dehydration resulted in a significant positive correlation between the rate of oxygen consumption and the total body water content of B. antarctica larvae. Whether the reduction in the rate of oxygen consumption was compensated for by an increase in anaerobic metabolism is unknown. However, we previously reported a similar reduction of aerobic metabolism of larval B. antarctica following dehydration in air (Benoit et al., 2007). Such hypometabolism in response to dehydration is well known among arthropods and has been suggested as a mechanism to reduce respiratory water loss (Hadley, 1994). In our study, the reduced rate of oxygen consumption would not be expected to curtail water loss during submergence in hyperosmotic seawater. Osmoregulating mosquito larvae typically display increased rates of oxygen consumption during exposure to increased environmental salinity, presumably in an effort to ionoregulate (Bradley, 1987). However, in species unable to effectively osmoregulate during hyperosmotic seawater exposure, rates of oxygen consumption generally decline with increased environmental salinity and salt load within the body fluids (Bradley, 1987; Plaut, 1999; Ordiano et al., 2005; Irwin et al., 2007). Together with the present results, this suggests that rather than an active physiological down regulation of metabolism in an effort to limit water loss, the reduced rate of oxygen consumption may be a passive response to osmotic and ionic perturbations of cellular function.
66 Osmotic dehydration, and any additional salt load from the external medium, incurred during the exposure to hyperosmotic seawater may have resulted in the observed hypometabolism due to effects on protein function. In addition to the aforementioned effects of inorganic ions on protein stability, solute crowding, as result of reduced body water, can also impose osmophobic and hydrophobic effects on macromolecules (Hochachka and Somero, 2002). Such osmo- and hydrophobicity have important effects on the assembly and folding of proteins and nucleic acid structures and, therefore, could result in reduced protein function and metabolic rate in the dehydrated state. Additionally, we cannot rule out a role for the accumulation of organic solutes (e.g., urea; Muir et al., 2007) contributing to the observed reduction of oxygen consumption.
Cross tolerance to other environmental stressors Acclimation to hyperosmotic seawater (1000 mOsm kg-1) increased larval tolerance to freezing and dehydration, but reduced tolerance of high temperature. A link between cold tolerance and dehydration is now well established (Ring and Danks, 1994; 1998). At the cellular level, freezing and dehydration present a similar challenge of maintaining membrane integrity as water is osmotically removed from cellular stores. Further, in many cases the physiological response(s) elicited by organisms, such as the accumulation of osmolytes, to these related stressors is similar. Among arthropods, enhanced cold tolerance following dehydration is known from several taxa (Hadley, 1994). Similar to our present results, Hayward and colleagues (2006) recently documented enhanced freeze tolerance of B. antarctica following slow dehydration in air. The simple reduction of body water content would be expected to slow and reduce ice formation within the body fluids (Lee et al., 1991), and likely contributed to the enhanced survival of subsequent freezing. However, even after larvae were rehydrated their subsequent tolerance of freezing was increased, suggesting other physiological mechanisms, independent of reductions of body water content, were involved (see below). In contrast to increased cold tolerance following prior hydric stress, enhanced desiccation tolerance is less well documented. However, a drought acclimation response that enhances subsequent tolerance of dehydration and low temperature is known from the soil-dwelling springtail Folsomia candida (Sjursen et al., 2001). When this collembolan was acclimated at 98.2% RH for 6 d, survival of subsequent drought stress, to as low as 94% RH, and the tolerance
67 of water loss was dramatically increased. We recently documented a similar drought acclimation response, analogous to the enhanced desiccation tolerance following acclimation in hyperosmotic seawater, in B. antarctica following desiccation at high relative humidities (Benoit et al., 2007). Accumulation of organic solutes appears to be a common component of the drought acclimation response (Sjursen et al., 2001; Benoit et al., 2007; present study), and likely contributes mechanistically, by protecting membranes and proteins, to the increased tolerance of dehydration. In F. candida, drought acclimation also results in a higher degree of unsaturation of membrane phospholipid fatty acids (Bayley et al., 2001; Holmstrup et al., 2002), a change that resembles membrane alterations seen in ectothermic animals acclimated to low temperature. Holmstrup and colleagues (2002) suggest such membrane desaturation may counter the increased packing of membrane lipids that occurs as water is removed from the cell during dehydration, thereby maintaining membrane fluidity and metabolic function. Such changes in B. antarctica during acclimation to seawater could help to explain their enhanced tolerance of desiccation and low temperature. Additionally, desaturation of phospholipid fatty acids may render the larvae more susceptible to heat shock, as cellular membranes may be more prone to phase transitions at high temperatures resulting in a loss of membrane integrity. It is also worth noting that as with B. antarctica larvae, F. candida experiences reduced tolerance to high temperatures following drought acclimation (Holmstrup et al., 2002).
Conclusions In the harsh and highly variable environment of the Antarctic Peninsula, B. antarctica may be regularly challenged by osmotic stress, including freezing, dehydration, and periodic submergence in hyperosmotic seawater. It has become apparent that the terrestrial larvae possess an impressive tolerance and rely upon a number of behavioral and physiological mechanisms to ameliorate to such stress. During hyperosmotic seawater submergence, the inherent tolerance of osmotic stress likely allows the larvae to maintain metabolic function and escape to more favorable microhabitats. However, prolonged submergence in seawater presents a severe threat to survival, likely as a result of water loss and salt load from the external environment. Cellular water loss may be slowed and protein structure and function preserved by the accumulation of several organic osmolytes. These osmolytes likely also contribute to the enhanced subsequent
68 tolerance of freezing and desiccation following a brief seawater acclimation, and reflect a conserved physiological response to these related yet distinct forms of osmotic stress.
Acknowledgements This research was supported by NSF grants OPP-0337656, OPP-0413786, and IAB-0416750. We are especially thankful to the support staff at Palmer Station for their assistance in Antarctica. We also wish to thank Tim Muir and Jon Costanzo for their assistance with collection of the oxygen consumption data.
69 References
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70 Elnitsky, M. A., Hayward, S. A. L., Rinehart, J. P., Denlinger, D. L., and Lee, R. E., Jr. (2008). Cryoprotective dehydration and the resistance to inoculative freezing in the Antarctic midge Belgica antarctica. J. Exp. Biol. 211, 524-530. Garrett, M. A. and Bradley, T. J. (1987). Extracellular accumulation of proline, serine and trehalose in the haemolymph of osmoconforming brackish-water mosquitoes. J. Exp. Biol. 129, 231-238. Hayward, S. A. L., Rinehart, J. P., Sandro, L. H., Lee, R. E., and Denlinger, D. L. (2007). Slow dehydration promotes desiccation and freeze tolerance in the Antarctic midge Belgica antarctica. J. Exp. Biol. 210, 836-844. Hadley, N. F. (1994). Water Relations of Terrestrial Arthropods. Academic Press, San Diego. Hochachka, P. W. and Somero, G. N. (2002). Biochemical adaptation. Oxford University Press, New York. Holmstrup, M., Costanzo, J. P., and Lee, R. E. (1999). Cryoprotective and osmotic responses to cold acclimation and freezing in freeze-tolerant and freeze-intolerant earthworms. J. Comp. Physiol. B 169, 207-214. Holmstrup, M., Hedlund, K., and Boriss, H. (2002). Drought acclimation and lipid composition in Folsomia candida: implications for cold shock, heat shock and acute desiccation stress. J. Insect Physiol. 48, 961-970. Holmstrup, M. and Sømme, L. (1998). Dehydration and cold hardiness in the Arctic collembolan Onychiurus arcticus Tullberg 1876. J. Comp. Physiol. B 168, 197-203. Irwin, S., Wall, V., and Davenport, J. (2007). Measurement of temperature and salinity effects on oxygen consumption of Artemia franciscana K., measured using fibre-optic oxygen microsensors. Hydrobiol. 575, 109-115. Kokkinn, M. J. (1986). Osmoregulation, salinity tolerance and the site of ion excretion in the halobiont chironomid, Tanytarsus baritarsis Freeman. Austral. J. Mar. Fresh. Res. 37, 243-250. Lee, R. E. (1991). Principles of insect low temperature tolerance. In: Insects at Low Temperature (ed. R. E. Lee and D. L. Denlinger), pp. 17-46. New York: Chapman and Hall.
71 Lee, R. E., Elnitsky, M. A., Rinehart, J. P., Hayward, S. A. L., Sandro, L. H., and Denlinger, D. L. (2006). Rapid cold-hardening increases the freezing tolerance of the Antarctic midge, Belgica antarctica. J. Exp. Biol. 209, 399-406. Michaud, M. R., Benoit, J. B., Lopez-Martinez, G., Elnitsky, M. A., Lee, R. E., and Denlinger, D. L. (2008). Metabolomics reveals unique and shared metabolic changes in response to heat shock, freezing, and desiccation in the Antarctic midge, Belgica antarctica. J. Insect Physiol. 54, 645-655. Muir, T. J., Costanzo, J. P., and Lee, R. E. (2007). Osmotic and metabolic responses to dehydration and urea-loading in a dormant, terrestrially hibernating frog. J. Comp. Physiol. B 177, 917-926. Neumann, D. (1976). Adaptations of chironomids to intertidal environments. Annu. Rev. Entomol. 21, 387-414. Ordiano, A., Alvarez, F., and Alcaraz, G. (2005). Osmoregulation and oxygen consumption of the hololimnetic prawn, Macrobrachium tuxtlaense at varying salinities (Decapoda, Palaemonidae). Crustaceana 78, 1013-1022. Patrick, M. L. and Bradley, T. J. (2000). The physiology of salinity tolerance in larvae of two species of Culex mosquitoes: the role of compatible solutes. J. Exp. Biol. 203, 821-830. Plaut, I. (1999). Effects of salinity acclimation on oxygen consumption in the freshwater blenny, Salaria fluviatilis, and the marine peacock blenny, S. pavo. Mar. Fresh. Res. 50, 655-659. Ring, R. A. and Danks, H. V. (1994). Desiccation and cryoprotection: overlapping adaptations. Cryo-Letters 15, 181-190. Ring, R. A. and Danks, H. V. (1998). The role of trehalose in cold-hardiness and desiccation. Cryo-Letters 19, 275-282. Sano, R., Asakawa, N., Inoue, Y., and Sakurai, M. (1999). A dual role for intracellular trehalose in the resistance of yeast cells to water stress. Cryobiology 39, 80-87. Sjursen, H., Bayley, M., and Holmstrup, M. (2001). Enhance drought tolerance of a soil- dwelling springtail by pre-acclimation to a mild drought stress. J. Insect Physiol. 47, 1021-1027. Somero, G. N. and Yancey, P. H. (1997). Osmolytes and cell-volume regulation: physiological and evolutionary principles. In The Handbook of Physiology, section 14, Cell Physiology.
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73 Table 1. Estimated osmotic contribution of initial osmolytes in the hemolymph and osmolytes produced during exposure of Belgica antarctica larvae to hyperosmotic seawater. Values are mean ± SEM. Different letters represent significant differences between days of exposure (ANOVA; Bonferroni-Dunn test).
Days of exposure
Seawater (1000 mOsm kg-1) Control (freshwater)
0 1 3 6 6
Osmolality (mOsm kg-1) 407 ± 7a 491 ± 10b 646 ± 12c 779 ± 15d 391 ± 10a
-1 a b b c a Total body water content (g H2O g DM ) 2.51 ± 0.07 2.17 ± 0.05 2.07 ± 0.03 1.89 ± 0.05 2.46 ± 0.06
Osmotically active water (OAW) 2.04 1.72 1.63 1.46 1.99 -1 content* (g H2O g DM )
Loss of OAW (%) --- 15.7 21.1 28.5 ---
Osmotic contribution of original solutes --- 480 516 569 --- due to loss of OAW (mOsm kg-1) Osmolyte concentration (µg mg DM-1) Glycerol 2.17 ± 0.27a 2.96 ± 0.46a 6.87 ± 0.58b 9.56 ± 0.46c 1.94 ± 0.31a
Glucose 3.75 ± 0.34a 4.71 ± 0.42a 8.30 ± 0.48b 12.68 ± 0.54c 3.82 ± 0.28a
Trehalose 4.91 ± 0.31a 6.62 ± 0.36b 9.11 ± 0.41c 14.38 ± 0.53d 4.23 ± 0.37a
Osmotic contribution of synthesized --- 11 54 108 --- osmolytes† (mOsm)
74
Total explainable osmotic --- ~100 88.2 86.9 --- pressure (%)
* OAW was calculated from Worland et al. (1998). [(OIW) = 0.069(TBW) + 0.3, where OIW is osmotically inactive water content, TBW is total body water content and is the sum of OIW and OAW]. † Assuming that osmolytes are only dissolved in OAW.
75 Figure legends
Fig.1. (A) Survival (N = five groups of 10 larvae), (B) water content (N = 25-30), and (C) hemolymph osmolality (N = 6) of Belgica antarctica larvae exposed to a various concentrations of seawater or to freshwater (~0 mOsm kg-1). Values are mean ± 1 SEM.
Fig. 2. Effect of acclimation to seawater (~1000 mOsm kg-1) and resultant changes of body water content on the rate of oxygen consumption of Belgica antarctica larvae (y = -0.853 + 0.602x; N = 15; R2 = 0.822; P<0.001).
Fig. 3. Effect of acclimation to seawater on the freeze tolerance of Belgica antarctica larvae. Larvae were acclimated to either seawater (~1000 mOsm kg-1) or freshwater (~0 mOsm kg-1) for 3 d prior to assessment of freeze tolerance. A third group of larvae (rehydrated) were acclimated to seawater for 3 d followed by rehydration for 24 h in freshwater. Larvae were frozen in groups of 10 individuals in ~100 µL of freshwater for 6 h. Values are mean ± 1 SEM of five groups of 10 larvae. Asterisks denote a significant difference relative to the freshwater (control) treatment (ANOVA, Dunnett’s test, P<0.05).
Fig. 4. Effect of seawater acclimation on the desiccation tolerance of Belgica antarctica larvae at 98.2 (A) or 75.0% RH (B) and 4oC. Larvae were acclimated to either seawater (~1000 mOsm kg-1) or freshwater (~0 mOsm kg-1) for 3 d prior to assessment of desiccation tolerance. Rehydrated larvae were acclimated to seawater for 3 d and then allowed to rehydrate for 24 h in freshwater prior to desiccation. Values are mean ± 1 SEM of five groups of 10 larvae. Asterisks denote a significant difference relative to the freshwater (control) treatment (ANOVA, Dunnett’s test, P<0.05).
Fig. 5. Effect of seawater acclimation on the heat shock tolerance (time at 30oC) of Belgica antarctica larvae. Larvae were acclimated to either seawater (~1000 mOsm kg-1) or freshwater (~0 mOsm kg-1) for 3 d prior to assessment of heat shock tolerance. Rehydrated larvae were acclimated to seawater for 3 d and then allowed to rehydrate for 24 h in freshwater prior to heat
76 shock. Values are mean ± 1 SEM of five groups of 10 larvae. Asterisks denote a significant difference relative to the freshwater (control) treatment (ANOVA, Dunnett’s test, P<0.05).
77 Fig. 1.
A 100
75
50
Survival (%) 0 mOsm kg-1 acclimation -1 25 400 mOsm kg acclimation 1000 mOsm kg-1 acclimation -1 1500 mOsm kg acclimation 2000 mOsm kg-1 acclimation 0
) B -1 2.6 2.4 O g DM 2 2.2 2.0 1.8 1.6 Water content (g H Water 1.4
1000 C
) 900 -1
800
700
600
500 Osmolality (mOsm kg 400
300 0246810
Time (days)
78 Fig. 2. ) -1 h
-1 0.7 g DM 2 0.6
0.5
0.4
0.3 Oxygen consumption (ul O Oxygen 1.81.92.02.12.22.32.42.52.6 Water content (g H O g DM-1) 2
79 Fig. 3.
* Freshwater acclimation 100 Rehydrated Seawater acclimation
75 * 50 * Survival (%)
25 *
0% 0% 0 -10 -12 -15 -20 o Temperature ( C)
80 Fig. 4.
A 100 *
* 75 * 50
Survival (%) 25 Freshwater acclimation Rehydrated Seawater acclimation
0 0246810
Time at 98.2% RH (d)
B 100 * 75 * 50 Survival (%)
25
0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 Time at 75% RH (d)
81 Fig. 5.
100 Freshwater acclimation Rehydrated Seawater acclimation 75 *
50 *
Survival (%) * 25 * 0 * 0.0 0.5 1.0 1.5 2.0 2.5 3.0 o Time at 30 C (h)
82 Chapter 5 Concluding remarks Terrestrial organisms inhabiting the Antarctic Peninsula may be faced with a myriad of harsh and unpredictable environmental conditions, including extremes of low temperature, limited water availability, and exposure to hyperosmotic seawater, which result in potentially severe osmotic stress. The success of Antarctic arthropods in this environment, therefore, necessitates an inherent tolerance to osmotic challenge. This dissertation sought to assess the tolerance and physiological response of the Antarctic midge Belgica antarctica and the springtail Cryptopygus antarcticus to such osmotic stress. During the austral winter, the freeze-tolerant larvae of the Antarctic midge B. antarctica may be encased within a matrix of soil and ice at subzero temperatures for 7-8 months. If the larvae resist inoculative freezing, these environmental conditions create a substantial gradient for water loss from the unfrozen larvae, owing to the lower vapor pressure of the surrounding ice. Our results demonstrate that B. antarctica do indeed dehydrate when exposed to subzero temperatures at equilibrium with the vapor pressure of ice. This water loss, along with the de novo synthesis and accumulation of several organic osmolytes, depressed the melting point of the larvae’s body fluids to near equilibrium with the ambient temperature, suggesting the larvae have the capacity to undergo cryoprotective dehydration at ecologically-relevant subzero temperatures. While such protective dehydration was previously known from other arthropods, this study represented the first report in a true insect. For B. antarctica larvae in nature, the use of cryoprotective dehydration versus freeze tolerance for winter survival likely depends upon the ambient microenvironmental conditions, especially soil moisture, upon entrance into winter. Monitoring of Cryptopygus antarcticus populations in the maritime Antarctic suggests they too are faced with drought stress. During summer, microhabitats may dry due to increased temperature, wind, and extended periods of reduced precipitation. In addition to behavioral strategies, our results suggest that the springtails also rely upon a number of physiological mechanisms to limit water loss and/or increase the tolerance of desiccation. Under ecologically- relevant conditions the collembolans possess an impressive basal tolerance of water loss. This tolerance was further enhanced through a drought acclimation response involving the synthesis of several organic osmolytes. During winter, the collembolans, similar to larval B. antarctica,
83 likely rely upon the partial dehydration and the accumulation of sugars and polyols as important components of their over-wintering survival. Finally, midge larvae may also be osmotically challenged by periodic submergence in freshwater, from summer rain and melting snow and ice, or hyperosmotic seawater, from splash from Antarctic storms. The larvae readily survived and were effective osmoregulators at osmotic concentrations less than the osmolality of the body fluids. In hyperosmotic seawater, larvae were unable to osmoregulate and survival declined with prolonged submergence. However, the effects of such osmotic perturbation were likely ameliorated again by the accumulation of organic solutes. These osmolytes likely also contributed to the enhanced subsequent tolerance of freezing and dehydration following a brief seawater exposure. Low environmental temperature and freezing of the soil water, dehydration, and hyperosmotic seawater all present these terrestrial arthropods with a similar osmotic stress. In all three cases, the arthropods accumulate organic osmolytes, likely to slow/limit cellular water loss and preserve protein structure and function. The similar physiological mechanisms used by these and other arthropods to ameliorate the effect of water stress likely represents an evolutionarily conserved response to related, yet distinct forms of osmotic perturbation. At least indirectly, this finding also supports the notion that freeze tolerance as a strategy for low temperature survival likely evolved from adaptations to desiccation stress. The studies presented herein further illustrate the impressive tolerance of environmental stress that likely contributes to the success of B. antarctica and C. antarcticus on the Antarctic Peninsula.
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