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Physiology and Biochemistry |
2DuPont Bio-Based Materials, Chestnut Run Plaza 728/1411, P.O. Box 80728, Wilmington, Delaware 19880-0728 USA; 3Halophyte Biotechnology Center, Graduate College of Marine Studies, University of Delaware, 700 Pilottown Road, Lewes, Delaware 19958 USA
Received for publication April 23, 2004. Accepted for publication January 24, 2005.
ABSTRACT
Callus cultures of the salt marsh grass Spartina patens were examined to determine changes and consistencies in membrane lipid composition in response to salt. Major membrane lipid classes remained stable at all salinity levels (0, 170, 340 mmol/L). However, the membrane protein to lipid ratio decreased significantly in response to elevated NaCl. Callus plasma membrane (PM) consisted predominantly of sterols, about 60% (mol%) of the total lipids. Glycolipid was the second largest lipid class, making up about 20% (mol%) of the total. With increasing salinity, the relative percentage of sitosterol decreased, while that of campesterol increased. The phospholipid species detected were phosphatidylethanolamine (PE), phosphatidylcholine (PC), phosphatidylserine (PS), and phosphatidylinositol (PI). When callus was grown at 340 mmol/L NaCl, PC increased significantly. PI and PS were also significantly elevated in salinity treatments. Only 24–32% of the PM fatty acids were common plant membrane fatty acids, C16, C18, C20, and C22, while over 60% were the less common fatty acids, C11 and C14. Membrane fluidity remained stable in response to growth medium salinity. The findings on membrane responses to salinity will facilitate a better understanding of this halophyte's tactics for salt tolerance.
Key Words: halophyte • lipid • membrane composition • plasma membrane • Poaceae • salinity • salt tolerance • Spartina patens
Salinity tolerance in plants is a complicated phenomenon that includes many cellular processes, including osmoticum production, inter- and intracellular compartmentation, ion exclusion and excretion, photosynthate allocation, and signal transduction (Greenway and Munns, 1980
; Flowers, 1985
; Cheeseman, 1995
). Differentiated cells in various plant tissues and organs may function differently in coordination with neighboring cells to achieve salinity tolerance. Whether the salt tolerance or sensitivity of a plant is the result of fundamental differences in cellular level adaptations or is built upon the additive and synergistic combination of various cellular, organ-based, and tissue-based tolerance mechanisms is not clear.
One approach to studying cellular mechanisms of salinity tolerance is to use undifferentiated cell cultures. Plant cell lines from halophytes and glycophytes differ in their salt tolerance. The external level of NaCl required to inhibit plant growth to 50% of its maximum growth rate has been the criterion used in the few studies done. Cells of the halophytes Distichlis spicata (Warren and Gould, 1982
) and Spartina pectinata (Warren et al., 1985
) tolerated higher external NaCl than did cells of the glycophytes Nicotiana tabacum, N. sylvestris, or Zea mays (Hasegawa et al., 1980
). However, cells of the halophyte Suaeda maritima exhibited salt tolerance similar to that of glycophytes (von Hedenstrom and Breckle, 1974
). The optimal salinity levels for growth of Salicornia europaea and Suaeda maritima suspension cultures were about half the optima for whole plants (von Hedenstrom and Breckle, 1974
). In cell cultures of Mesembryanthemum crystallinum, the induction of a 100-kD phosphoenolpyruvate carboxylase protein and other Crassulacean acid metabolism enzymes were tissue-dependent and could not be induced in cell culture upon salinity treatment (Thomas et al., 1992
). The discrepancy of salinity tolerance in halophyte cell lines indicates that some cellular tolerance mechanisms that function in tissues with a higher level of organization, such as shoots or roots, may not function in cells at a lower level of organization, such as undifferentiated callus cell cultures or suspension cultures. On the other hand, cellular-based salinity tolerance seems to be independent of the anatomical and physiological specializations of the whole plant. For example, Spartina pectinata is a halophyte that displays less salt tolerance than the halophyte D. spicata, yet salinity tolerance of suspension cultures of S. pectinata was similar to those of D. spicata (Warren et al., 1985
).
Independently grown cells differ from whole plants in that they do not have mechanisms involving organizational complexity, such as root ion exclusion, shoot ion excretion, and ion deposition in older tissues. The undifferentiated cells depend largely upon the adaptation of ion exclusion at the plasma membrane (PM), ion compartmentation into the vacuole, and cytoplasmic osmotica production (Ben-Hayyim and Kochba, 1983
; Handa et al., 1986
; Hasegawa et al., 1986
; Binzel et al., 1988
; LaRosa et al., 1989
). Regulation of the membrane transport process is an important adaptation in cell salinity tolerance (Schobert, 1980
). Studies of sorghum [Sorghum bicolor L. and S. halepense (L.) Pers.] showed that a Na+ exclusion mechanism appears to operate at both cellular and whole plant levels for the relatively salt-tolerant S. halepense (Yang et al., 1990a
, b
). The Na+ to K+ ratios were lower in callus and roots of S. halepense than in S. bicolor at salinity levels of 0.05, 0.1, and 0.15 mmol/L. Cell suspension cultures of Kosteletzkya virginica are at least as salt-tolerant as the whole plants (Blits et al., 1993
). Still, it is not clear whether the regulation of the cell membrane in response to NaCl depends on the physiological and metabolic processes of the whole plant or is simply a strictly cellular-based adaptation.
Not until recent decades has the significance of membrane lipids in plant salinity tolerance been appreciated (Douglas and Walker, 1984
; Kuiper, 1984
; Brown and DuPont, 1989
; Blits and Gallagher, 1990
; Mansour et al., 1994
; Kerkeb et al., 2001
; Mansour et al., 2002
). Here, we studied callus tissue cultures of the halophytic C-4 grass, Spartina patens, which grows at the upper elevations of the salt marsh, to further our understanding of whether and how the membrane lipid composition in undifferentiated cells can be maintained or regulated in response to salt. The objectives were to (1) examine the salinity tolerance of undifferentiated callus cultures of S. patens and (2) investigate the expression of salinity tolerance in plasma membranes from S. patens callus.
MATERIALS AND METHODS
Callus adaptation to NaCl
Spartina patens callus was induced and maintained on BND medium (Murashige and Skoog salts [Murashige and Skoog, 1962
] +3% sucrose +0.5 mg/L 6-benzylaminopurine, 1.0 mg/L 1-naphthaleneacetic acid, 0.5 mg/L 2,4-dichlorophenoxyacetic acid, and 50 mL/L coconut water) (Li et al., 1995
) and transferred to fresh medium every 5 wk. Callus cultures were grown in 150 x 25 cm culture tubes under 30 µE · m–2 · s–1 fluorescent light with a 14 : 10 photoperiod at 22°C. Callus grown on BND medium was transferred to the same medium, and salinity in the growth media was then increased by 170 mmol/L NaCl at 5-wk intervals until the respective salinity level (0, 170, 340, or 510 mmol/L NaCl) was reached. Callus was grown for another 10 wk (with fresh media after 5 wk) at that level, then harvested for plasma membrane isolation.
Growth analysis
Callus tissue in 10 culture tubes was weighed at the beginning of the salinity treatment. Callus tissue in 5–10 tubes was harvested at the end of the experiment for growth measurements, including fresh mass, dry mass, and ash mass. Dried samples were ashed in a muffle furnace at 500°C for 2 h and measured for ash weight.
Ion analysis
Ions were extracted from the various tissues by heating ashed tissue (5–20 mg) in 2.5 mL of 1% nitric acid at 80°C for 2 h. Na+, K+, Ca2+, and Mg2+ concentrations were determined by inductively coupled plasma atomic emission spectroscopy (JY70 plus ICP-ASE, Jobin-Yvon, Edison, New Jersey, USA).
Isolation of plasma membranes
Plasma membranes were isolated from S. patens callus as described earlier (Wu and Seliskar, 1998
). Briefly, 90–100 g of callus were homogenized with a Waring blender in ice-cold 50 mmol/L (1,3)-bis[Tris(hydroxymethyl)-methylamino] propane [BTP]/Mes buffer (pH 7.2) containing 250 mmol/L sucrose, 3 mmol/L EDTA, 5 mmol/L dithiothreitol, 1 mmol/L phenylmethylsulphonyl fluoride (PMSF), and 2 mmol/L mercaptoethanol. The homogenate was further disrupted with a Polytron homogenizer and filtered through four layers of cheesecloth. Following differential centrifugation of the filtered homogenate at 14 000 x g for 20 min, total microsomal membranes were collected through ultracentrifugation at 141 000 x g for 60 min. Membrane pellets were suspended in 5 mmol/L phosphate buffer with 250 mmol/L sucrose, and then partitioned in a dextran–polyethylene glycol (PEG) aqueous biphasic system at polymer concentrations of 7.0% (w/w), each phase with 250 mmol/L sucrose and 7 mmol/L KCl in 5 mmol/L phosphate buffer at pH 7.8. After partitioning two times in the phase system, purified plasma membranes were collected from the upper phase and washed with suspension buffer, which contained 250 mmol/L sucrose in 5 mmol/L BTP/Mes (pH 7.2). Plasma membranes were then pelleted at 141 000 x g for 1.5 h and stored frozen at –80°C in suspension buffer. The purity of the plasma membrane at all salinity levels was verified with membrane enzyme markers (Wu and Seliskar, 1998
).
Membrane lipid extraction and analysis
Membrane lipid was extracted according to the methods of Bligh and Dyer (1959)
adapted by Higgins (1987)
for membrane suspension. Two mL of membrane suspension were mixed thoroughly with 7.5 mL of a chloroform and methanol mixture (1 : 2), 2.5 mL of chloroform were added, followed by 2.5 mL H2O. The mixture was shaken well, then spun at 1730 g for 5 min to separate two phases. The lower phase was collected and condensed under nitrogen to a small volume and then transferred to small, preweighed vials. The solvents were further evaporated with nitrogen, and total membrane lipids were weighed. Membrane lipids were then dissolved in a chloroform and methanol mixture (1 : 1) with 0.05% butylated hydroxytoluene and stored at –20°C for later analysis.
The total membrane lipid extract was separated into neutral lipid, glycolipid, and phospholipid fractions on silica SepPak cartridges (Waters, Milford, Massachusetts, USA) using a modified procedure based on Lynch and Steponkus (1987)
and Yongmanitchai and Ward (1992)
. Neutral lipids containing sterols were eluted with 30 mL of a chloroform and methanol mixture (100 : 1). Glycolipids were eluted with 30 mL of a chloroform and acetone mixture (24 : 52). Polar lipids containing phospholipids were eluted with 30 mL of methanol. Each fraction was condensed on a rotary evaporator and stored in small volumes in amber vials at –20°C.
Free sterols and total sterols were analyzed according to the methods of Kates (1972)
. Sterol species were determined routinely with a gas chromatograph Carle 100 (Hach Carle, Loveland, Colorado, USA) equipped with a flame ionization detector and a 15 m x 0.53 mm DB-7 capillary column (J & W Scientific, Folsom, California, USA) at a column temperature of 245°C and a helium flow rate of 30 mL min–1. Cholestan was used as the internal standard. Sterol components were identified by comparison with authentic compounds under the same conditions.
The phospholipid fraction was quantified for individual species using thin-layer chromatography with precoated glass plates (Silica Gel GHL, 250 µm layer thickness, Analtech, Newark, Delaware, USA). The mobile phase consisted of chloroform : methanol : acetic acid : H2O in the ratio of 75 : 5 : 30 : 2.2 (Baxter, McGaw Park, Illinois, USA). Individual phospholipid species were identified by co-chromatography with standards of phosphatidylcholine (PC), phosphatidylinositol (PI), phosphatidylserine (PS), phosphatidylethanolamine (PE), and phosphatidylglycerol. Lipid spots were visualized with 55% H2SO4 (by mass) containing 0.6% K2CrO3 and charred under low heat. Lipid spots were scraped off the plates, then digested with perchloric acid. Inorganic phosphorus from phospholipid was determined spectrophotometrically according to the methods of Rouser et al. (1969)
. Total glycolipids were determined according to Dubois et al. (1956)
in a reaction mixture with 5% phenol and concentrated H2SO4, using glucose as the standard.
Fatty acids of total membrane lipid extract were analyzed as fatty acid methyl esters (Sasser, 1990
). Total lipid extract was saponified in a boiling water bath for 5 min with 3.75 mol/L NaOH dissolved in an equal volume of H2O and methanol. The free fatty acids were then methylated with methyl alcohol in 3.25 N HCl at 80°C for 10 min. The methylated fatty acids were extracted with a mixture of hexane and methyl tertbutyl ether (1 : 1), then washed with 0.3 mol/L NaOH. The fatty acid methyl esters were analyzed on a Hewlett-Packard GC 5890 equipped with a 25 x 0.2 mm phenyl methyl silicone fused silica capillary column (Hewlett-Parkard, San Fernando, California, USA) and a flame ionization detector using hydrogen as the carrier gas, nitrogen as the "makeup" gas, and air to support the flame. The Hewlett-Packard 3365 ChemStation (Hewlett-Packard) was used for automated sample handling and analysis. The Sherlock software (MIDI Inc., Newark, Delaware, USA) was adopted for external calibration analysis, peak indexing, multivariate analysis, and additional database searching.
Plasma membrane fluidity was measured by steady-state fluorescence polarization using the resuspended PM fractions, diluted to give a protein concentration of 100 µg in 2 mL of the assay medium, which consisted of 40 mmol/L HEPES-KOH pH 7.0 containing 100 mmol/L KCl, 5 mmol/L MgCl2, and 0.1 mmol/L EGTA (Cooke et al., 1991
). The fluorescence probe used was 5 µL of 0.845 µg µL–1 1,6-diphenyl-1,3,5-hexatriene (DPH) in 1 : 1 ethanol and tetrahydrofuran (THF). The steady-state fluorescence polarization measurements were made with an SLM 4800S fluorescence spectrometer (SLM Instruments, Urbana, Illinois, USA) at room temperature. The fluorescence emission intensities were recorded at 428 nm, with an analyzer oriented parallel (I||) and perpendicular (I
) to the direction of polarization of the excitation beam of 362 nm. The steady-state fluorescence anisotropy (rs) and polarization (P) were calculated according to the following equations (Van Blitterswijk et al., 1981
):
 | (1) |
Statistical analysis
Using SYSTAT software (SYSTAT Software Inc, Richmond, California, USA), one-way analysis of variance (ANOVA) was conducted, and either Fisher's LSD or Tukey's multiple comparison test (as indicated in the tables and figures) was used to determine significant differences among the data.
RESULTS
Callus clumps similar in size were measured for fresh mass and used in the salinity treatments (Table 1). Callus grown at 0, 170, and 340 mmol/L salinity had a net gain in fresh mass, but callus grown at 510 mmol/L did not. Compared to the other NaCl levels, callus grown on 170 mmol/L NaCl media obtained the highest fresh mass, dry mass, and total ash mass at the end of 10 wk of salinity treatment (Table 1).
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Callus maintained a similar K+ concentration at all the tested salinity levels (Fig. 1). Salinity significantly increased Na+ concentrations in callus tissue. Callus Na+ concentration grown on 170 mmol/L NaCl increased significantly over that grown on medium without NaCl. The Na+ concentration in callus grown on 340 mmol/L NaCl was similar to that grown on 170 mmol/L, but another significant increase in callus Na+ occurred at 510 mmol/L NaCl. The relative change in Na+ and K+ concentrations resulted in a Na+ to K+ ratio change from 0.01 to 0.58, 0.73, and 2.69 as medium salinity increased from 0 to 170, 340, and 510 mmol/L, respectively.
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= 0.05). The same was true of the anistropy values. DISCUSSION
It was evident from the four salinity levels tested here (0, 170, 340, and 510 mmol/L) that S. patens callus grew well up to 340 mmol/L NaCl and that growth was stimulated at 170 mmol/L NaCl compared to callus grown without NaCl (Table 1), indicating a cellular salinity tolerance in this halophytic species. Callus grown on medium without NaCl was green, and as salinity increased, the color became lighter until, at 510 mmol/L, it was light brown (data not shown). The color change of callus in response to salinity was also observed in S. bicolor at 100 and 150 mmol/L NaCl, while there was no color change in callus of S. halepense, a salt-tolerant relative of S. bicolor (Yang et al., 1990b
). Tissue browning could be an indication of necrosis, or tissue damage, or the production of stress response compounds such as phenolic compounds.
Callus clumps were smaller in size as growth medium salinity increased from 0 to 510 mmol/L. The significant decrease in callus water content (Table 1) could be a passive adaptation to maintain favorable osmolyte concentrations to counteract growth medium salinities, as was suggested by Glenn (1987)
for salt-tolerant grass species.
Contrary to sorghum (Weimberg et al., 1984
; Yang et al., 1990a
, b
) and trifolium (Stavarek and Rains, 1984
), where callus accumulated more Na+ than whole-plant leaves, callus of S. patens accumulated less Na+ (about 15%; Fig. 1) than whole-plant shoot and root tissue (30% and 25%, respectively; Wu et al., 1998
), up to the growth medium salinity of 340 mmol/L NaCl. Therefore, Na exclusion was expressed in callus to a greater extent than it was in whole plants. But at the highest salinity tested in this study, 510 mmol/L NaCl, callus failed to maintain a low Na+ concentration. The stagnant growth of callus at 510 mmol/L NaCl (Table 1) could be a result of ion toxicity, as seen in the total ash content and ion composition (Fig. 1).
Membranes have been shown to play an important role in the ability of plants to cope under saline conditions (Douglas and Walker, 1984
; Kuiper, 1984
; Brown and DuPont, 1989
; Blits and Gallagher, 1990
; Mansour et al., 1994
). Determining how salinity affects the membrane composition of the halophyte S. patens will increase our understanding of plant salt tolerance. We found that over half of the molar lipid species in S. patens callus were free sterols, and the callus had a higher free sterol content than whole-plant roots (Wu et al., 1998
). Mansour et al. (2002)
reported a higher free sterol to phopholipid ratio, as well as a higher total sterol to phospholipid ratio in plasma membranes of wheat roots in response to NaCl. The high free sterol to phospholipid ratio raises the question of plasma membrane structural configuration, i.e., the integrity of the membrane bilayer in the presence of a high sterol content. Experimental evidence suggests that the long-range order of acyl chains would persist even at 50 mol% cholesterol, where a 1 : 1 cholesterol with diacylphospholipid interact cooperatively (Jain et al., 1984
; Presti, 1985
). The "saturated" sterol packing in the membrane bilayer would favor a very impermeable membrane, because sterols are the major component contributing to membrane rigidity (Chapman, 1973
; Van Blitterswijk et al., 1981
). Interestingly, a higher free sterol to phospholipid ratio was also observed in plasma membranes of salt-tolerant tomato callus compared to salt-sensitive tomato callus (Kerkeb et al., 2001
). Plasma membranes of S. patens callus demonstrated a stable free sterol to phospholipid ratio in response to salinity.
The relative compositional change in free sterols (namely, the increase in campesterol and decrease in sitosterol) was in line with the theory that "more planar" sterols enhance ion exclusion (Douglas and Walker, 1983
). Campesterol differs from sitosterol in that it has a methyl group at C-24 instead of an ethyl group (Nes and McKean, 1977
). Relative to sitosterol, campesterol has a smaller freedom of rotation at the C-17–hydrocarbon chain due to the smaller side chain, which would result in tighter packing in the phospholipid bilayer (Bruckdorfer et al., 1969
; Edwards and Green, 1972
) that could reduce membrane permeability to charged ions (Demel et al., 1972
; Benz and Cros, 1978
). Douglas and Walker (1984)
found that the Cl– exclusion capability of roots of different citrus genotypes was in accordance with a greater amount of "less planar" plasma membrane sterols. A high Na to K ratio (0.417, molar) also resulted in more planar free sterols in sugar beet root plasma membrane than did a low Na to K ratio (0.125, molar) (Yahya et al., 1995
). Reducing less planar sterols in the membrane might be advantageous in ion exclusion. It is interesting to note that an increase in campesterol was not evident in S. patens whole-plant roots (Wu et al., 1998
), but was significant in callus cultures. The correlation between cellular salinity tolerance vs. whole-plant salinity tolerance is still unclear (Dracup, 1991
).
The relative compositional change of phospholipid species in callus resulted in a trend of higher PC to PE ratio at higher salinities. The differences in the relative head and tail sizes of PC and PE favor two different phase configurations, the lamellar and inverted hexagonal phases (Williams, 1988
; Leshem, 1992
). Besides the interruption of bilayer structure, the inverted hexagonal phase also tends to induce hydrophilic water channels throughout the biological membranes (Caffrey, 1985
; Cullis et al., 1985
; Gruner et al., 1985
). The decrease in PC to PE ratio in oat in response to dehydration (Norberg et al., 1992
) and in wheat in response to NaCl (Mansour et al., 1994
) might result from stress damage that not only disrupts the integrity of the plasma membrane, but also causes channels to form that allow passive water diffusion across the membranes. However, for the halophyte S. patens, the observation is an unchanged or even higher PC to PE ratio in callus cultures and roots of whole plants (Wu et al., 1998
). It is also worth noting that externally supplied choline stimulated the growth of rice cell culture and resulted in an increase in the PC to PE ratio. This increase in membrane PC content was presumably related to the fast proliferation of callus (Sathishkumar and Manoharan, 1996
).
It is interesting to note the abundance of C14 and C11 fatty acids in membrane lipid of S. patens callus, up to 48 mol% for C14 and up to 23 mol% for C11, as shown in Table 4. This is unusual in higher plants. Typical in glycophytic higher plants are longer-chain fatty acids such as C16, C18, and C20 (Rochester et al., 1987
; Norberg and Liljenberg, 1991
; Cowan et al., 1993
; Mansour et al., 1994
). Studies have documented a higher percentage of short-chain fatty acids in marine algal species (Fleurence et al., 1994
; Saoudi-Helis et al., 1994
). Renaud and Parry (1994)
reported that the percentage of C14 increased in response to salinity in an Isochrysis species. The unicellular salt-tolerant green alga Dunaliella salina also responded to higher salinity in the growth medium with an increase of C14:2 in plasma membrane PE (Peeler et al., 1989
). C12 and C14 were minor components of the salt-tolerant yeast Zygosaccharomyces rouxii lipid, but increased significantly in plasma membranes when cells were grown on 2.5 mol/L NaCl (Hosono, 1992
). In S. patens, C11:0 is present in both root tissue (Wu et al., 1998
) and callus tissue plasma membrane, while absent in shoot tissue plasma membrane (unpublished data), which makes it tempting to suggest that C11:0 is related to the direct contact of tissue with the salinity medium.
Also interesting to note is that the primary C18 species is unsaturated, i.e., C18:2 (Table 4). Membrane fatty acid unsaturation has been suggested to affect cell salinity tolerance; for example, the mutation that deactivated 
-12 and -6 desaturase affected Na+/H+ antiport activity in cyanobacteria Synechocystis sp. PCC 6803 (Allakhverdiev et al., 1999
). In that study, mutant cells lost almost all antiport activity after a 30-h incubation in 1.0 mol/L NaCl, whereas the wild-type cells retained about half of the original activity in the same treatment, suggesting membrane saturation makes the antiport more sensitive to salt stress. Yu et al. (1998)
observed increased C18:3 and decreased total phospholipid content in plasma membrane isolated (using the method of sucrose gradients) from barley roots under NaCl (200 mmol/L). The prevalence of C18:2 in S. patens callus warrants more study to determine the role of membrane unsaturation in the salinity tolerance of this species.
Both membrane fluidity and changes in PM lipid composition directly affect cross-membrane transport. Changes in PM lipid composition may result in PM fluidity changes as suggested by previous studies that a less fluid PM bilayer supports reduced NaCl permeability (Kuiper, 1984
; Mansour et al., 1994
). We have reported reduced membrane fluidity in response to salinity in root PM of S. patens (Wu et al., 1998
). In the present study, independently grown cells of S. patens were able to maintain membrane fluidity throughout the salinity levels examined.
In summary, S. patens callus grew well up to 340 mmol/L NaCl, and major plasma membrane lipid classes remained stable at all salinity levels, whereas the PM protein to lipid ratio decreased as salinity increased. When calli were grown at higher salinities (170 and 340 mmol/L, as compared to 0 mmol/L), PM lipid had a higher percentage of campesterol and a higher PC to PE ratio, which may contribute to less permeable plasma membranes. Steady-state fluorescence measurements indicate that PM rigidity was maintained at the salinity levels tested. The abundance of short-chain fatty acids in S. patens PM suggests that they may play a role in the salt tolerance of cells. Our findings indicate significant plasma membrane responses to salinity in this halophytic salt marsh grass. This knowledge about membrane changes, coupled with future and past findings about other subcellular responses, may lead to a better understanding of the integrated response of whole cells.
FOOTNOTES
1 Support for this research came from the University of Delaware Sea Grant College Program under Grant No. NA16RG0162-01, project R/B 30 from the office of Sea Grant, National Oceanic and Atmospheric Administration (NOAA), U.S. Department of Commerce. The authors thank Microbial ID Inc. (Newark, DE, USA) for their assistance in analyzing plasma membrane fatty acids. 
4 E-mail: seliskar{at}udel.edu
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