Limnol. Oceanogr., 44(4), 1999, 1155–1159

Root tips from the marsh grass Spartina alterniflora, collected from areas of high and low pore-water sulfide, exhibited a substantial capacity to catalyze sulfide oxidation, as determined by closed-chamber respirometry. A large proportion of this catalysis was apparently nonenzymatic and was higher in roots of plants from the high-sulfide versus the low-sulfide site. Activity exhibiting characteristics of enzymatic sulfide oxidation was significantly higher in plants from the low-sulfide site. Results from elemental analysis of root tissue were consistent with the theory that metals play a role in nonenzymatic catalysis. These results indicate that estuarine plants may detoxify environmental sulfide via sulfide oxidation. Hydrogen sulfide is a common metabolic poison that is abundant in marine-reducing environments. Sulfide (H2S, HS, and S22) blocks aerobic respiration by inhibition of mitochondrial cytochrome c oxidase (Nicholls 1975; Wilson and Erecinska 1978) and spontaneously oxidizes in the presence of dissolved oxygen, thereby reducing oxygen availability. Despite this, many aerobic marine organisms can survive chronic sulfide exposure. The mechanisms that enable these organisms to avoid sulfide toxicity are beginning to be characterized. Most of the work in this area has involved marine invertebrates, many of which are symbiotic with sulfur-oxidizing chemoautotrophic bacteria (reviewed in Somero et al. 1989; Childress and Fisher 1992). Although plants are found in many of the same environments as the invertebrates that exhibit mechanisms of sulfide detoxification (e.g., salt marshes, eel-grass beds, and mangrove swamps), no studies have investigated the sulfide-detoxification mechanisms of marine plants. One plant that is subject to chronic sulfide exposure is Spartina alterniflora, a dominant salt marsh grass. This grass extends roots into reduced sediments that are rich in sulfide (Carlson and Forrest 1982; King et al. 1982). It is well established that sulfide can have deleterious effects (e.g., Howes et al. 1986; Pearson and Havill 1988; Pezeshki et al. 1993; Pezeshki and Delaune 1996). Environmental sulfide enters the root tissues (Carlson and Forrest 1982) and can inhibit metalloenzymes, including cytochrome c oxidase (Allam and Hollis 1972; Havill et al. 1985). Sulfide exposure correlates with stunted growth in the field (King et al. 1982) and results in reduced growth, alcohol dehydrogenase activity, adenylate charge, and nitrogen uptake in laboratory experiments (Koch and Mendelssohn 1989; Koch et al. 1990). Although sensitive to sulfide, S. alterniflora is more tolerant of sulfide than are other freshwater marsh species that do not encounter elevated sulfide (Koch and Mendelssohn 1989; Koch et al. 1990). Thus, it is apparent that while sulfide is an important environmental stressor of S. alterniflora, physiological mechanisms of sulfide tolerance may be exhibited. S. alterniflora may detoxify sulfide in ways similar to those exhibited by marine invertebrates. An important feature of the lifestyle of invertebrates that tolerate sulfidic environments is that they bridge oxic and anoxic environments. For example, invertebrates at deep-sea hydrothermal vents inhabit areas where sulfide-laden water mixes with oxygenated water. Sediment-dwelling invertebrates aerate their environment by pumping oxygenated waters into their burrows. Another strategy, exhibited by some symbiotic clams, is to live partially in aerated seawater and to extend part of the body into sulfide-rich sediments or fissures. The presence of oxygen allows sulfide to be oxidized to less toxic species, such as thiosulfate, sulfite, sulfate, and elemental sulfur. Although sulfide is spontaneously oxidized by oxygen, invertebrates have a variety of means of catalyzing sulfide oxidation in order to gain greater protective benefit and, in some cases, in order to allow sulfide oxidation to be coupled to the production of cellular energy. Mechanisms that enhance the rate of sulfide detoxification but that do not result in energy gain are sulfide oxidases and catalysis by heavy metals (reviewed in Somero et al. 1989). Mechanisms that can result in energetic gain are sulfide oxidation by bacteria symbionts (Cavanaugh et al. 1981; Felbeck et al. 1981) or mitochondrial sulfide oxidation (e.g., Powell and Somero 1986; Lee et al. 1996; Völkel and Grieshaber 1997). Like sulfide-tolerant marine invertebrates, S. alterniflora bridge oxic and anoxic environments. In S. alterniflora and other aquatic plants, a well-developed aerenchyma system facilitates the transport of oxygen from the atmosphere to the roots, where oxidation of sulfide potentially reduces its toxicity (Teal and Kanwisher 1966; Hwang and Morris 1991; Arenovski and Howes 1992; Armstrong et al. 1994; Howes and Teal 1994). Thus, sulfide oxidation may be a mechanism that allows S. alterniflora to tolerate sulfide. It is not known whether S. alterniflora actively facilitate oxidation. This plant-mediated oxidation could be catalyzed by sulfide oxidases, metals, mitochondria, or hitherto uncharacterized mechanisms. In the present study, we measured the potential for sulfide oxidation in roots of S. alterniflora and then determined whether this capacity is catalyzed by known mechanisms. S. alterniflora used in the first series of experiments were collected at Airport Marsh on Dauphin Island, Alabama, adjacent to a tidal channel (16‰). Pore-water sulfide in sediments associated with S. alterniflora at this site was high, ranging from 1.3–8.1 mM total sulfide (Lee et al. 1996). Entire plants with or without associated sediment were transported in seawater to the laboratory for further investigation. Roots of S. alterniflora were subjected to several rinses in seawater to remove associated sediments. Several tips of roots (1–3 cm; 11–44 mg dry weight) were excised and given a final rinse with filtered artificial seawater (FASW; Tropic Marin, pH ø8.0), until no sediment particles were evident under a dissecting microscope. These tips were then placed in a closed-chamber respirometer (Cyclobios Oxygraph 67097) containing 6 ml stirred (500 rpm) air-equilibrated FASW at 208C. Chamber pH was likely in the range of 8