Elemental sulfur reduction to H2S by Tetrahymena thermophila

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ScienceDirect European Journal of Protistology 62 (2018) 56–68

Elemental sulfur reduction to H2 S by Tetrahymena thermophila Dennis G. Searcy Biology Department, University of Massachusetts, Amherst, MA 01003, USA Received 14 September 2017; received in revised form 22 November 2017; accepted 24 November 2017 Available online 2 December 2017

Abstract Eukaryotic nucleocytoplasm is believed to be descended from ancient Archaea that respired on elemental sulfur. If so, a vestige of sulfur reduction might persist in modern eukaryotic cells. That was tested in Tetrahymena thermophila, chosen as a model organism. When oxygenated, the cells consumed H2 S rapidly, but when made anoxic they produced H2 S mostly by amino acid catabolism. That could be inhibited by adding aminooxyacetic acid, and then H2 S production from elemental sulfur became more evident. Anoxic cell lysates produced H2 S when provided with sulfur and NADH, but not with either substrate alone. When lysates were fractionated by centrifugation, NADH-dependent H2 S production was 83% in the soluble fraction. When intact cells that had just previously oxidized H2 S were shifted to anoxia, the cells produced H2 S evidently by re-using the oxidized sulfur. After aerobic H2 S oxidation was stopped, the oxidation product remained available for H2 S production for about 10 min. The observed H2 S production is consistent with an evolutionary relationship of nucleocytoplasm to sulfur-reducing Archaea. Mitochondria often are the cellular site of H2 S oxidation, suggesting that eukaryotic cells might have evolved from an ancient symbiosis that was based upon sulfur exchange. © 2017 Elsevier GmbH. All rights reserved.

Keywords: H2 S production; Origin mitochondria/eukaryotes; Sulfur (S8 ) reduction; Tetrahymena thermophila

Introduction The currently accepted “Tree of Life” shows the Eukaryotic Domain originating from near the base of the Archaeal Domain, either from within the lowest Archaeal branches or below them (Thiergart et al., 2012; Embley and Williams, 2015). Low-branching clades of Archaea often respire on elemental sulfur (S8 ) producing H2 S. Sulfur respiration can be Abbreviations: AOAA, 2-(aminooxy)acetic acid; CoQ, Coenzyme; Q10 , ubiquinone; DTT, dithiothreitol; GSH, glutathione; SM buffer, 50 mM MOPS (= 3-(N-morpholino)propanesulfonic acid) adjusted to pH 7.0 with NaOH; SQR, sulfide:quinone oxidoreductase; RC-1 through RC-4, respiratory complexes of the mitochondrial Electron Transport System (ETS). E-mail address: [email protected] https://doi.org/10.1016/j.ejop.2017.11.004 0932-4739/© 2017 Elsevier GmbH. All rights reserved.

thought of as analogous to O2 respiration except that H2 S is produced instead of H2 O (Achenbach-Rrichter et al., 1988; Schauder and Kröger, 1993; Kletzin et al., 2004). The clade of Archaea most closely allied to eukaryotes has been called the “TACK Group.” Several of the most interesting TACK organisms have never been seen, but are known only through DNA sequences obtained by environmental sampling (Guy and Ettema, 2011; Zaremba-Niedzwiedzka et al., 2017). Many TACK “genomes” have been annotated to contain sulfur reductase (= sulfhydrogenase) (see www.ncbi.nlm.nih.gov/gene). When living cultures have been available, it has been validated that cellular growth was

D.G. Searcy / European Journal of Protistology 62 (2018) 56–68

linked to S8 reduction and H2 S production1 (Zillig et al., 1981; Kletzin, 2007). Since dissimilatory S8 reduction (see below) is common among basal Archaea, by conjecture it could be a primitive feature of eukaryotes. The purpose of the study described below was to test for that in Tetrahymena thermophila, which was chosen as a representative eukaryotic organism. “Dissimilatory sulfur reduction” is defined as using sulfur as a terminal electron acceptor and excreting the product, which usually is H2 S. In contrast, assimilatory sulfur reduction reduces sulfur for the purpose of incorporation into biological molecules. It is important to keep in mind that both types of sulfur reduction are distinct from SO4 2− reduction, which is chemically difficult and is restricted to certain specialized bacteria (Muyzer and Stams, 2008). In animals, the most frequent source of H2 S is catabolism of sulfurcontaining amino acids (Julian et al., 2002; Kimura, 2011), which was observed during the study described below. It could be inhibited so as to better measure H2 S production from S8 . Animals cannot reduce SO4 2− (Fromageot, 1947), and that is true probably of many Protists. Other potential substrates for H2 S production contain sulfur that is less fully oxidized, such as sulfane sulfur (S0 , elemental sulfur). Sulfane is covalently bound to only other sulfur atoms, such as in cyclo-octal sulfur (S8 ) (Wood, 1987). Sulfane atoms occur also in polysulfides such as − S S S S− and organic polysulfides: R S S S− (Hedderich et al., 1999). Persulfides (R S SH) have the chemical reactivity of sulfanes because persulfides can tautomerize into “thiosulfoxide”, which contains a sulfane atom. See Toohey and Cooper (2014). Sulfane has been detected in many animal species (Ip et al., 1997; Thiermann et al., 2000; Arndt et al., 2001; Cooper and Williams, 2004). Some bivalve mollusks with symbiotic bacteria accumulate sulfane to the point of becoming yellow (Vetter, 1985). In humans, sulfane has been detected in blood, brain, liver, and kidneys at concentrations of up to 0.5 mg-atom S0 kg−1 (Westley and Westley, 1991; Kimura et al., 2015). Thus, at least in animals, sulfane is present and might be available for H2 S production. Elemental sulfur is a fungicide. When its mechanism was examined, fungi were found to reduce S8 to H2 S (Sciarini and Nord, 1943; Miller et al., 1953). Evidently, S8 causes uncontrolled oxidation of the cytoplasm to the point of death. There was one report of S8 reduction supporting anoxic fungal growth presumably by serving as a metabolic electron acceptor (Abe et al., 2007). Later the same research group wrote that “. . . elemental sulfur causes too much toxicity to measure an effect on fermentative growth” (Sato et al., 2011). Assuming there is no valid report of a eukaryote using S8 as a metabolic electron acceptor, the observations below on T. thermophila are the first example.

1 At pH 7 “H S” is a mixture of H S and HS− . I refer to the mixture 2 2 simply as “H2 S”.

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The first biochemists were fascinated by the reactions of sulfur with cellular extracts. One was de Rey-Pailhade (1888), who made ethanol extracts of yeasts and of animal tissues. When he added S8 to the extracts there was H2 S production. Such soluble extracts should not include proteins, but they do contain smaller molecules such as glutathione (GSH), and GSH can react spontaneously with S8 to produce H2 S (Sluiter, 1930). Another example used a human with medical intent was that of Monaghan and Garai (1924): a patient suffering from arthritis was given an intravenous injection of colloidal sulfur. Shortly thereafter H2 S was detected in his breath, suggesting that humans can reduce S0 . The patient’s arthritis was ameliorated, consistent with current reports that H2 S indeed can be anti-inflammatory (Szabo, 2007; Li et al., 2011). The H2 S was most likely explained by S8 reaction with GSH, which is abundant in red blood cells. Regarding the chemistry of sulfur, it is said to be “a large soft atom that often reacts without a catalyst” (Widdel and Hansen, 1992). For example, S8 reacts with organic sulfhydryl groups such GSH to produce H2 S, such as described above. Cytoplasmic proteins have abundant SH groups, and if accessible they react with S0 and produce H2 S (Roy and Trudinger, 1970). During the studies described here, in order to minimize the spontaneous reactions of S0 it was important to keep its concentration low. Toohey (1989) observed, “A remarkable feature of sulphane sulphur in biological systems is the very low and narrow concentration range at which it is effective.” Toohey (1989) estimated the physiological concentration of sulfane to be less than 1 ␮g-atom S0 L−1 . In the experiments that follow below, that was achieved by adding slowly a dilute solution of S8 to cell suspensions. The S8 was consumed almost immediately so that there was no accumulation of S0 . Tetrahymena thermophila was chosen for this study because it is a eukaryotic cell that has become a model laboratory organism (Orias et al., 1999; Asai and Forney, 1999; Collins, 2012). It is not specialized for a sulfurous environment. A disadvantage of T. thermophila is that it has proteases that interfere with investigation of cell lysates (Straus et al., 1992). This study made use of a potentiometric H2 S electrode. The electrode has certain advantages and disadvantages. Advantages are that the electrode is specific and sensitive, responding to H2 S concentrations as low as 10−15 M (Light and Swartz, 1968; Frant and Ross, 1972; Searcy and Peterson, 2004). And at equilibrium it consumes no H2 S, which can be significant with amperometric electrodes. Disadvantages of the electrode are that it responds slowly to the lowest concentrations of H2 S and that it reversibly absorbs small amounts of H2 S. And there is no agreement as to exactly how the sulfide electrode works. The specificity of the electrode is attributable to a disk of Ag2 S. The presumed mechanism is as follows: H2 S diffuses into the Ag2 S disk and then as S2− migrates through the disk by lattice diffusion. At the inner side of the disk there is a silver electrode that is oxidized

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to produce Ag2 S while creating an equilibrium electrical potential.

Material and Methods Cell culture and preparation Tetrahymena thermophila B 2086 was obtained from the Tetrahymena Stock Center, Cornell University, Ithaca, New York, USA. It was cultured in “Modified Neff Medium” (Cassidy-Hanley, 2012). When grown for an experiment, 3% inoculum was used and incubated at 21 ◦ C for about 22 h. Cultures were 1 cm deep and swirled for aeration. Actively growing cultures were used at between 25% and 70% of their maximum cell density, which was measured by optical density at 540 nm using 2 cm diameter round cuvettes in a Bausch and Lomb Spectronic 21 spectrophotometer. For each experiment the volume collected was adjusted to obtain a specific quantity of cells. The volume collected (V) was calculated using the following expression: V (mL) = 2.875/OD540 nm . For example, when OD540 nm = 0.5 then 5.75 mL of culture were collected. That contained 3,060,000 cells with 3.59 mg cellular protein. The volume collected was chilled in ice and then centrifuged at 900 × g (8800 m s−2 ) for 20 s. The cells were washed by resuspension in cold 50 mM sodium 3-(N-morpholino)propanesulfonate, pH 7.0 (SM buffer) and centrifuged again. The final cell pellet was suspended in cold 5 mL SM buffer and tested immediately.

Cell lysates and centrifugal fractions A separate cell lysate was prepared immediately before each experiment. Cells equivalent to 3.59 mg protein were collected, washed, and resuspended in cold 5 mL SM buffer. The suspension was sheared twice using a gas pressure cell (Simpson, 2010) and 600 psi N2 (4000 kPa). Lysis was >99%. The lysate was centrifuged at 900 × g for 20 s, producing no visible pellet. The supernatant suspension was tested immediately. Centrifugal fractions were obtained from a 2× concentrated lysate (7.18 mg protein in 5 mL SM). It was centrifuged by acceleration to 17,000 rpm (30,000 × g) and stopping the centrifuge immediately. Acceleration plus deceleration took 5.5 min.

collar around each adapter 15 mm from the lower end. When simultaneous recordings from 2 probes were needed, a larger chamber and adapter were available. Stirring was provided by a glass-covered stirring magnet. The sample chambers were surrounded by 30 ◦ C circulating water. The sulfide electrode was a combination-type from Thermo Scientific (Orion 4216BN). At the end of each day the electrode was drained, rinsed, and stored dry. When put back into use the electrode was filled with water and conditioned by soaking for 10 min each in 10 mM AgNO3 , then in 10 mM Na2 S, and in 2 mM dithiothreitol. Then the reference part of the electrode was drained and refilled with 10% (w/v) KNO3 , 1 mM KCl, and 100 ␮M AgNO3 . The electrode was attached through a high input impedance voltmeter (signal conditioner) to a computer equipped to digitize and record the data. The electrode voltage was sampled the 10,000 times s−1 , and each 5 s the data were averaged and recorded. The electrode was calibrated using 50 ␮M Na2 S in SM buffer. The apparatus was similar to the “Sulfidostat” of Searcy and Peterson (2004), but modified to inject S8 solution instead Na2 S. Oxygen was measured with a polarographic Clarktype probe (Yellow Springs Instruments, Yellow Springs, Ohio) fitted with a Teflon membrane. For calibration, airequilibrated SM buffer at 30 ◦ C was calculated to contain 241 ␮M O2 (Rasmussen and Rasmussen, 2003). Sulfur (S8 ) dissolved in ethanol was added to the sample using a microliter syringe and a syringe pump. A Hamilton “gas-tight” syringe (250 ␮L) was fitted with Intramedic PE20 polyethylene tubing 10.5 cm long × 0.38 mm ID. The tip of the tubing was heated and drawn to 0.1 mm opening. The S8 solution did not contact metal. The syringe pump was set to deliver typically 2.12 nL s−1 .

H2 S production For anoxic conditions purified N2 gas was bubbled through the sample for 1 min, and the electrode then lowered into the sample while carefully excluding air. Measurements showed that bubbling with N2 removed O2 with a half-time of about 10 s. When the sample was cell lysate, foaming was reduced by smearing 1 ␮g Antifoam A (Sigma Chemical) onto the vial wall. During experiments, to ensure anoxia, the overflow collar above the sample was flushed continuously with N2 . The rate of H2 S production usually became constant after 15 min and then was measured using the increase in H2 S concentration.

Experimental apparatus The experimental chamber was a glass vial 20 mm ID × 65 mm high. Specific electrodes for either H2 S or O2 were fitted with acrylic adapters so as to fit snugly inside the vial. A 1 mm groove down the side of each adapter allowed removal of bubbles and access to the sample for making additions. Sample overflow was accommodated by an indented

Blank corrections When the chamber contained only anoxic SM buffer and H2 S there was loss of H2 S from the solution at a rate that was approximately proportional to the H2 S concentration. The rate constant for H2 S loss was −5.5 × 10−9 M min−1 ␮M−1 , which usually was insignificant. When polysulfide (5 mgatom L−1 ) was added to 5 mL SM buffer at 254 nL min−1 ,

D.G. Searcy / European Journal of Protistology 62 (2018) 56–68

due to H2 S in the polysulfide solution the H2 S concentration increased at a rate 44 nM min−1 , and that was subtracted from the experimental data.

1

In polysulfide stock solutions, S0 was measured using the cold cyanolysis assay (Wood, 1987). A second technique, for validation, was to dilute the polysulfide into SM buffer and then add dithiothreitol to 5 mM, measuring the release of H2 S (Kwasniewski et al., 2011). The results of the two techniques did not differ significantly. On occasion, H2 S was assayed using the methylene blue colorimetric assay (Greenberg et al., 1981), using a gravimetric Na2 S standard. Protein was measured using the BCA method (Smith et al., 1985) calibrated by using a series of bovine serum albumin concentrations. The standard curve was fit by a polynomial that then was used for calculations. Cell numbers were determined by dilution of cell suspensions into 4% formaldehyde and counted using a hemocytometer and microscope. Solutions Elemental sulfur (S8 ) was dried for 2 days in an oven and then dissolved to 5 mg-atom S0 L−1 in ethanol (= 625 ␮M S8 ). Several days were allowed to ensure complete dissolution. A stock solution of 1 M Na2 S was made by dissolving Na2 S·10H2 O in water using only transparent glass-like crystals. The solution could be stored without loss for several months in a polypropylene bottle at −70 ◦ C. Polysulfide stock solutions (5 mL) were made by adding 100 mg S8 to 0.1 M Na2 S, 0.01 M NaOH solution. The suspension was stored under vacuum for several days with occasional agitation. Then the clear yellow solution decanted and stored under N2 . According to cold cyanolysis assay, the polysulfide stock solution contained about 240 mgatoms S0 L−1 . That was diluted to 10 mg-atoms S0 L−1 in 10 mM NaOH and stored under N2 . Each day the diluted stock solution was assayed using cold cyanolysis, adjusted to 5 mg-atom S0 L−1 in 10 mM NaOH, and placed into the microliter syringe for experimentation. Other stock solutions: 50 mM 2-(aminooxy)acetic acid (AOAA, Sigma–Aldrich); 100 mM KCN; 4 M NaN3 ; 0.2 M NADH; and 0.02 M NADPH, each dissolved in water. Rotenone (5 mM, Fluka) was dissolved in ethanol.

Results My initial experiments with T. thermophila and sulfur used a bolus of 1% S8 (w/v), which is typical for bacterial growth experiments. There was substantial H2 S production, but boiled controls produced even more (see later) and so the project was abandoned. Later it was reasoned that nonspecific sulfur reactions might be reduced by reducing the

H2 S

O2

0.5

Assays

59

75 50

[O2]

25 0

0 0

10

20

30

40

50

Fig. 1. Sulfide production and consumption by intact cells. Starting with 65 ␮M O2 in the buffer (blue line), it was consumed in 2.5 min. After about 8 min delay, H2 S production began (black line). At 19 min and again at 41 min 20 ␮L bubbles of O2 were injected into the sample chamber, and then H2 S was rapidly consumed. Near 20 min and 42 min the O2 bubbles were removed, and after the cells had consumed the O2 then H2 S production resumed. Procedure: growing cells containing 7.18 mg protein were washed and suspended in 10 mL SM buffer. The suspension was placed in a chamber equipped with both an H2 S electrode and an O2 probe. Sulfur (S8 ) dissolved in ethanol was pumped into the chamber continuously at the rate of 21.2 × 10−12 g-atom S0 s−1 . The inhibitor AOAA was not present. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

concentration of S8 , which was achieved by adding dissolved S8 slowly and continuously using a microliter syringe and syringe pump. A typical rate of S8 addition was 11 × 10−12 gatom S0 s−1 , which was chosen as the slowest rate that yielded easily measured results. It was found that the first experiment each day produced more H2 S than subsequent experiments. After storage, when the H2 S electrode was treated with dithiothreitol, H2 S was released. A second treatment with dithiothreitol released no additional H2 S. That suggested that after storage there was S0 in the electrode, and it became standard procedure at the start of each day to treat the electrode with dithiothreitol. In addition, results of each day’s first experiment were discarded. Those two precautions reduced experimental variation.

General features of H2 S production Fig. 1 shows that when O2 was available the cells rapidly consumed H2 S, and when anoxic they produced H2 S somewhat more slowly. The rate of H2 S oxidation in cell-free buffer was insignificant compared to that by cells. Aerobic H2 S consumption in Tetrahymena was studied previously (Searcy, 2006), and should be consulted for the relevant controls. Since H2 S oxidation occurs much faster than H2 S production, if both production and consumption occur simultaneously then there will be no net H2 S accumulation. Anoxia is required for accumulation of H2 S. That is a common pattern observed in animals, fungi, Bacteria, and Archaea (Zillig et al., 1986; Abe et al., 2007; Muyzer and Stams, 2008; Olson et al., 2009, 2013). The production of H2 S was confirmed by the methylene blue assay and also by a novel “Silverstat” experiment, as follows. The Silverstat was a variation on the earlier Sulfi-

60

Growing cells were washed and resuspended in 5 mL 50 mM sodium MOPS, pH 7.0. Data were collected using either an H2 S electrode or an O2 electrode. When anoxic, suspensions had been purged with N2 . When S8 was added, it was injected at a rate of 10.6 × 10−12 g-atom S s−1 . Data are means ± SE (N). Negative zero (−0.00) means that the trace of H2 S already present was slowly consumed at a rate that rounded to zero.

± ± ± ± ± 807 187 115 804 125 ± ± ± ± ± 815 166 130 964 126 ± ± ± ± ± −0.00 −0.00 +0.49 +0.01 −0.00 ± ± ± ± ± −0.00 −0.00 −0.00 +0.00 −0.00 No inhibitor KCN, 1 mM NaN3 , 40 mM AOAA, 100 ␮M Rotenone, 30 ␮M

1.37 0.12 0.25 0.00 0.90

± ± ± ± ±

0.18 (13) 0.08 (5) 0.05 (4) 0.00 (5) 0.17 (8)

Aerobic

0.00 (15) 0.00 (5) 0.00 (8) 0.00 (2) 0.00 (4)

5.16 0.78 3.00 4.14 3.59

± ± ± ± ±

1.01 (13) 0.27 (4) 0.64 (6) 0.37 (10) 0.52 (11)

Aerobic

0.00 (9) 0.00 (6) 0.19 (7) 0.01 (7) 0.00 (4)

Aerobic

7 (39) 10 (5) 11 (6) 24 (10) 4 (15)

Aerobic

60 (17) 14 (6) 12 (5) 70 (11) 8 (6) Anoxic

S8 added

Anoxic

Inhibitor

Table 1. Rates of sulfide production and O2 consumption.

dostat technique (Searcy and Peterson, 2004). In a Silverstat experiment H2 S is kept at a constant concentration by addition of AgNO3 . When H2 S increased, AgNO3 is added by a computer-controlled syringe pump, precipitating Ag2 S and keeping the H2 S concentration constant. A typical set point was 1 ␮M H2 S. The rate of H2 S production was calculated as one-half the rate of AgNO3 addition. When measured by this technique, the rate of cellular H2 S production was nearly the same as that measured by H2 S accumulation. A concern with the “Silverstat” technique was that Ag+ might be toxic. But, the solubility product of Ag+ and H2 S at pH 7 is 10−43 (Licht, 1988), so that in 1 ␮M H2 S the concentration of Ag+ calculates to less than one atom per liter. Cell culture experiments showed that adding 1 ␮M Ag+ to culture medium did not affect growth. Disadvantages of the “Silverstat” included fouling of the apparatus by the Ag2 S precipitate and possible toxicity to cells due to ingestion of Ag2 S particles. After this experiment the “Silverstat” technique was not used again. With no added S8 , cell suspensions nonetheless produced H2 S (see Fig. 2). That can be explained by catabolism of sulfur-containing amino acids since it was inhibited by adding 100 ␮M aminooxyacetic acid (AOAA). With AOAA present, H2 S production was dependent upon added S8 . Table 1 summarizes the effects of certain inhibitors on H2 S production and O2 consumption by T. thermophila. Conventional inhibitors of O2 consumption (KCN, NaN3 , and rotenone) were not entirely effective. The sister species, T. pyriformis, is similarly resistant to complete inhibition (Turner et al., 1971; Unitt et al., 1983). Intact cells of T. thermophila consumed O2 at about 2/3 the specific rate (per mg protein) reported for purified mitochondria from T. pyriformis (Turner et al., 1971). Regarding aerobic H2 S consumption, the rates in Table 1 are much slower than those reported previously (Searcy, 2006). That is easily explained because in 2006 the concentration of H2 S was maintained at about 10−7 M by

No S8

Fig. 2. Anoxic H2 S production occurred spontaneously, but not when aminooxyacetic acid (AOAA) had been added. With AOAA present, H2 S production resumed when S8 was added. Cells (3.68 mg cellular protein) were washed and suspended in 5 mL SM buffer and made anoxic by flushing with N2 . (A) Control, washed cells. (B) Cells with 100 ␮M AOAA. (C) Cells with AOAA and with S8 added continuously at 21.2 × 10−12 g-atom S0 s−1 .

S8 added

B. With AOAA

No S8 added

C. With AOAA + S8

Rates of O2 consumption (nmol s−1 g protein−1 )

A. Anoxic cells only

Rates of H2 S production (nmol s−1 g protein−1 )

D.G. Searcy / European Journal of Protistology 62 (2018) 56–68

D.G. Searcy / European Journal of Protistology 62 (2018) 56–68

Table 2. Rates of H2 S production in 100 ␮M AOAA in combination with other inhibitors. Inhibitor added in addition to AOAA

Rates of H2 S production (nmol s−1 g protein−1 )

None KCN, 1 mM NaN3 , 40 mM Rotenone, 30 ␮M

4.14 ± 0.37 (10) 0.02 ± 0.01 (5) 3.96 ± 0.52 (5) 2.67 ± 0.43 (7)

Conditions as in Table 1. Cell suspensions were anoxic and S8 added slowly.

additions of Na2 S whereas in the present study no Na2 S was added and concentrations of H2 S dropped as low as 10−12 M. When NaN3 had been added there was the unusual phenomenon of significant H2 S accumulation in aerobic conditions. That is inconsistent with the generalization that dissimilatory sulfur reduction occurs only in the absence of O2 (please see Discussion). Since AOAA inhibits H2 S production from the catabolism of amino acids, H2 S production from S8 was measured more easily when AOAA was present. In all experiments after this point 100 ␮M AOAA was present. For example, in Table 2 several inhibitors were tested in combination with AOAA. With AOAA present, KCN prevented H2 S production from S8 , but NaN3 did not, and rotenone was partially inhibitory.

Sulfanes other than S8 Fig. 3 compares 3 types of sulfane for H2 S production. Sulfide production from polysulfide started with less delay and was faster than from S8 .

2

Polysulfide S8

1

61

Controls showed that S8 and Na2 S2 O3 injected into SM buffer with no cells resulted in no detectable H2 S. In contrast, when polysulfide was injected into buffer then H2 S accumulated at a steady rate. The explanation may be that polysulfides can disproportionate to produce H2 S (Giggenbach, 1972), such as: 4HS3 − + 4H+ ↔ 4H2 S + S8

(1)

The effect of pH is evident. The polysulfide stock solution was pH 12 and was injected into SM buffer, pH 7. With decreased alkalinity, H2 S will be released. Nonetheless, when cells were present the amount of H2 S released exceeded that in the blank. When polysulfide (5 mg-atom S0 L−1 ) was added at the standard rate to 5 mL SM buffer, H2 S concentration increased at 44.2 nM min−1 . When cells were present, without subtracting the blank H2 S increased at 199 nM min−1 . Thiosulfate (S2 O3 2− ) was not a substrate for significant H2 S production by T. thermophila. Previously Olson et al. (2013) reported that in homogenates of mammalian tissues 10 mM S2 O3 2− was a good substrate. That is at least 1000× more than in my experiments, but to test their conditions I added 10 mM S2 O3 2− to a T. thermophila cell suspension (with AOAA present), and no H2 S was produced. A cell lysate that contained 1 mM NADH and 100 ␮M AOAA was tested with 5 mM S2 O3 2− , and it produced no H2 S. (For more about cellular lysates, see later.) Several other sulfur compounds were tested but none resulted in detectable H2 S production. Compounds tested included SO3 2− , SO4 2− , and S4 O6 2− . When S8 was injected into a cell suspension there was about 10 min delay before H2 S production began. In contrast, when polysulfide was injected the delay was significantly less. As a control, a syringe was filled with 5 mM Na2 S and pumped at the same rate into a chamber containing 5 mL SM buffer. After a lag of about 2 min the electrode began to report an increase in H2 S concentration. Thus, in cell suspensions the 10 min delay in H2 S accumulation is explained mostly by a lag in the onset of cellular H2 S production. In the experiments that follow, sulfane was added usually as S8 rather than polysulfide. That was because polysulfide stock solutions were poorly defined and were unstable, and because polysulfide by itself produced significant rates of H2 S accumulation.

Na2S2O3 0 0

10

20

Fig. 3. Tests of 3 different substrates for H2 S production. Washed cells (2.60 mg protein) were suspended in 5 mL SM buffer containing 100 ␮M AOAA, made anoxic, and then the different sulfane compounds each injected at the rate of 21.2 × 10−12 g-atom S0 s−1 . The injected solutions were: 0.625 mM S8 dissolved in ethanol, 5 mM Na2 S2 O3 in water, or 5 mg-atoms S0 as polysulfide dissolved in 10 mM NaOH. The accumulation of H2 S in buffer when polysulfide was injected has been subtracted from the polysulfide data. Please see the text. Each line is the average of at least 2 replicates.

Rates of S0 addition The prediction was that the rate of H2 S production would be proportional to the rate of S0 addition, but data in Fig. 4 show otherwise. When S8 was injected at the slowest rate then H2 S production accounted for only about half of the injected sulfur. At higher rates of S8 injection the data plot became parallel to a 45 ◦ line suggesting that from the additional S8 the yield was near 100%. To test whether the rates in Fig. 4 had reached their full velocity and whether the missing sulfur was absorbed into a

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was accumulation of 0.2–0.4 ␮M H2 S, which is the estimate of the S0 concentration in the cell suspension during S8 injection. If some S0 has a fate other than H2 S production, that will be an underestimate, while the slow response of the H2 S electrode can cause an overestimate. Altogether, during S8 injection a plausible estimate of S0 pool is 0.2–0.4 ␮g-atom S0 L−1 , which includes S0 both in the cells and in the buffer. A second estimate of the steady-state S0 concentration can be the amount of S8 injected before H2 S production reaches steady state. Examining the 2nd and 3rd restarts of S8 addition led to an estimate similar to the value above.

30

20

10

0 0

20

40

Fig. 4. Rates of H2 S accumulation at differing rates of S0 addition. Rates were measured >20 min after the start of sulfane addition when rates had become steady. (䊉): 5 mg-atom S8 L−1 dissolved in ethanol. (): S8 was first injected at 22 × 10−12 g-atom S s−1 and then after 20 min the rate was reduced to 10.8 × 10−12 . The cell suspensions each contained 3.8 mg protein in 5 mL SM buffer with 100 ␮M AOAA. (): controls in which 5 mM Na2 S instead of S8 was injected into 5 mL of SM buffer. Conditions were anoxic. Data have been corrected for loss of H2 S from cell-free buffer, which was about 5% of the rates of cellular H2 S production.

saturable sink, S8 was added first at a higher rate for 20 min and then shifted to a slower rate (open circle). That manipulation did not significantly affect the result. As an additional control, Fig. 4 includes the injection of Na2 S solution into cell-free buffer. The rates of H2 S accumulation were close to expectations, eliminating a variety of possible instrumental errors and validating the technique.

Estimate of steady-state S0 in a cell suspension during injection of S8 Fig. 5 shows how stopping and starting S8 injection affected H2 S production. After stopping S8 injection there

20

5 4 3 2

Rate of S8

(gS0 sec−1 12 0 × 10 )

1 0 0

20

40

60

Fig. 5. Production of H2 S while starting and stopping S8 injection. S8 was pumped into an anoxic cell suspension at rates of either 21.2 pg-atom S0 s−1 or zero (right axis, blue line). Black line, left axis: concentration of H2 S. While S8 injection was paused the delivery tube was removed from the sample chamber. Small jumps at 30 min and 50 min were caused by reinserting the delivery tube. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

H2 S oxidation produced a compound that could be used again for H2 S production Cellular oxidation of H2 S occurs by the mitochondrial enzyme sulfide:quinone oxidoreductase (SQR). The initial product is thought to be a persulfide attached directly to the enzyme (Libiad et al., 2014). Shortly after that the S0 atom is transferred to other molecules that have not been identified with certainty. Then the S0 is oxidized to S2 O3 2− or to SO4 2− , which in animals are mostly excreted. But while it remains S0 there is the possibility that it can be reduced again to H2 S. To test that, cells were allowed to oxidize H2 S for 10 min with O2 present. Then O2 was removed, and when incubation continued the cells produced H2 S (Fig. 6A). Cells that had not oxidized H2 S just previously did not produce H2 S (Fig. 6C). The yield of H2 S was not great: during the initial aerobic phase 4.2 × 10−7 mol H2 S were consumed while during the anoxic phase 2.3 × 10−10 mol H2 S were produced. Without S8 injection the cells produced H2 S only when they had oxidized H2 S just previously. By conjecture, the H2 S was produced from an ephemeral product of H2 S oxidation. To measure how long the oxidation product remained available for H2 S oxidation, cells were incubated aerobically with H2 S for 10 min, and then without added H2 S aerobic incubation was continued for varying periods of time. Finally the suspensions were made anoxic and H2 S production measured. The results are summarized in Fig. 7. The capacity for H2 S production declined with a half-time of about 12 min, which therefore is the estimate of the lifetime of the recyclable sulfur product of H2 S oxidation. When the product of H2 S oxidation is recycled, might that occur by reversal of the normally oxidative SQR reaction? NADH is a strong enough electron donor that potentially it can reduce S0 to H2 S (see Discussion). If so, then electrons from NADH are likely to enter through mitochondrial RC-1, then to CoQ, SQR, and finally S0 . To test that, RC-1 activity can be inhibited by rotenone, and that was tested as shown in Fig. 6D. Unexpectedly, adding rotenone increased H2 S production 3.3-fold. Evidently, for this type of H2 S production, electrons do not pass through RC-1 and it is unlikely that H2 S is produced by a reversal of the normal SQR reaction.

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Fig. 6. H2 S oxidation followed by anoxic H2 S production. (A) For 10 min the cells were provided with both O2 and 1 ␮M H2 S. Then O2 and H2 S were removed by vigorously purging the suspension with N2 for 30 s. The anoxic cell suspension produced H2 S. (B) Similar to A except that aerobic incubation was continued for 7 min after Na2 S addition was stopped. (C) Control in which no Na2 S was added during the initial aerobic incubation. (D) Rotenone (30 ␮M) present, but otherwise as in A. All cell suspensions included 100 ␮M AOAA. No S8 was added. During aerobic incubation Na2 S addition was by a computer-controlled syringe pump that maintained 1 ␮M H2 S in the cell suspension. When aerobic, the O2 concentration was maintained near 75% of equilibrium with air (∼180 ␮M O2 ) by manually adding small bubbles of O2 . Oxygen concentrations fluctuated, but for simplicity are shown as constant in the figure.

H2 S production in cell lysates y = e-0.060x R² = 0.85 0.5

0 0

10

20

Fig. 7. Cells were incubated aerobically for 10 min with 1 ␮M H2 S as in Fig. 6A. After 10 min Na2 S addition was stopped and aerobic incubation continued for the time intervals shown on the x-axis. Then the cell suspensions were made anoxic and the rates of H2 S production measured. Experiments from 4 separate days were combined. Variation between days was reduced by normalizing the “zero time” H2 S production to 1.0. The average “zero time” rate of H2 S production was 0.015 ␮M H2 S min−1 .

NADH and NADPH are likely electron donors for H2 S production, but since they cannot pass through membranes they were tested using cell lysates. Data are shown in Fig. 8. Lysates supplied with 0.1 mM NADH produced 1.6 nmol H2 S s−1 (g protein) −1 and there was no significant difference between NADH and NADPH. The concentrations of 1 mM NADH and 0.1 mM NADPH were chosen to approximate physiological concentrations (Reiss et al., 1984; Cambronne et al., 2016). In a typical cell the total concentration of NADH plus NAD+ is about 1 mM, which in aerobic conditions is 90% oxidized. But in anoxic conditions the NAD+ /NADH pool is likely to become mostly reduced and therefore ∼1 mM NADH. When polysulfide was tested on cell lysates H2 S was produced at a rate not significantly different from S8 addition, although with polysulfide H2 S production started more quickly (Fig. 9). When the lysate was provided with polysulfide but not NADH, the data dipped below zero (Line 2, Fig. 9). The

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Fig. 8. Cell lysates were tested for H2 S production with addition of S8 and either NADH or NADPH. Each lysate contained 4.2 mg protein and 100 ␮M AOAA in 5 mL SM buffer. The lysate was made anoxic by purging with N2 , and then S8 was added at a rate of 10.6 × 10−12 g-atom S0 s−1 . NADH or NADPH was added as shown. The control (dotted, lying on the X-axis) received S8 but no NAD(P)H. Each curve is the average of 2 experiments.

Fig. 9. Polysulfide added to lysates. Line #1 is a lysate containing 1 mM NADH and with polysulfide added at a rate of 10.6 × 10−12 gatom S0 s−1 . Other lines of data are controls or are shown for comparison. #2 (blue)—lysate plus polysufide, no NADH. #3 (red)—lysate plus NADH, no polysulfide. #4 (orange)—intact cells (not a lysate) plus polysulfide. #5 (black dashed line)—S8 added to a lysate with 1 mM NADH. Blank (not shown)—when the polysulfide stock solution was added to SM buffer there was accumulation H2 S, and that has been subtracted from each experiment to which polysulfide was added. Other conditions were as in Fig. 8, including 100 ␮M AOAA in each lysate.

explanation is that addition of polysulfide to SM buffer resulted in slow accumulation of H2 S, and that was subtracted from the experiments that included lysates. But in this experiment, for about 10 min, H2 S accumulated more slowly in the lysate than in the blank. Thus, when the blank was subtracted the data were negative. After 10 min there was slow net accumulation of H2 S, which perhaps can be explained by enzymatic disproportionation of the polysulfide.

Centrifugal fractions To test whether the site of H2 S production from S8 was in the soluble cytosol or in a particulate fraction, lysates

were fractionated by centrifugation. Table 3 shows that of the NADH-stimulated H2 S production, 83% was in the cytosol. There was H2 S production in the particulate fraction, but it was stimulated less by added NADH. To confirm that mitochondria were exclusively in the particulate fraction, the conventional mitochondrial marker is succinate-dependent O2 consumption. Such activity was easily measured in the lysate, and after centrifugation was present in only the particulate fraction, as expected. The activity diminished with time, as also did NAD(P)H-stimulated H2 S production. Further purification of mitochondria was attempted, including Percoll gradient centrifugation (Sims and Anderson, 2008), but then succinate-stimulated O2 consumption and NADH-dependent H2 S production became undetectable in any fraction.

Boiled lysates A standard criterion for a reaction to be enzymatic is that it should be inhibited after heating to 100 ◦ C. When that test was done there was S8 -stimulated H2 S production in both boiled cells and boiled lysates. After boiling some intact cells could be seen, and so in the following tests for simplicity the cells were first lysed and then placed in boiling water for 10 min. Without S8 , the lysate produced H2 S, but produced more when S8 was added (Fig. 10). Adding NADH did not increase H2 S production. Most importantly, a not-boiled lysate with S8 but no added NADH did not produce H2 S (Fig. 8, dotted line). Evidently, H2 S production was activated by boiling and then it was not dependent upon NADH. By conjecture, the source of the H2 S might have been from persulfides (R-SSH). To test that, dithiothreitol was added, which is expected to release H2 S from persulfides

Fig. 10. H2 S production in boiled lysates. (1) boiled lysate to which S8 was added slowly at 10.6 × 10−12 g-atom S0 s−1 . (2) Same as 1 except with 1 mM NADH. (3) Boiled lysate, no additions. (4) Same as 3 except with 1 mM NADH. (5) Unboiled lysate with S8 slowly added. Line (5) is from Fig. 8. Each lysate was prepared and then heated to 100 ◦ C for 10 min immediately before each experiment. After cooling each lysate was adjusted to 100 ␮M AOAA and, as needed, 1 mM NADH. Each lysate contained 4.3 mg protein in 5 mL SM buffer. Each line is the mean of 4–7 replicates.

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Table 3. Centrifugal fractions of cell lysates tested for H2 S production. Centrifugal fraction

Soluble, no NADH added “ + 1 mM NADH Particulate, no NADH “ + 1 mM NADH

Rate of H2 S accumulation (mol L−1 s−1 × 1012 ) 19 920 56 239

± ± ± ±

NADH-dependent H2 S production (mol L−1 s−1 × 1012 )

12 (6) 140 (9) 14 (8) 30 (8)

901 183

Specific activities (mol s−1 (g protein)−1 × 1012 ) 20 905 96 483

± ± ± ±

12 (6) 135 (9) 28 (8) 92 (8)

Cells were washed and suspended in 5 mL SM buffer, lysed, and then centrifuged briefly by acceleration to 20,000 × g. The centrifugal pellet was resuspended in 5 mL SM buffer. Each preparation (soluble and particulate) was adjusted to 100 ␮M AOAA and 1 mM NADH, made anoxic and, and then S8 added at 10.4 × 10−12 g-atom S0 s−1 . Protein concentrations were- soluble fraction about 1.0 g L−1 and particulate fraction about 0.5 g protein L−1 .

(Ubuka, 2002). Nonetheless, adding 2 mM dithiothreitol to boiled lysates increased by only 5%–10% the amount of H2 S released during the first 10 min of incubation. Thus, the source H2 S released by boiled lysates has not been identified.

Discussion T. thermophila cytosol reduced S8 to H2 S using either NADH or NADPH as an electron donor. This is the first report of dissimilatory S0 reduction by a eukaryote– that is, using S0 as a terminal acceptor for metabolic electrons and excreting the product.

“Accidental” H2 S production It is possible that the enzyme(s) responsible for H2 S production from S8 might have been described already, but reported to catalyze a different reaction. For example, the sulfur reductase of Pyrococcus furiosus initially was described to reduce CoA dimmers, and only later was it shown to reduce CoA-SSH producing H2 S (Schut et al., 2007). In eukaryotes such an enzyme has not been reported, but there are other disulfide reductases that might accidentally produce H2 S. The resolution of these possibilities awaits purification of the putative S0 reductase enzyme.

Membrane permeabilities to H2 S and to S8 Non-enzymatic S8 reduction An operational criterion for testing whether a given reaction is enzymatic is that should be inhibited after heating to 100 ◦ C. When T. thermophila lysates were boiled and then tested, H2 S production occurred nonetheless. When the boiled lysates were given S8 , H2 S production increased still more. But adding NADH to a boiled lysate, either with or without S8 , caused no increase in H2 S production. In contrast, not-boiled (native) lysates behaved differently. Adding S8 by itself did not cause significant H2 S production, but only when both S8 and NADH were added was there substantial H2 S production (See Figs. 8 and 10). Without lysate, NADH and S8 in SM buffer did not interact. Thus, although having failed the “boiling test” for enzymatic catalysis, evidence suggests that cellular production of H2 S from S8 is enzymatic. The best reason for concluding that H2 S production was enzymatic was that it was NADH-dependent. Sulfide production increased in boiled lysates (Fig. 10). A likely explanation is that boiling exposed buried R-SH and R-SSH groups making them available for reaction with each other and with added S8 . For example:

The “solubility-diffusion” model describes how small molecules pass through a membrane: the molecule dissolves in lipid bilayer and then diffuses through. Overton’s Rule (1896) states that membrane permeability correlates with the molecule’s oil/water partition coefficient (Walter and Gutknecht, 1986). The solubilities of H2 S is high in lipids and in water, and it diffuses easily through membranes (Cuevasanta et al., 2012). Elemental sulfur (S8 ) has not been studied in this regard, but an approximation can be made from its solubilities in water and lipids. The solubility of S8 in water is 5 ␮g L−1 and in olive oil it is 250 g L−1 (Seidell, 1911; Boulegue, 1978). Olive oil and membrane lipids both are mostly fatty acids, so that probably is a valid approximation. Thus, S8 is predicted to dissolve in a membrane and then perhaps diffuse out the other side. But the solubility of S8 in fat is so much greater than in water, it might also be expected to remain in the lipid bilayer. If so, then its initial enzymatic reaction might occur there. That is problematic because the data indicated that S8 reduction occurred mostly in the soluble fraction of the cells whereas membranes are in the particulate fraction.

R-SH + R-SSH → R-SS-R + H2 S

(2)

Respiratory inhibitors: NaN3 , KCN, and rotenone

2R-SH + S0 → R-SS-R + H2 S

(3)

The “hierarchy of electron acceptors” is the generalization that in nature the electron acceptor with the most strongly

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positive reduction potential will be used exclusively until it is gone, and then the next most positive electron acceptor will be used (Stumm and Morgan, 1981). When O2 is available, it is used first, and not sulfur. Nonetheless, in T. thermophila when azide had been added there was significant S0 reduction in the presence of O2 (Table 1). The explanation is that azide inhibited the mitochondrial electron transport system (ETS). There are two possible schemes: (1) H2 S may be produced continuously in aerobic conditions, but remained undetected because SQR immediately re-oxidized all of it. When azide inhibited RC-4 that inhibited SQR activity, and then there was net accumulation of H2 S. (2) Perhaps H2 S production is inactive in aerobic cells. When azide was added then reducing equivalents accumulated. Electrons that normally would go to O2 were diverted to S0 , and H2 S production was induced. Cyanide (KCN) is similar to azide in specifically inhibiting mitochondrial RC-4. Thus, it is interesting that it had an opposite effect: KCN inhibited H2 S production. The explanation may be “cold cyanolysis”, which is the spontaneous reaction of CN− with S0 to produce SCN− (Wood, 1987). In addition to cold cyanolysis, transfer of S0 to CN− can be augmented by the enzyme “rhodanese” (Cipollone et al., 2007). Thus, KCN inhibited H2 S production by removing the S0 substrates for H2 S production. The effect of rotenone on H2 S production is harder to explain. In intact cells with AOAA, rotenone inhibited H2 S production by about 1/3 (Table 2), suggesting that there may be more than one mechanism of H2 S production. But in experiments where cells were incubated first with H2 S aerobically and then anoxically, adding rotenone more than doubled the amount of H2 S produced. (Fig. 6D) The increase might be explained by considering “fumarate respiration”. During fumarate respiration NADH is oxidized by RC-1. The electrons then flow to CoQ and then to succinate dehydrogenase, which is run in reverse reducing fumarate. When rotenone inhibits RC-1 then more electrons become available for S0 reduction and H2 S production is increased.

Polysulfides and S2 O3 2− When provided with polysulfide instead of S8 , H2 S production started with less delay (Fig. 3 and was faster (Fig. 9). That can be explained if polysulfides are intermediates in the pathway from S8 to H2 S, which is credible because polysulfides are substrates for H2 S production also in certain Bacteria and Archaea (Schauder and Müller, 1993; Hedderich et al., 1999). Olson et al. (2013) reported S2 O3 2− to be a good substrate for H2 S production in homogenates of mammalian tissues. That was not observed in T. thermophila. Olson’s homogenates included 10 mM dithiothreitol, which can produce H2 S non-enzymatically from most forms of S0 , and the reaction might be augmented by rhodanese. In contrast, I used NADH, which is a natural electron donor that does not react

Table 4. Standard redox potentials at pH 7. Redox pair

E’0 (volts)

NAD+ /NADH CoQ/CoQH2 FAD/FADH2 in SQR enzyme S0 /HS−

−0.32 +0.11 −0.12 −0.27

Sources: Brock et al. (1984); Mishanina et al. (2015).

spontaneously with S8 . But another explanation may be that Olson and I studied different organisms.

Can reversal of the mitochondrial SQR reaction explain H2 S production? Standard redox potentials suggest that S0 reduction by NADH is possible (Table 4). The hypothetical pathway from NADH to S0 would be: NADH → RC-1 → Coenzyme Q → SQR → S0 . That entails the SQR reaction running in reverse. The difficulty lies with positive standard reduction potential of CoQ; the difference from CoQ to SQR is −380 mV, which makes electron transfer from CoQ to SQR nearly impossible. The possibility of reverse electron flow was suggested by Lagoutte et al. (2010), who observed electron flow from H2 S to NAD+ , which entailed running RC-1 in reverse. The electron path would be: H2 S → SQR → CoQ → RC-1 → NAD+ . In this case electron transfer from CoQ to RC-1 looks impossible, but the explanation was that reverse electron flow through RC-1 can occur because there is an electrochemical gradient across the mitochondrial inner membrane and reverse electron transport was coupled to the inward transport of H+ .

Sulfur as an electron shuttle, and evolutionary origin of mitochondria After H2 S has been aerobically oxidized, for about 10 min a product remained available that could be reduced to H2 S again (Figs. 6 and 7) Thus, there can be a sulfur cycle, and since H2 S oxidation is mitochondrial and H2 S production is cytosolic, in effect sulfur is a shuttle carrier of electrons from cytosol to mitochondria. Presumably, the ancestors of the cytoplasm and mitochondria were both sulfur-metabolizing organisms. The ancestor of the cytoplasm produced H2 S and that of mitochondria consumed H2 S. Vestiges of both activities now have been detected in their predicted cellular compartments (see Searcy(2003)).

Acknowledgments I thank the Biology Department for letting me use laboratory space after my retirement. A reviewer for pointing out that the stimulatory effect of rotenone might be explained

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by fumarate respiration. Instruments and supplies were purchased by the author. This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

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